Effects of leucokinin-VIII on Aedes Malpighian tubule segments lacking stellate cells
Department of Biomedical Sciences, VRT 8004, Cornell University, Ithaca, NY 14853, USA
* Author for correspondence (e-mail: kwb1{at}cornell.edu)
Accepted 27 October 2003
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: leucokinin, paracellular Cl- conductance, tight junction, septate junction, Malpighian tubule, yellow fever mosquito, Aedes aegypti
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
There is good agreement that the transepithelial Cl- conductance
activated by leucokinin does not pass through principal cells of Malpighian
tubules (O'Donnell et al.,
1996; Pannabecker et al.,
1993
). Stellate cells are thought to offer the transepithelial
Cl- conductance in Drosophila melanogaster Malpighian
tubules, responding to drosokinin, the leucokinin-like diuretic in this
species (O'Donnell et al.,
1998
). Currents measured near stellate cells, not principal cells,
were found to be sensitive to Cl- channel blockers, consistent with
the activation of a transcellular Cl- conductance through stellate
cells (O'Donnell et al.,
1998
). Moreover, the drosokinin receptor and Ca2+
signaling pathway have been localized in stellate cells
(Pollock et al., 2003
;
Radford et al., 2002
). In
contrast, studies of Aedes aegypti Malpighian tubules in our
laboratory have found that leucokinin activates a Ca2+ signaling
pathway in principal cells (Yu and
Beyenbach, 2002
), which in turn increases the transepithelial
Cl- conductance of the paracellular pathway
(Pannabecker et al., 1993
).
Thus, the site of the transepithelial Cl- conductance activated by
kinins seems to be species-specific: the paracellular pathway in
Aedes Malpighian tubules and a transcellular pathway through stellate
cells in Drosophila Malpighian tubules.
In Malpighian tubules of four mosquito species, including Aedes
aegypti, stellate cells comprise 14-21% of the cell population
(Satmary and Bradley, 1984).
Thus, 1 in 5 epithelial cells along the length of the tubule is a stellate
cell. The paucity of stellate cells makes it possible to select and dissect
for study tubule segments that do not contain stellate cells. Leucokinin-VIII
activated a transepithelial Cl- conductance in these tubule
segments free of stellate cells, indicating that stellate cells are neither
needed for signaling nor for mediating the effects of leucokinin on
transepithelial Cl- conductance. These observations strengthen the
case for a diuretic mechanism mediated by principal cells and executed
via alterations in the paracellular pathway through septate
junctions.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
|
Ringer solution contained, in mmol l-1: 150.0 NaCl, 7.5 NaOH,
3.4 KCl, 1.8 NaHCO3, 1.7 CaCl2, 1.0 MgSO4,
5.0 glucose and 25.0
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid
(Hepes). The pH was adjusted to 7.1 with 1 mol l-1 NaOH. Synthetic
leucokinin-VIII was a gift from Ron Nachman (USDA, College Station, TX, USA),
and was used at a concentration of 1 µmol l-1, which is required
to exert maximal effects on tubule electrophysiology and fluid secretion
(Hayes et al., 1989;
Veenstra et al., 1997
).
Chemicals were purchased from Sigma-Aldrich (St Louis, MO, USA) and Fisher
Scientific (Suwanee, GA, USA).
In vitro microperfusion of Malpighian tubules
Fig. 1A illustrates the
method for measuring transepithelial voltage and resistance in isolated
perfused Malpighian tubules (Burg et al.,
1966; Helman,
1972
). The tubule lumen was cannulated with a double-barreled
perfusion pipette of outer diameter approximately 10 µm (Theta-Borosilicate
glass, #1402401; Hilgenberg, D-34323 Malsfeld, Germany). One barrel of this
pipette was used to perfuse the tubule lumen with Ringer solution and to
measure transepithelial voltage (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 measurements of the
transepithelial resistance (Rt) by cable analysis
(Helman, 1972
). The
peritubular bath (500 µl) was perfused with Ringer solution at a rate of 6
ml min-1. Vt was recorded continuously, and
Rt was measured periodically when of interest. The
electric isolation of the tubule lumen was ensured with pipette dimensions
that fitted the outer and inner diameters of the tubule. In addition, a
viscous resin of high dielectric constant, Sylgard® 184 (Dow Corning Inc.,
Auburn, MI, USA), was used to secure electrical insulation in the collecting
pipette (Fig. 1A,B). All
voltage measurements were done with custom-made high impedance amplifiers
(Burr-Brown, 1011
). A permanent recording of voltage during
the experiment was produced using a strip chart recorder (Model BD 64, Kipp
and Zonen, Bohemia, NY, USA).
Fig. 1A illustrates a tubule segment that includes a stellate cell, and Fig. 1B the perfusion of a tubule segment devoid of stellate cells. Fig. 1C shows an equivalent electrical circuit of transepithelial electrolyte secretion in Aedes Malpighian tubules. The transcellular active transport pathways for Na+ and K+ through principal cells at basolateral and apical membranes are lumped together in a single resistor Rc, the transcellular resistance. Active transcellular transport is driven by an electromotive force (Ec) generated at the apical membrane by the V-type H+-ATPase. Parallel to active transcellular cation transport is passive transport of Cl- through the epithelial shunt (Rsh) located outside principal cells. The active and passive transport pathways form an intraepithelial electric circuit (Fig. 1C). Since the intraepithelial loop current (I) is the same in active and passive transport pathways, it follows that the rate of transcellular cation (Na+ and K+) secretion equals the rate of Cl- secretion through the shunt, thereby conserving the electroneutrality of the fluid on both sides of the epithelium.
Transepithelial Cl- diffusion potential
The amplitude of transepithelial Cl- diffusion potentials was
measured as the change in transepithelial voltage upon a tenfold isosmotic
replacement of peritubular Cl- with isethionate. The measurement
assumes a low permeability of the tubule to isethionate
(Yu and Beyenbach, 2001). The
Cl- concentration in the tubule lumen was maintained at 156.8 mmol
l-1 by perfusing the tubule lumen with normal Ringer at rates less
than 5 nl min-1. The tenfold reduction in the peritubular
Cl- concentration drove the diffusion of Cl- from the
tubule lumen to the peritubular bath, generating lumen-positive
transepithelial diffusion potentials with magnitudes proportional to
transepithelial Cl- conductance.
Light and electron microscopy
For light microscopy, Aedes Malpighian tubules were transferred to
80 µl Ringer solution on a poly-L-lysine coated microscope slide
and covered with a 22 mm x 30 mm coverslip. The edges of the coverslip
were sealed with Permount® to prevent evaporation of the Ringer solution.
Digital images were taken with an upright microscope (Leica DMLB, Wetslar,
Germany) equipped with a digital camera (MagnaFire S99802, Optronics, Goleta,
CA, USA), hardware (IEEE-1394 PCI host controller) and software (MagnaFire
2.1A). For electron microscopy, the Malpighian tubules were prepared as
described previously (O'Connor and
Beyenbach, 2001). Sections were cut to a thickness of 70 nm.
Electron micrographs of the tubules were produced with a Philips Tecnai 12
Biotwin transmission electron microscope (FEI, Eindhoven, Netherlands).
Statistical evaluation of data
Each tubule served as its own control. Accordingly, the data were analyzed
for the differences between paired samples, control versus
experimental (leucokinin-VIII), using the paired Student's
t-test.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Principal cells are large and spindle-shaped, 80-120 µm long
(Fig. 2B;
Masia et al., 2000). Their
thickness is greatest (
30 µm) near the cell center, where the large
nucleus is located. As illustrated in Fig.
2C, cell thickness decreases towards the lateral edges of
principal cells where they contact other principal or stellate cells at
septate junctions. The lateral interstitial space between epithelial cells is
much diminished in view of the fusiform shape of principal cells. Accordingly,
transepithelial diffusion potentials reflecting paracellular permeability
reflect primarily the permeability of septate junctions.
In contrast to the large, fusiform, and opaque principal cells, stellate
cells are small, thin and transparent (Fig.
2B,C). In addition, stellate cells have the characteristic shape
of a star with rounded points (Fig.
2B; Wessing et al.,
1992). Stellate cells are 50-150 µm long and less than 5 µm
thick (Fig. 2B,C). Their
thinness and lack of intracellular concretions make them transparent, yielding
a view of the tubule lumen (Fig.
2B). Due to their small size and number, stellate cells appear
sporadically along the length of the tubule. Therefore, tubule segments
200-250 µm long frequently lack stellate cells
(Fig. 2A,B).
Fig. 2C illustrates
principal and stellate cells and their septate junctions. Principal cells have
a prominent brush border where each long microvillus is home to a
mitochondrion. The brush border of stellate cells is short and devoid of
mitochondria. In addition, stellate cells have many deep infoldings of the
basal membrane facing the hemolymph
(Bradley et al., 1982).
The effects of leucokinin-VIII on Malpighian tubule segments with and without stellate cells
Fig. 3 summarizes the
effects of leucokinin-VIII in segments of Aedes Malpighian tubules
with (A-D) and without (E-H) stellate cells. A stellate cell is clearly seen
in Fig. 3A. Incontrast,
Fig. 3E illustrates a tubule
segment consisting only of principal cells and septate junctions. Principal
cells thinning out towards their lateral edges also make them transparent near
the septate junctions. However, the presence of intracellular concretions in
these lateral transparent zones clearly identifies principal cells rather than
transparent extensions of stellate cells.
Eleven Malpighian tubule segments containing both principal and stellate
cells were studied in the absence (control) and presence of leucokinin-VIII
(Fig. 3A-D). The perfused
tubule segments were 290.9±16.3 µm long. In the absence of
leucokinin-VIII and under symmetrical perfusion with Ringer solution in the
tubule lumen and peritubular bath, the transepithelial voltage was
39.3±7.3 mV and the transepithelial resistance was 12.4±2.6
kcm (Fig. 3B,C). The
tubule segments displayed small transepithelial Cl- diffusion
potentials, 8.2±1.2 mV, in response to a tenfold replacement of
peritubular Cl- with isethionate
(Fig. 3D). Upon the addition of
1 µmol l-1 leucokinin-VIII to the peritubular bath, the
transepithelial voltage dropped significantly from 39.3 mVto 2.3±0.7
mV, and the transepithelial resistance dropped significantly from 12.4
k
cm to 2.4±0.3 k
cm
(Fig. 3B,C), turning these
moderately 'tight' epithelia into 'leaky' epithelia. In parallel with the
effects on transepithelial voltage and resistance, leucokinin-VIII
significantly increased the transepithelial Cl- diffusion potential
from 8.2 mV to 42.1±5.4 mV (Fig.
3D), indicating the activation of a transepithelial Cl-
conductance.
The above experiments were repeated in six Malpighian tubule segments
containing only principal cells and their septate junctions
(Fig. 3E-H). The perfused
tubule segments were 146.7±15.8 µm long. The leucokinin response of
these tubule segments without stellate cells was identical to that of tubule
segments composed of principal and stellate cells. Again, leucokinin-VIII
significantly decreased the transepithelial voltage from 37.8±7.0 mV to
3.4±0.6 mV, and it reduced the transepithelial resistance from
8.8±2.1 kcm to 1.7±0.2 k
cm
(Fig. 3F,G). At the same time,
leucokinin significantly increased transepithelial Cl- diffusion
potentials from 5.8±2.6 mV to 50.0±2.1 mV
(Fig. 3H). Clearly, the absence
of stellate cells did not impair, or in any way diminish, the effects of
leucokinin-VIII on epithelial electrophysiology.
One noteworthy feature of the leucokinin effects is the speed of switching
the tubule between the 'tight' and the 'leaky' states
(Fig. 4). Within seconds of
adding leucokinin-VIII to the peritubular bath, transepithelial voltage
dropped from 58.5 mV to 5.1 mV and transepithelial resistance dropped from 7.9
to 1.5 kcm, producing the 'leaky' epithelium state. The epithelium
remained 'leaky' with low values of transepithelial voltage (4.6 mV) and
resistance (1.5 k
cm) as long as leucokinin-VIII was present. However,
once leucokinin-VIII was removed from the peritubular bath, the
transepithelial voltage and resistance quickly returned towards control levels
(50.1 mV and 7.1 k
cm), switching the epithelium back to the 'tight'
state, albeit somewhat slower than the rapid on-effect of leucokinin.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Site of the leucokinin signaling pathway
Where their mechanism of action has been studied in Malpighian tubules, the
leucokinins consistently increase transepithelial NaCl and KCl secretion,
suggesting an effect on the secretion of Cl-, the counterion common
to transepithelial secretion of Na+ and K+
(Coast, 2001;
Pannabecker et al., 1993
).
Either principal or stellate cells, or both, may respond to leucokinin. In
Drosophila Malpighian tubules, stellate cells mediate the effects of
drosokinin, the Drosophila leucokinin
(Radford et al., 2002
;
Terhzaz et al., 1999
). In the
house cricket Acheta domesticus, principal cells must by default
mediate the effects of achetakinins (the Acheta leucokinin) because
Acheta Malpighian tubules do not possess stellate cells
(Coast et al., 1990
;
Hazelton et al., 1988
).
However, Malpighian tubules of the blood-sucking bug Rhodnius
prolixus, which also lack stellate cells, do not respond to leucokinin
(Bradley, 1983
;
Te Brugge et al., 2002
). Thus,
responsiveness to leucokinin is not a universal property of Malpighian
tubules, and in those Malpighian tubules that do respond to leucokinin, the
presence of stellate cells is not necessary.
The large size of principal cells in Malpighian tubules of Aedes
aegypti has allowed us to study signaling and transport processes in
these cells. In contrast, the small size of stellate cells has precluded their
study. For this reason we must infer the functions of stellate cells
indirectly, by comparing the effects of leucokinin in the presence or absence
of stellate cells. Since leucokinin elicited similar qualitative and
quantitative effects in tubules regardless of the presence of stellate cells
(Fig. 3), it is clear that
stellate cells are not required to express the effects of leucokinin in
Malpighian tubules of Aedes aegypti. Indeed, in a previous study that
probed principal cells with intracellular microelectrodes we found that
leucokinin activates a Ca2+-signaling pathway
(Yu and Beyenbach, 2002). In
particular, leucokinin activated Ca2+ channels in the basolateral
membrane of principal cells, allowing the entry of Ca2+ into the
cell from the peritubular medium as one critical step in the signaling
pathway.
Site of the transepithelial Cl- conductance activated by leucokinin
Like the Ca-signaling pathway of leucokinin, the site of the
transepithelial Cl- conductance activated by leucokinin is
species-specific. Both transcellular and paracellular pathways have been
proposed. The evidence for a transcellular Cl- pathway is strongest
in Malpighian tubules of the fruit fly, whereas the evidence for a
paracellular Cl- conductance is strongest in Malpighian tubules of
the yellow fever mosquito.
In Drosophila Malpighian tubules, stellate cells are thought to
provide a transcellular route for Cl- secretion in the presence of
leucokinin for the following reasons: (1) the location of the drosokinin
receptor in stellate cells and (2) the response of stellate cells to
drosokinin with elevated intracellular Ca2+ concentration
(Radford et al., 2002;
Terhzaz et al., 1999
). Other
observations by O'Donnell et al.
(1998
) are consistent with a
Cl- transport pathway through stellate cells but do not prove it:
(1) the identification of maxi-Cl- channels in unspecified apical
membrane domains of the tubule, (2) the paucity of maxi-Cl-
channels in only 5% of apical membrane patches and (3) the measurement of high
currents sensitive to low extracellular Cl- concentrations and
Cl- channels blockers in the vicinity of stellate cells. Together,
these observations support the conclusion that leucokinin activates apical
maxi-Cl- channels of stellate cells via intracellular
Ca2+, thereby increasing transepithelial Cl- secretion
by Drosophila Malpighian tubules.
Studies of Aedes Malpighian tubules in our laboratory have
revealed a dense population of low-conductance Cl- channels in the
apical membrane of stellate cells, which could mediate transepithelial
Cl- secretion in Malpighian tubules under control conditions
(O'Connor and Beyenbach,
2001). However, in the presence of leucokinin a paracellular
Cl- conductance is activated, which overpowers any contribution
that stellate cells might make to transepithelial Cl- secretion.
Our evidence supporting a paracellular Cl- conductance activated by
leucokinin is as follows: (1) leucokinin doubles the rate of transepithelial
fluid secretion via a non-selective increase in NaCl and KCl
secretion, suggesting the stimulation of the transport pathway for
Cl-, the counterion of Na+ and K+
(Pannabecker et al., 1993
),
(2) leucokinin drops both transepithelial resistance and voltage to values
close to zero, turning the tubule into 'leaky' epithelium with high
paracellular Cl- conductance that allows transepithelial
Cl- diffusion potentials to reach 80% of the Cl- Nernst
potential (Pannabecker et al.,
1993
; Yu and Beyenbach,
2001
), (3) the large transepithelial Cl- diffusion
potentials induced by leucokinin are similar for lumen-to-bath and
bath-to-lumen transepithelial Cl- gradients, pointing to diffusion
potentials across a single barrier such as that of septate junctions
(Pannabecker et al., 1993
),
(4) leucokinin depolarizes the apical membrane voltage and hyperpolarizes the
basolateral membrane voltage, consistent with an increased paracellular
conductance (Pannabecker et al.,
1993
; Yu and Beyenbach,
2002
), and (5) leucokinin decreases the transepithelial resistance
4.3-fold but the input resistance of the principal cells only 1.7-fold
(Masia et al., 2000
;
Yu and Beyenbach, 2001
),
pointing to a major resistance change outside principal cells. Thus,
experimental data collected in studies employing four different experimental
methods and conceptual approaches are internally consistent with leucokinin
increasing the Cl- conductance of the paracellular pathway in
Malpighian tubules of the yellow fever mosquito.
Observing the full effects of leucokinin on transepithelial voltage,
resistance and Cl- diffusion potentials in tubule segments without
stellate cells narrows the site of the activated Cl- conductance to
principal cells and/or the paracellular pathway associated with principal
cells (Fig. 3). Principal cells
can be ruled out because only an increase in paracellular Cl-
conductance can account for all the experimental voltage changes that are
induced by leucokinin (Pannabecker et al.,
1993). Furthermore, analysis of the epithelial circuit model
yields only one conclusion that is supported by the experimental data: the
increase in paracellular Cl- conductance in the presence of
leucokinin.
The strongest evidence that leucokinin activates a paracellular rather than
a transcellular Cl- conductance in Aedes tubules is
obtained by examining the effect of leucokinin on transepithelial resistance
(Pannabecker et al., 1993).
Leucokinin lowers the shunt resistance from 52.5
cm2 to 5.8
cm2. The unilateral reduction of the Cl-
concentration to 5 mmol l-1 in the peritubular bath or tubule lumen
increases the transepithelial resistance to only 16.9 and 20.1
cm2, respectively. Apparently, the reduction of the
Cl- concentration on just one side of the epithelium leaves
sufficient Cl- in the Cl- conductive pathway to elicit
only a partial increase in transepithelial resistance. However, lowering
Cl- concentration on both sides of the epithelium to 5 mmol
l-1 increases the transepithelial resistance to 55.8
cm2, fully reversing the effect of leucokinin on
transepithelial resistance. Thus, the epithelial conductance activated by
leucokinin is an extracellular conductance, as would be expected from a
septate or tight junction.
Dynamic regulation of the paracellular pathway
Like tight junctions in vertebrate tissues, septate junctions in
invertebrate tissues provide contacts between epithelial cells that not only
prevent the lateral mixing of apical and basolateral membrane domains but also
define the permselectivity and magnitude of the paracellular transport pathway
(Matter and Balda, 2003). For
a long time, tight junctions were thought to be rather fixed structures locked
into strict functional limits of differentiated epithelia. However, the
identification of three types of tight junction proteins, occludin, claudins
and junctional adhesion molecule, as well as their associated adaptor
proteins, has begun to unveil the dynamic regulation of tight junctions with
various speeds. Changes in tight junction properties involving gene expression
or molecular remodeling of tight junction proteins occur in days or hours
(Van Itallie et al., 2001
).
Dynamic formation and reorganization of tight junctions between
claudin-transfected fibroblasts can occur within minutes
(Sasaki et al., 2003
). In
Malpighian tubules of the yellow fever mosquito, we observe the regulation of
the paracellular Cl- conductance with switch-like speed that has
not been observed in other epithelia
(Beyenbach, 2003
). Malpighian
tubules, in particular, may have developed the regulation of tight junction
permeability to an extraordinary degree. In the absence of glomerular
filtration, Malpighian tubules must rely entirely on tubular mechanisms of
renal regulation. Rapid, reversible changes in paracellular Cl-
conductance may endow the tubule with powers of diuresis not unlike those of
glomerular kidneys.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Beyenbach, K. W. (1993). Extracellular fluid homeostasis in insects? In Molecular Comparative Physiology; Structure and Function of Primary Messengers in Invertebrates, vol. 12 (ed. K. W. Beyenbach), pp.146 -173.Basel: Karger.
Beyenbach, K. W. (2001). Energizing epithelial
transport with the vacuolar H+-ATPase. News Physiol.
Sci. 16,145
-151.
Beyenbach, K. W. (2003). Regulation of tight junction permeability with switch-like speed. Curr. Opin. Nephrol. Hypertens. 12,543 -550.[Medline]
Blackburn, M. B., Wagner, R. M., Shabanowitz, J., Kochansky, J. P., Hunt, D. F. and Raina, A. K. (1995). The isolation and identification of three diuretic kinins from the abdominal ventral nerve cord of adult Helicoverpa zea. J. Insect Physiol. 41,723 -730.[CrossRef]
Bradley, T. J. (1983). Functional design of microvilli in the Malpighian tubules of the insect Rhodnius prolixus.J. Cell Sci. 60,117 -135.[Abstract]
Bradley, T. J., Stuart, A. M. and Satir, P. (1982). The ultrastructure of the larval Malpighian tubules of a saline water mosquito Aedes taeniorhynchus. Tissue Cell 14,759 -774.[CrossRef][Medline]
Burg, M., Grantham, J., Abramow, M. and Orloff, J.
(1966). Preparation and study of fragments of single rabbit
nephrons. Am. J. Physiol.
210,1293
-1298.
Coast, G. M. (2001). The neuroendocrine regulation of salt and water balance in insects. Zool. (Jena) 103,179 -188.
Coast, G. M., Holman, G. M. and Nachman, R. J. (1990). 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.
Coast, G. M., Kay, I. and Wheeler, C. H. (1993). 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, vol. 12 (ed. K. W. Beyenbach), pp. 38-66. Basel: Karger.
Coast, G. M., Orchard, I., Phillips, J. E. and Schooley, D. A. (2002). Insect diuretic and antidiuretic hormones. Adv. Insect Physiol. 29,279 -409.
Hayes, T. K., Pannabecker, T. L., Hinckley, D. J., Holman, G. M., Nachman, R. J., Petzel, D. H. and Beyenbach, K. W. (1989). Leucokinins, a new family of ion transport stimulators and inhibitors in insect Malpighian tubules. Life Sci. 44,1259 -1266.[CrossRef][Medline]
Hazelton, S. R., Parker, S. W. and Spring, J. H. (1988). Excretion on the house cricket (Acheta domesticus): fine structure of the Malpighian tubules. Tissue Cell 20,443 -460.[CrossRef]
Helman, S. I. (1972). Determination of electrical resistance of the isolated cortical collecting tubule and its possible anatomical location. Yale J. Biol. Med. 45,339 -345.[Medline]
Holman, G. M., Cook, B. J. and Nachman, R. J. (1987). Isolation, primary structure and synthesis of leucokinin-VII nad VIII: the final members of the new family of cephalomyotropic peptides isolated from head extracts of Leucophaea maderae. Comp. Biochem. Physiol. 88C, 31-34.[CrossRef]
Iaboni, A., Holman, G. M., Nachman, R. J., Orchard, I. and Coast, G. M. (1998). Immunocytochemical localisation and biological activity of diuretic peptides in the housefly, Musca domestica.Cell Tissue Res. 294,549 -560.[CrossRef][Medline]
Leyssens, A., Steels, P., Lohrmann, E., Weltens, R. and Van Kerkhove, E. (1992). Intrinsic regulation of potassium transport in Malpighian tubules Formica electrophysiological evidence. J. Insect Physiol. 38,431 -446.[CrossRef]
Masia, R., Aneshansley, D., Nagel, W., Nachman, R. J. and
Beyenbach, K. W. (2000). Voltage clamping single cells in
intact Malpighian tubules of mosquitoes. Am. J. Physiol. Renal
Physiol. 279,F747
-F754.
Matter, K. and Balda, M. S. (2003). Signalling to and from tight junctions. Nat. Rev. Mol. Cell. Biol. 4,225 -236.[CrossRef][Medline]
O'Connor, K. R. and Beyenbach, K. W. (2001).
Chloride channels in apical membrane patches of stellate cells of Malpighian
tubules of Aedes aegypti. J. Exp. Biol.
204,367
-378.
O'Donnell, M. J., Dow, J. A., Huesmann, G. R., Tublitz, N. J.
and Maddrell, S. H. (1996). Separate control of anion and
cation transport in Malpighian tubules of Drosophila melanogaster.J. Exp. Biol. 199,1163
-1175.
O'Donnell, M. J., Rheault, M. R., Davies, S. A., Rosay, P.,
Harvey, B. J., Maddrell, S. H., Kaiser, K. and Dow, J. A.
(1998). Hormonally controlled chloride movement across
Drosophila tubules is via ion channels in stellate cells.
Am. J. Physiol. Regul. Integr. Comp. Physiol.
274,R1039
-R1049.
Pannabecker, T. L., Hayes, T. K. and Beyenbach, K. W. (1993). Regulation of epithelial shunt conductance by the peptide leucokinin. J. Mem. Biol. 132, 63-76.[Medline]
Plawner, L., Pannabecker, T. L., Laufer, S., Baustian, M. D. and Beyenbach, K. W. (1991). Control of diuresis in the yellow fever mosquito Aedes aegypti: evidence for similar mechanisms in the male and female. J. Insect Physiol. 37,119 -128.[CrossRef]
Pollock, V. P., Radford, J. C., Pyne, S., Hasan, G., Dow, J. A.
and Davies, S. A. (2003). NorpA and itpr mutants reveal roles
for phospholipase C and inositol (1,4,5)-trisphosphate receptor in
Drosophila melanogaster renal function. J. Exp.
Biol. 206,901
-911.
Radford, J. C., Davies, S. A. and Dow, J. A. T.
(2002). Systematic GPCR analysis in Drosophila
melanogaster identifies a leucokinin receptor with novel roles.
J. Biol. Chem. 277,38810
-38817.
Rosay, P., Davies, S. A., Yu, Y., Sozen, A., Kaiser, K. and Dow,
J. A. (1997). Cell-type specific calcium signalling in a
Drosophila epithelium. J. Cell Sci.
110,1683
-1692.
Sasaki, H., Matsui, C., Furuse, K., Mimori-Kiyosue, Y., Furuse,
M. and Tsukita, S. (2003). Dynamic behavior of paired claudin
strands within apposing plasma membranes. Proc. Natl. Acad. Sci.
USA 100,3971
-3976.
Satmary, W. M. and Bradley, T. J. (1984). The distribution of cell Types in the Malpighian tubules of Aedes taeniorhynchus (Wiedemann) diptera culicidae. Int. J. Insect Morphol. Embryol. 13,209 -214.[CrossRef]
Te Brugge, V. A., Schooley, D. A. and Orchard, I. (2002). The biological activity of diuretic factors in Rhodnius prolixus. Peptides 23,671 -681.[CrossRef][Medline]
Terhzaz, S., O'Connell, F. C., Pollock, V. P., Kean, L., Davies,
S. A., Veenstra, J. A. and Dow, J. A. (1999). Isolation and
characterization of a leucokinin-like peptide of Drosophila melanogaster.J. Exp. Biol. 202,3667
-3676.
Thompson, K. S., Rayne, R. C., Gibbon, C. R., May, S. T., Patel, M., Coast, G. M. and Bacon, J. P. (1995). Cellular colocalization of diuretic peptides in locusts: a potent control mechanism. Peptides 16,95 -104.[CrossRef][Medline]
Van Itallie, C., Rahner, C. and Anderson, J. M.
(2001). Regulated expression of claudin-4 decreases paracellular
conductance through a selective decrease in sodium permeability. J.
Clin. Invest. 107,1319
-1327.
Van Kerkhove, E., Weltens, R., Roinel, N. and De Decker, N. (1989). Hemolymph composition in Formica hymenoptera and urine formation by the short isolated Malpighian tubules electrochemical gradients for ion transport. J. Insect Physiol. 35,991 -1004.[CrossRef]
Veenstra, J. A., Pattillo, J. M. and Petzel, D. H.
(1997). 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.
Wessing, A., Zierold, K. and Hevert, F. (1992). Two types of concretions in Drosophila Malpighian tubules as revealed by X-Ray microanalysis a study on urine formation. J. Insect Physiol. 38,543 -554.
Yu, M.-J. and Beyenbach, K. W. (2001). Leucokinin and the modulation of the shunt pathway in Malpighian tubules. J. Insect Physiol. 47,263 -276.[CrossRef][Medline]
Yu, M.-J. and Beyenbach, K. W. (2002).
Leucokinin activates Ca2+-dependent signal pathway in principal
cells of Aedes aegypti Malpighian tubules. Am. J. Physiol.
Renal Physiol. 283,F499
-F508.