The dependence of electrical transport pathways in Malpighian tubules on ATP
Department of Biomedical Sciences, VRT 8014, Cornell University, Ithaca, NY 14853, USA
* Author for correspondence (e-mail: kwb1{at}cornell.edu)
Accepted 10 October 2002
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Malphigian tubule, ATP, yellow fever mosquito, Aedes aegypti, metabolism, electrogenic transport, electroconductive transport
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Since the product of oxidative phosphorylation is ATP, the present study
was undertaken to examine the relationship between intracellular ATP
concentration ([ATP]i) and electroconductive and electrogenic
transport mechanisms in principal cells of Malpighian tubules of the yellow
fever mosquito. Principal cells are the sites of Na+ and
K+ secretion from the hemolymph into the tubule lumen
(Beyenbach, 1995). The entry of
K+ into the cell from the hemolymph is electroconductive through
K+-channels in the basolateral membrane
(Fig. 1). On the apical side of
the cell, extrusion of K+ from the cytoplasm into the tubule lumen
is thought to be mediated by an nH+/K+
exchanger that, in turn, is driven by the electrochemical H+
potential generated across the membrane by the activity of the electrogenic
V-type H+-ATPase, also located in the apical membrane. To examine
the relationship between [ATP]i and electroconductive and
electrogenic transport mechanisms in principal cells of the tubule, we
measured the effects of metabolic inhibition on [ATP]i in one set
of tubules, and the effects on membrane voltage and resistance of principal
cells in another set of tubules. We found that a severe reduction in
[ATP]i not only inhibited the electrogenic V-type
H+-ATPase at the apical membrane but also blocked the
K+-conductance of the basolateral membrane at the same time.
Simultaneous changes at both apical and basolateral membranes as a function of
[ATP]i suggest that ATP integrates the transport steps at these two
cell membranes. In the absence of ATP synthesis, the drop of [ATP]i
to 10% of control levels effectively isolates the cell from its luminal and
peritubular environment, apparently preserving intracellular homeostasis that
allows electrogenic and electroconductive transport mechanisms to spring back
again when ATP can be synthesized again.
|
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ringer solution and inhibitors
The control Ringer solution consisted of the following solutes, in
mmoll-1: 150.0 NaCl, 25.0 Hepes, 3.4 KCl, 1.8 NaHCO3,
1.0 MgCl2, 1.7 CaCl2 and 5.0 glucose. The pH was
adjusted to 7.1 with 1 moll-1 NaOH. The osmolality of the Ringer
solution was 320 mosmol kg-1 H2O. High K+
solutions contained 34 mmoll-1 KCl, where K+ replaced
Na+ in equimolar quantities. BaCl2, a
K+-channel blocker, was used at a concentration of 5
mmoll-1 and, to balance osmolarity, 15 mmoll-1 mannitol
was added to the control Ringer solution. To inhibit ATP synthesis we used
dinitrophenol (DNP, Sigma) or KCN (CN, Fisher Scientific) at 0.1
mmoll-1 and 0.3 mmoll-1, respectively. Control or test
Ringer solutions continuously flowed through the Lucite perfusion bath at a
rate of 2.4 ml min-1. The actual bath volume was adjusted to
approximately 250 µl by positioning the height of the suction outflow line.
Accordingly, the half-time for changing the bath was approximately 4.3 s, on
the assumption that the bath change obeyed first-order kinetics. The bath was
considered completely exchanged after 10 half-times, i.e. 43 s.
Intracellular ATP concentrations
We used the luciferin/luciferase method to measure ATP concentrations,
using the Calbiochem kit no. 119108, (La Jolla, CA, USA). Luminescence was
measured with the Lumat B9501 luminometer (Berthold Australia Pty Ltd,
Bundoora, Australia). Relative light units (RLU) of unknown quantities of ATP
were read against those of known quantities of ATP (Calbiochem) and corrected
for the background luminescence of a blank, consisting of 100 µl Ringer
solution plus 100 µl releasing reagent (Calbiochem kit). A double-log plot
of moles of ATP versus RLU yielded linear standard curves with
regression coefficients >0.99. Exploratory ATP assays in Malpighian tubules
advised the use of ATP standards ranging between 10-9 to
10-11 moles to bracket the ATP content of five Malpighian
tubules.
The ATP content was routinely measured in the complete set of five Malpighian tubules isolated from a single adult female mosquito. After depositing the five tubules in 500 µl Ringer solution, 500 µl of ATP- releasing reagent (Calbiochem) was added and stirred with a Vortex mixer for 10 s. To a 200 µl sample of this tubule extract, 100 µl of luciferin/luciferase was added and immediately read in the luminometer. The average RLU was taken from three replicates. After regression of RLU to ATP concentration, the average [ATP]i was estimated from the average dimension of the female Aedes Malpighian tubule (length, 3.5 mm; o.d., 100 µm, i.d. 10 µm).
To observe how rapidly [ATP]i would drop after metabolic inhibition with cyanide or dinitrophenol, we added 50 µl of 3 mmoll-1 KCN or 1 mmoll-1 DNP to a set of five Malpighian tubules at time zero and measured [ATP]i between 15 s to 8 min later. To measure the recovery of ATP concentrations after treatment with CN, tubules were exposed to 0.3 mmoll-1 CN for 8 min. CN was then washed out and ATP concentrations were measured 1-18 min later.
Two-electrode voltage-clamp and electrophysiological studies
Isolated Malpighian tubules lying on the bottom of the perfusion bath were
approximately 3.5 mm long. Since the open end of the tubule lumen is
continuous with the electrical ground in the peritubular bath, concerns about
short-circuiting transepithelial and membrane voltages were minimized by
selecting principal cells approximately 0.5 mm from the blind end of the
tubule, i.e. approx. 10 length constants away from the open end of the tubule
(Aneshansley et al., 1988). We
used piezo-electric remote control drives (PCS-5000; Burleigh Instruments
Inc., Fishers, NY, USA) to impale the principal cell with both current and
voltage electrodes (Fig. 1C).
Impalements near the center of the cell yielded the most stable current and
voltage records, some lasting as long as 3 h.
Conventional current and voltage microelectrodes (Kwik-Fil, Borosilicate
Glass Capillaries, TW 100F-4; World Precision Instruments, Sarasota, FL, USA)
were prepared as follows. After washing the glass capillaries with
sulfuricchromic acid cleaning solution and rinsing with distilled
water, the capillaries were stored in a drying oven at 170°C until use.
Capillaries were pulled on a programmable horizontal puller (Model P-87;
Sutter Instruments, Novato, CA, USA) to yield resistances of 20-30 M
when filled with 3 moll-1 KCl. Each principal cell was impaled with
two electrodes, a voltage electrode for measurement of the basolateral
membrane voltage (Vbl), and a current electrode for
measurement of the input resistance, Rpc
(Fig. 1C). Ag/AgCl wires
connected the microelectrodes to the clamp circuit. The Ag/AgCl wire serving
as ground in the peritubular Ringer solution was shielded with a 4%
agar-Ringer bridge.
For most of the time, the impaled principal cell was left in the open-circuit condition, i.e. the cell was not voltage-clamped. The voltage-clamp circuit was engaged only when measurements of input resistance (Rpc) were of interest. We used the GeneClamp Model 500 Voltage and Patch Clamp Amplifier (Axon Instruments, Foster City, CA, USA), head stage HS-2A-x1LU for continuous measurements of the basolateral membrane voltage (Vbl) and the head stage HS-2A-x10MGU for current injection. Rpc was determined from steady currents in response to five progressive 10 mV hyperpolarizing voltage clamp steps of 11 ms duration, beginning at -20 mV on the hyperpolarizing side and ending at +20 mV on the depolarizing side of the basolateral membrane voltage (Vbl). Currentvoltage (I-V) plots of the data were drawn using the program Clampfit (pClamp 6; Axon Instruments, Foster City, CA, USA). The plots yielded the input conductance, gpc, as the slope of the best linear fit to the I-V plot. Rpc was determined as the inverse of gpc. Current and voltage data were displayed on an oscilloscope (Iwatsu, Japan) and a strip chart recorder (model BD64; Kipp and Zonen, Crown Graphic). In addition we acquired and stored the data using the following hard- and software: a Macintosh computer (7300/200), Multifunction I/O Board PCI-1200, Signal Conditioning and Termination Board Model SC-2071, and LabView program version 4.1 (National Instruments Manufacturer, Austin, TX, USA).
Doseresponse curves and measures of EC50
The effect of each concentration of DNP or CN on the basolateral membrane
voltage (Vbl) and cell input resistance
(Rpc) was measured at least six times. The
doseresponse data were analyzed by the Hall, 4 parameter Regression of
SigmaPlot (SPSS Science, Chicago, IL, USA) to yield values and standard errors
of the EC50.
Circuit analysis
In order to explain the analysis of data obtained by the methods of
two-electrode voltage-clamp (TEVC) of a single principal cell of the tubule,
it is useful to review the model of transepithelial NaCl and KCl secretion
across the Aedes aegypti Malpighian tubule
(Fig. 1A). The model circuit
was originally derived from studies of isolated perfused Malpighian tubules
(Beyenbach, 1995,
2001
;
Beyenbach and Petzel, 1987
;
Pannabecker et al., 1993
;
Yu and Beyenbach, 2001
). Under
control conditions the isolated Aedes Malpighian tubule secretes an
approximately equimolar solution of NaCl and KCl
(Fig. 1A). The cations
Na+ and K+ take a transcellular pathway that passes
through principal cells. The `accompanying' anion, Cl-, takes a
shunt pathway located outside principal cells, apparently the paracellular
pathway. The active transport pathway for Na+ and K+
through principal cells consists of electromotive forces (E) and
resistances (R) at the basolateral (bl) and apical (a) membranes,
respectively (Fig. 1B). The
passive transport pathway for Cl- consists of the shunt resistance
(Rsh) alone. Active and passive transport pathways are
parallel to each other, forming an intraepithelial current loop, such that
cationic current through principal cells equals anionic current through the
shunt. It follows that Cl- is the `accompanying' counterion for
both Na+ and K+ secreted into the tubule lumen, which
was confirmed experimentally by observing that rates of transepithelial
Na+ and K+ secretion approximate the rate of
transepithelial Cl- secretion (Williams and Beyenbach,
1983
,
1984
). Transepithelial
diffusion potentials are negligible for Na+ and K+
because the shunt pathway is virtually impermeable to these two major cations
secreted into the tubule lumen (Williams
and Beyenbach, 1984
). Instead, the shunt pathway is permeable to
Cl-. However, transepithelial Cl- diffusion potentials
do not develop because the Cl- concentration in the tubule lumen
(161 mmoll-1, Williams and
Beyenbach, 1983
) is similar to that in the peritubular Ringer (159
mmoll-1). For these reasons, the shunt pathway
Rsh has no transepithelial diffusion potential
(Esh) in the equivalent electrical circuit of the tubule
(Fig. 1D).
In TEVC studies of a single principal cell
(Fig. 1C), the resistance
measured via intracellular electrodes is the input resistance
(Rpc) of the cell, i.e. the resistance of the basolateral
membrane in parallel to the resistances of the apical membrane and the shunt
as shown in Fig. 1D:
![]() | (1) |
![]() | (2) |
![]() | (3) |
![]() | (4) |
Statistical analysis
Most experiments were suitable for analysis by the Student's
t-test for paired samples where each principal cell served as its own
control. When this was not possible we consulted the t-test for
unequal sample size and unequal variance. EC50 values were
considered not significantly different if 3 S.E.M. of one EC50
included the mean of the other EC50. Data are summarized as mean
± S.E.M. (N, number of observations).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Plots of the peritubular CN concentration versus Vbl and Rpc yielded remarkably similar doseresponse curves (Fig. 2A). CN had no effect on voltage or resistance at concentrations less than 100 µmol l-1 but a nearly complete effect at 300 µmol l-1. Both doseresponse curves were steep, with an EC50 of 184.0±22.3 µmol l-1 for the decrease in voltage and 164.4±3075 µmol l-1 for the increase in resistance. The two EC50 values were not significantly different. Steep doseresponse curves usually result from cooperative interactions of reaction sites, but the evaluation of Hill coefficients is inappropriate here in view of the multiple steps in the cascade between the inhibition of ATP synthesis and its effect on voltage and resistance.
|
Doseresponse curves of the effects of dinitrophenol on
basolateral membrane voltage and cell input resistance
Plots of the peritubular DNP concentration versus Vbl
and Rpc were also strikingly similar. DNP had no effect on
voltage and resistance at concentrations less than 10 µmol l-1,
but a nearly complete effect at 100 µmol l-1
(Fig. 2B). Again, the
doseresponse curves were steep with an EC50 of
50.3±3.6 µmol l-1 for the decrease in voltage and
71.7±7.7 µmol l-1 for the increase in resistance. Again,
the two EC50 values were not significantly different.
Effect of cyanide on intracellular ATP concentration, voltage and
resistance of principal cells
Fig. 3 illustrates the
temporal relationship between [ATP]i and the electrophysiological
variables in Malpighian tubules, first under control conditions, then in the
presence of cyanide, and finally upon washout. Since [ATP]i was
measured in many Malpighian tubules these data are summarized as mean ±
S.E.M. The electrophysiological data corresponding in time are from a single
representative principal cell of a Malpighian tubule.
|
In freshly isolated Malpighian tubules, the control [ATP]i was
0.91±0.14 mol l-1 in 16 determinations of five tubules each
(90 Malpighian tubules in total). Control values of Vbl,
Rpc and Iat(virt) in the
representative principal cell were -76 mV, 334 k and 227.5 nA,
respectively. The addition of cyanide to Malpighian tubules caused
[ATP]i to plummet to 0.26±0.07 mmol l-1 within 15
s (11 sets of five Malpighian tubules; P<0.0005).
[ATP]i measured 30 s after CN addition rose insignificantly to
0.32±0.09 mmol l-1 (11 sets of five Malpighian tubules) and
then fell to 0.081±0.03 mmol l-1 at 2 min (5 sets of five
Malpighian tubules). [ATP]i remained near this low value for the
remaining time in cyanide.
The electrophysiological effects of CN lagged behind the effects on
[ATP]i. By the time [ATP]i had already decreased by 91%
to 0.08 mmol l-1 (2 min in cyanide), Vbl had
decreased only by 6.6% to -71 mV, with only a slight increase in
Rpc from 334.1 to 439.5 k.
Iat(virt) had decreased only 29.0% to 161.6 nA. After 3
min in the presence of CN, Vbl began to accelerate its
depolarization to 0 mV, and Rpc began to rise in parallel
while Iat(virt) continued to decrease. Shortly before
wash-out of CN, when [ATP]i was 0.11±0.03 mmol
l-1 (after 8 min in cyanide), Vbl had reached a
steady state of -8 mV, Rpc had climbed to 3150.8 k
,
and Iat(virt) had decreased to values close to zero
(Fig. 3).
The effects of CN on Vbl, Rpc and
Iat(virt) clearly lagged behind the effects on
[ATP]i. Upon washout of CN, the reverse was observed as the
recovery of [ATP]i lagged behind the recovery of
electrophysiological variables. 2 min after washout of CN,
Vbl had already returned to control values
(Fig. 3). At the same time the
input resistance had dropped 80.4%, from 3150.8 k in cyanide to 616.9
k
, and Iat(virt) had returned to 57.7% of control
values. However, [ATP]i remained near the low levels measured in
the presence of CN. When Vbl, Rpc and
Iat(virt) had fully returned to control values
approximately 12 min after washout, [ATP]i was just beginning to
recover. It would take another 5 min for [ATP]i to approach
control, pre-cyanide values.
The electrical responses to CN shown in
Fig. 3 for a single
representative principal cell were observed in seven additional principal
cells where the effects of CN were investigated. The addition of 0.3 mmol
l-1 KCN to the peritubular bath significantly
(P<0.0001) depolarized Vbl from
-85.9±2.5 mV (8 principal cells) to -13.4±3.2 mV (7), increased
Rpc significantly (P<0.001) from
383.4±22.5 k (8) to 3659.9±554.9 k
(7), and
significantly (P<0.0001) decreased Iat(virt)
from 229.0±8.5 nA (8) to 4.2±0.15 nA (7). The
electrophysiological effects of CN were observed 7-12 min after addition to
the bath, with a mean time of approximately 9 min.
Effects of dinitrophenol on intracellular ATP concentrations and the
conductance of basolateral and apical membranes of principal cells
Metabolic inhibition of the tubules with DNP yielded effects similar to
those of KCN. [ATP]i significantly (P<0.02) fell from
0.91±0.27 mmol l-1 (9 principal cells) to 0.088±0.05
mmol l-1 2 min after adding DNP. Approximately 3 min later,
Vbl significantly (P<0.0001) depolarized from
-82.4±4.0 mV (7) to -10.3±1.3 mV (7), Rpc
significantly (P<0.001) rose from 381.3±69.3 k (7)
to 4055.5±495.9 k
(7), and Iat(virt)
significantly (P<0.001) decreased from 250.4±11.4 nA (7) to
2.6±0.3 nA (7). The effects of DNP on electrophysiological variables
confirm previous observations (Beyenbach
and Masia, 2002
; Masia et al.,
2000
).
The effect of metabolic inhibition on the basolateral membrane
K+-conductance of principal cells
In view of the marked increase in the cell input resistance during
metabolic inhibition, we explored the effects of CN on the
K+-conductance of the basolateral membrane of principal cells. The
K+-conductance was of interest because it is the dominant ionic
conductance of that membrane in principal cells of Aedes aegypti
(Beyenbach, 2001;
Beyenbach and Masia, 2002
;
Masia et al., 2000
).
Fig. 4 summarizes
observations made in five principal cells where the response of the
basolateral membrane to brief tenfold increases in the peritubular
K+ concentration was evaluated first in the absence and then in the
presence of 0.3 mmol l-1 KCN. Control values in this series of
experiments were Vbl -80.0±4.1 mV and
Rpc was 495.3±61.8 k. When the peritubular
[K+] was increased from 3.4 mmol l-1 to 34 mmol
l-1, Vbl significantly (P<0.05)
depolarized to -50.2±4.9 mV. In parallel, Rpc
significantly (P<0.05) decreased to 434.9±50.1 k
.
These effects were fully reversible upon restoring the control K+
concentration.
|
Upon changing the peritubular Ringer solution to include 0.3 mmol
l-1 CN, Vbl significantly
(P<0.0002) decreased to -0.4±2.8 mV while Rpc
significantly (P<0.05) increased to 1378.4±282.9 k.
In the presence of CN, the basolateral membrane voltage failed to respond to
the tenfold increase in peritubular K+ concentration
(Fig. 4). Likewise,
Rpc was not affected by the increase in peritubular
K+ concentration.
The loss of the basolateral membrane K+-conductance in the
presence of CN was confirmed with barium
(Fig. 5). Barium is known to
block K+-channels in principal cells of the Aedes
Malpighian tubule (Masia et al.,
2000). In the present study, the addition of 5 mmol l-1
Ba2+ to the peritubular bath significantly (P<0.0001)
hyperpolarized Vbl from -79.5±2.2 mV (50) to
-89.1±3.1 mV (50), and significantly (P<0.0001) increased
Rpc from 488.9±32.3 k
(50) to
769.4±40.2 k
(50). These effects were reversible upon
Ba2+ washout. When Ba2+ was added to the peritubular
bath in the presence of 0.3 mmol l-1 CN, there were no effects on
Vbl and Rpc, indicating that
K+-channels had already closed
(Fig. 5).
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
ATP synthesis was inhibited in two distinct steps of oxidative phosphorylation: cyanide was used to prevent the generation of proton gradients across the inner mitochondrial membrane, and dinitrophenol was used to collapse it. By inhibiting cytochrome oxidase complexes, cyanide inhibits the formation of proton gradients that are needed to drive synthases, the enzymes of ATP synthesis. In addition, cyanide may also inhibit glycolysis by binding to NAD+, thereby preventing its reduction to NADH. Cyanide caused [ATP]i to drop to values below 9% of control levels and to reduce transepithelial electrolyte secretion to less than 2%, as judged from the drop of the virtual active transport current Iat(virt) from 229.0 nA to 4.2 nA.
In contrast to cyanide, DNP allows the electron transport chain to continue, but proton gradients across the inner mitochondrial membrane cannot develop because of the H+-leak pathway provided by DNP. DNP is lipid soluble. Sequestered in lipid membranes including the inner mitochondrial membrane, DNP functions like a H+-carrier (H+-ionophore). The inward proton flux bypasses ATP synthases, thereby averting the formation of ATP. DNP caused [ATP]i to drop to values less than 10% of control, and it reduced transepithelial transport to 1% of control in view of the drop of the virtual active transport current Iat(virt) from 250.4 nA to 2.6 nA. Though the mechanisms of action of cyanide and DNP differ, their convergence by affecting Vbl, Rpc and Iat(virt) in similar ways points to the requirement of ATP for the maintenance of electrogenic and conductive transport pathways.
Coupling of intracellular ATP to transepithelial transport
The inhibition of ATP synthesis by famine or toxin should be devastating
for a cell. However, neither cyanide nor DNP reduced [ATP]i to
zero. Instead, [ATP]i dropped to levels 10% of control in the
presence of CN or DNP and remained at this new steady state concentration. The
nearly complete inhibition of Iat(virt), to 2% and 1% of
control in the presence of cyanide and DNP, respectively, suggests that
[ATP]i has fallen below threshold concentrations of the V-type
H+-ATPase, thereby preventing the further hydrolysis of ATP. For
this reason [ATP]i may not have fallen below 10% of control
concentrations.
Next to inhibiting the V-type H+-ATPase at the apical membrane, metabolic inhibition caused the cell input resistance Rpc to increase fivefold or more, reflecting in part the loss of K+-conductance at the basolateral membrane (Figs 3,4,5). Thus, the inhibition of ATP synthesis affected transport steps at opposite poles of the cell, suggesting a physiological control mechanism: when K+ exit across the apical membrane into the tubule lumen cannot be energized by the V-type H+-ATPase deprived of ATP, K+ entry into the cell across the basolateral membrane is blocked. The inhibition of both pump and channels effectively isolates the cell from epithelial transport loads, thereby maintaining intracellular homeostasis. When the environment for oxidative phosphorylation improves again, the cell can mount a quick recovery with little energy input. Indeed, upon washout of cyanide or DNP, which restored ATP synthesis, the recovery of Vbl, Rpc and open K+-channels was swift, needing only a minute or two (Fig. 3).
What was observed in the present study at the all-or-none extremes of oxidative phosphorylation may take place within the normal physiological range of [ATP]i, coupling available energy to rates of transepithelial transport. ATP may mediate this regulation directly and/or indirectly via changes in intracellular pH, Ca2+ concentrations and other signaling pathways.
Inhibition of the V-type H+-ATPase
Cyanide and DNP reduced [ATP]i to values of 80-90
µmoll-1, in the vicinity of the Km for ATP
of V-type H+-ATPases. The Km of V-type
H+-ATPase in the midgut of the tobacco hornworm Manduca
sexta is 250 µmoll-1 for ATP
(Schweikl et al., 1989). If
the Km is similar in the Aedes aegypti Malpighian
tubule, then the pump activity would drop to 26% of control rates when the
[ATP]i drops to 90 µmoll-1 in the presence of DNP or
CN. Since the virtual active transport current dropped to 1%, there must be
additional mechanisms for inhibiting the V-type H+-ATPase.
Inhibition of the proton pump by ADP comes first to mind. ADP is known to
inhibit V-type H+-ATPases
(Brauer and Tu, 1994
;
Simon and Burckhardt, 1990
).
As ADP concentrations go up, or ATP/ADP ratios go down, during metabolic
inhibition, the catalytic cytosolic V1 sector of the V-type
H+-ATPases dissociates from the transmembrane V0 sector
that now no longer conducts H+ ions
(Kane and Parra, 2000
). The
dissociation can be induced in yeast cells by depriving them of glucose, and
reversed by supplying the sugar again, suggesting that assembly and
disassembly of the holoenzyme is a general physiological mechanism that
regulates V-type H+-ATPases
(Kane and Parra, 2000
;
Merzendorfer et al., 1997
). A
second way that cyanide and DNP may inhibit the V-type H+-ATPase is
via redox modulation (Harvey and
Wieczorek, 1997
). V-type H+-ATPases require reducing
conditions for activity. In contrast, oxidizing conditions inhibit the proton
pump, apparently by facilitating the formation of disulfide bonds in the
catalytic V1 sector that prevent ATP from binding
(Feng and Forgac, 1994
).
Harvey and Wieczorek have suggested that mitochondria, often in close
proximity to the V-type H+-ATPase, may produce the reducing
conditions suitable for pump activity
(Harvey and Wieczorek, 1997
;
Merzendorfer et al.,
1997
).
Mechanisms that increase the input resistance of the cell
The similarity of the EC50 values for the effect of cyanide on
voltage and resistance is striking (Fig.
2A). Likewise, the EC50 values for the effect of DNP on
voltage and resistance are surprisingly alike
(Fig. 2B). Similar
EC50 values for the effects on voltage and resistance suggest a
single target, such as the V-type H+-ATPase, where the withdrawal
of ATP from the pump affects both electrogenic and conductive properties of
the proton pump. Withholding ATP from the proton pump is expected to reduce
the electromotive force of V-type H+-ATPase, thereby reducing the
apical membrane voltage as well as the basolateral membrane voltage
via coupling through the shunt
(Masia et al., 2000). In
parallel, the separation of the V1 sector from the holoenzyme
leaves a non-conducting V0 sector in the apical membrane that is
expected to increase the resistance of the apical membrane and with it, the
input resistance of the cell. Thus, concomitant effects on voltage and
resistance yielding similar EC50 values could largely stem from
effects of metabolic inhibition on the V-type H+-ATPase located at
the apical membrane. However, K+-diffusion potentials across the
basolateral membrane (Fig. 4),
and the effects of barium in the absence and presence of KCN
(Fig. 5) illustrate that
metabolic inhibition also affects the resistance of the basolateral membrane.
In particular, K+-channels in the basolateral membrane close during
metabolic inhibition, thereby contributing to the increase in cell input
resistance (Figs
3,4,5).
The mechanism that causes basolateral membrane K+-channels to shut
down is subject to speculation. ATP, Ca2+ and Mg2+
concentrations, pH, membrane voltage and phosphorylation are all known to
affect the activity of K+-channels
(Hille, 2001
). ATP-regulated
K+-channels (KATP) open when [ATP]i is low,
which is opposite to the closing we observe in the present study (Figs
4,
5). Voltage-dependent
regulation is another possibility, but most K+-channels open at
depolarization of the membrane voltage
(Hille, 2001
), whereas in the
present study depolarization of the basolateral membrane voltage was
associated with channel closure. On the other hand, depolarizing membrane
voltages are known to drive intracellular Mg2+ ions into the pore
of open K+-channels, thereby blocking them
(Lee et al., 1999
). Additional
studies are needed to characterize the type of K+-channels
inhabiting the basolateral membrane of principal cells and to learn their
mechanisms of regulation.
In addition to resistance changes at apical and basolateral membranes of
principal cells, the closing of gap junctions between principal cells may
contribute to the increase in input resistance during metabolic inhibition. In
a previous study we have estimated that on average 5-6 principal cells are
electrically coupled (Masia et al.,
2000). ATP depletion is associated with the uncoupling of gap
junctions in ischemic cardiac cells, which is apparently mediated via
elevated cytosolic Ca2+ concentration
(Dekker et al., 1996
). Further
experiments are necessary to elucidate the contribution of gap junctions to
the measurements of input resistance in single principal cells of the
Aedes Malpighian tubules.
Kinetics of metabolic and electrophysiological events
[ATP]i dropped rapidly during metabolic inhibition
(Fig. 3). It took only 15 s for
[ATP]i to fall by 73% upon addition of cyanide to the peritubular
medium. Technical limits of the method did not allow us to measure ATP at
times earlier than 15 s. Thus, metabolic rates might be even higher than those
indicated by the available data.
In view of the close proximity of the sites of ATP synthesis (mitochondria) and electrogenesis (ATP-dependent V-type H+-ATPase) in the brush border of principal cells (Fig. 6), we expected [ATP]i and electrical variables to change in parallel, without a time lag, upon the addition of cyanide. However, in the experiment, [ATP]i consistently dropped in advance of the effects on voltage, resistance and virtual active transport current (Fig. 3). For example, it took only 2 min for the average whole cell ATP concentration to drop to 10% of control values after the addition of cyanide to the peritubular medium, but nearly 6 min for the virtual active transport current to decrease by the same percentage (Fig. 3). Although some delay (33 s) can be attributed to differences in experimental protocol (see legend to Fig. 3), most of the delay, more than 5 min for Iat(virt) to drop to 10% of control, appears to have a physiological basis. As a first hypothesis, cytosolic ATP in the vicinity of the brush border can continue to fuel the V-type H+-ATPase for some time when mitochondrial ATP synthesis is blocked. Intracellular diffusion of ATP can also explain why Iat(virt) recovers well in advance of cytosolic ATP concentration upon washout of cyanide (Fig. 3). In this case, the first ATP generated by mitochondria in the brush border serves to fuel V-type H+-ATPases close by. Additional ATP can then move on and increase cytosolic ATP concentrations in the cell.
|
Entry of K+ from the hemolymph into the cell
Wherever the electrophysiological properties of basolateral membranes have
been studied in Malpighian tubules, a prominent K+-conductance has
been found, whether the tubule secretes or reabsorbs K+
(Beyenbach and Masia, 2002;
Haley and O'Donnell, 1997
;
Neufeld and Leader, 1998
;
Nicolson and Isaacson, 1990
;
Weltens et al., 1992
;
Wessing et al., 1993
).
Blocking the K+-conductance with Ba2+ or rubidium
suggests the presence of K+-channels, but their type have yet to be
identified in any Malpighian tubule (Haley
and O'Donnell, 1997
; Hyde et
al., 2001
; Leyssens et al.,
1994
; Wessing et al.,
1993
). The addition of Ba2+ to the peritubular medium
of the tubule leads to the hyperpolarization of basolateral membrane voltage,
together with an increase in membrane resistance, consistent with the
hypothesis that pump current returning to the cytoplasmic face of the V-type
H+-ATPase is carried by K+ across the basolateral
membrane (Beyenbach, 2001
;
Masia et al., 2000
).
Supporting the model of the electrophoretic entry of K+ from the
hemolymph into the epithelial cell is the distribution of intracellular
K+ at or near electrochemical equilibrium with extracellular
K+ in Malpighian tubules of ants and the alpine weta
(Neufeld and Leader, 1998
).
However, K+-channels are not the exclusive entry pathway for
K+. Other transport mechanisms mediating K+ entry may
include the Na+/K+ ATPase
(Anstee et al., 1986
;
Grieco and Lopes, 1997
;
Linton and O'Donnell, 1999
;
Xu and Marshall, 1999
) and
possibly KCl and Na/K/2Cl cotransport systems
(Leyssens et al., 1994
).
Aw and Jones (1985) have
hypothesized that, similar to cytosolic ATP gradients, pH gradients may exist
in the cytosol, especially in cells with active proton pumps such as the
V-type H+-ATPase. Indeed, the inhibition of the V-type
H+-ATPase with bafilomycin led to intracellular acidification in
Malpighian tubules of Drosophila
(Wessing et al., 1993
), and
metabolic inhibition of Malpighian tubules in the ant also acidified the
cytoplasm (Leyssens et al.,
1993
). Thus, K+-channels in basolateral membranes of
Malpighian tubules may be pH-sensitive, closing at intracellular
acidification. Consistent with this hypothesis is the finding that epithelial
K+-channels reduce their activity at intracellular acidification
(Harvey, 1995
;
McNicholas et al., 1998
;
Onken et al., 1990
).
Midgut goblet cells in Manduca sexta and principal cells in Aedes
aegypti Malpighian tubules
The observations of the effects of metabolic inhibition we made in
principal cells of the Aedes aegypti Malpighian tubule are remarkably
similar to those made in epithelial goblet cells of the lepidopteran midgut
(Zeiske et al., 2002). To
begin, both the Aedes principal cell and the Manduca goblet
cell secrete K+ from hemolymph to lumen (or goblet cavity)
via active, electrogenic transport processes. The entry of
K+ from the hemolymph into both cells is mediated via
Ba2+-blockable K+-channels, driven by current returning
to the cytoplasmic face of the V-type H+-ATPase
(Zeiske et al., 1986
). The
extrusion of K+ across the apical membrane of the goblet cell is
thought to be mediated by an electrogenic K+/2H+
transporter that in turn is driven by the H+ electrochemical
potential set up by the V-type H+-ATPase across the apical membrane
(Azuma et al., 1995
). In
essence, the model of transepithelial K+ secretion in goblet cells
does not differ from the present, widely accepted model of K+
secretion across Malpighian tubules.
Metabolic inhibition of goblet cells with azide led to the immediate
inhibition of the short-circuit current, documenting the inhibition of
transepithelial K+ secretion when the V-type H+-ATPase
at the apical membrane is deprived of ATP. In parallel, the K+
permeability of the basolateral membrane decreased
(Zeiske et al., 2002). Thus,
metabolic inhibition inhibits epithelial transport steps at both apical and
basolateral membranes, not only in principal cells of Aedes
Malpighian tubules but also in goblet cells of Manduca midgut. These
striking similarities support the hypothesis expressed above that ATP
integrates transport steps at basolateral and apical membranes of epithelial
cells. However, the study of Zeiske et al., went on to show that intracellular
acidification duplicated the effects of metabolic inhibition on (1) apical
membrane `K+ pump' currents, and (2) on basolateral membrane
K+ permeability in the Manduca midgut
(Zeiske et al., 2002
). The
effect of intracellular acidification on [ATP]i was not measured in
goblet cells. Hence it remains to be seen whether intracellular pH,
independently of ATP, can integrate transport steps at the apical and
basolateral cell membranes.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Aneshansley, D. J., Marler, C. E. and Beyenbach, K. W. (1988). Transepithelial voltage measurements in isolated Malpighian tubules of Aedes aegypti. J. Insect Physiol. 35,41 -52.
Anstee, J. H., Baldrick, P. and Bowler, K. (1986). Studies on ouabainbinding to (Na++K+)-ATPase from Malpighian tubules of the locust, Locusta migratoria L. Biochim. Biophys. Acta 860, 15-24.
Aw, T. Y. and Jones, D. P. (1985). ATP concentration gradients in cytosol of liver cells during hypoxia. Am. J. Physiol. 249,C385 -392.[Abstract]
Azuma, M., Harvey, W. R. and Wieczorek, H. (1995). Stoichiometry of K+/H+ antiport helps to explain extracellular pH 11 in a model epithelium. FEBS Lett. 361,153 -156.[CrossRef][Medline]
Beyenbach, K. W. (1995). Mechanism and regulation of electrolyte transport in Malpighian tubules. J. Insect Physiol. 41,197 -207.[CrossRef]
Beyenbach, K. W. (2001). Energizing epithelial
transport with the vacuolar H+-ATPase. News Physiol.
Sci. 16,145
-151.
Beyenbach, K. W. and Masia, R. (2002). Membrane conductances of principal cells in Malpighian tubules of Aedes aegypti.J. Insect Physiol. 48,375 -386.[CrossRef][Medline]
Beyenbach, K. W. and Petzel, D. H. (1987).
Diuresis in mosquitoes: Role of a natriuretic factor. News Physiol.
Sci. 2,171
-175.
Brauer, D. and Tu, S. I. (1994). Effects of ATP analogs on the proton pumping by the vacuolar H+-ATPase from maize roots. Physiol. Plant. 91,442 -448.[CrossRef]
Dekker, L. R., Fiolet, J. W., VanBavel, E., Coronel, R., Opthof,
T., Spaan, J. A. and Janse, M. J. (1996). Intracellular
Ca2+, intracellular electrical coupling, and mechanical activity in
ischemic rabbit papillary muscle. Circ. Res.
79,237
-246.
Feng, Y. and Forgac, M. (1994). Inhibition of
vacuolar H+-ATPase by disulfide bond formation between cysteine 254
and cysteine 532 in subunit A. J. Biol. Chem.
269,13224
-13230.
Grieco, M. A. B. and Lopes, A. G. (1997). 5-hydroxytryptamine regulates the Na+/K+-ATPase activity in Malpighian tubules of Rhodnius prolixus: Evidence for involvement of G-protein and cAMP-dependent protein kinase. Arch. Insect Biochem. Physiol. 36,203 -214.[CrossRef]
Haley, C. A. and O'Donnell, M. J. (1997).
K+ reabsorption by the lower Malpighian tubule of Rhodnius
prolixus: Inhibition by Ba2+ and blockers of
H+/K+-ATPases. J. Exp. Biol.
200,139
-147.
Harvey, B. J. (1995). Cross-talk between sodium and potassium channels in tight epithelia. Kidney Int. 48,1191 -1199.[Medline]
Harvey, W. R. and Wieczorek, H. (1997). Animal
plasma membrane energization by chemiosmotic H+ V-ATPases.
J. Exp. Biol. 200,203
-216.
Hille, B. (2001). Ion Channels of Excitable Membranes. Sunderland: Sinauer Associates, Inc.
Hyde, D., Baldrick, P., Marshall, S. L. and Anstee, J. H. (2001). Rubidium reduces potassium permeability and fluid secretion in Malpighian tubules of Locusta migratoria, L. J. Insect Physiol. 47,629 -637.[CrossRef][Medline]
Kane, P. M. and Parra, K. J. (2000). Assembly and regulation of the yeast vacuolar H+-ATPase. J. Exp. Biol. 203,81 -87.[Abstract]
Lee, J. K., John, S. A. and Weiss, J. N.
(1999). Novel gating mechanism of polyamine block in the strong
inward rectifier K channel Kir2.1. J. Gen. Physiol.
113,555
-564.
Leyssens, A., Dijkstra, S., Van Kerkhove, E. and Steels, P.
(1994). Mechanisms of K+ uptake across the basal
membrane of Malpighian tubules of Formica polyctena: The effect of
ions and inhibitors. J. Exp. Biol.
195,123
-145.
Leyssens, A., Zhang, S.-L., Van Kerkhove, E. and Steels, P. (1993). Both dinitrophenol and Ba2+ reduce KCl and fluid secretion in Malpighian tubules of Formica: The role of the apical H+ and K+ concentration gradient. J. Insect Physiol. 39,1061 -1073.[CrossRef]
Linton, S. M. and O'Donnell, M. J. (1999).
Contributions of K+:Cl- cotransport and
Na+/K+-ATPase to basolateral ion transport in Malpighian
tubules of Drosophila melanogaster. J. Exp. Biol.
202,1561
-1570.
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. 279,F747
-F754.
McNicholas, C. M., MacGregor, G. G., Islas, L. D., Yang, Y., Hebert, S. C. and Giebisch, G. (1998). pH-dependent modulation of the cloned renal K+ channel, ROMK. Am. J. Physiol. 275,F972 -F981.[Medline]
Merzendorfer, H., Graf, R., Huss, M., Harvey, W. R. and
Wieczorek, H. (1997). Regulation of proton-translocating
V-ATPases. J. Exp. Biol.
200,225
-235.
Neufeld, D. S. and Leader, J. P. (1998). Electrochemical characteristics of ion secretion in Malpighian tubules of the New Zealand alpine weta (Hemideina maori). J. Insect Physiol. 44,39 -48.
Nicolson, S. and Isaacson, L. (1990). Patch clamp of the basal membrane of beetle Malpighian tubules: Direct demonstration of potassium channels. J. Insect Physiol. 36,877 -884.
Onken, H., Zeiske, W. and Harvey, B. (1990). Effect of mucosal H+ and chemical modification on transcellular K+ current in frog skin. Biochim. Biophys. Acta 1024,95 -102.[Medline]
Pannabecker, T. L., Aneshansley, D. J. and Beyenbach, K. W.
(1992). Unique electrophysiological effects of dinitrophenol in
Malpighian tubules. Am. J. Physiol.
263,R609
-R614.
Pannabecker, T. L., Hayes, T. K. and Beyenbach, K. W. (1993). Regulation of epithelial shunt conductance by the peptide leucokinin. J. Membr. Biol. 132, 63-76.[Medline]
Schweikl, H., Klein, U., Schindlbeck, M. and Wieczorek, H.
(1989). A vacuolar-type ATPase, partially purified from potassium
transporting plasma membranes of tobacco hornworm midgut. J. Biol.
Chem. 264,11136
-11142.
Simon, B. J. and Burckhardt, G. (1990). Characterization of inside-out oriented H+-ATPases in cholate-pretreated renal brush-border membrane vesicles. J. Membr. Biol. 117,141 -151.[Medline]
Weltens, R., Leyssens, A., Zhang, S. L., Lohrmann, E., Steels, P. and Van Kerkhove, E. (1992). Unmasking of the apical electrogenic proton pump in isolated Malpighian tubules (Formica polyctena) by the use of barium. Cell. Physiol. Biochem. 2,101 -116.
Wessing, A., Bertram, G. and Zierold, K. (1993). Effects of bafilomycin A1 and amiloride on the apical potassium and proton gradients in Drosophila Malpighian tubules studied by X-ray microanalysis and microelectrode measurements. J. Comp. Physiol. B 163,452 -462.[Medline]
Williams, J. C. and Beyenbach, K. W. (1983). Differential effects of secretagogues on Na and K secretion in the Malpighian tubules of Aedes aegypti (L.). J. Comp. Physiol. 149,511 -517.
Williams, J. C. and Beyenbach, K. W. (1984). Differential effects of secretagogues on the electrophysiology of the Malpighian tubules of the yellow fever mosquito. J. Comp. Physiol. B 154,301 -309.
Xu, W. and Marshall, A. T. (1999). X-ray microanalysis of the Malpighian tubules of the black field cricket Teleogryllus oceanicus: The roles of Na K ATPase and the Na K 2Cl cotransporter. J. Insect Physiol. 45,885 -893.[CrossRef][Medline]
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]
Zeiske, W., Meyer, H. and Wieczorek, H. (2002).
Insect midgut K+ secretion: Concerted run-down of
apical/basolateral transporters with extra- /intracellular acidity.
J. Exp. Biol. 205,463
-474.
Zeiske, W., Van Driessche, W. and Ziegler, R. (1986). Current-noise analysis of the basolateral route of K+ ions across a K+-secreting insect midgut epithelium (Manduca sexta). Pfluegers Arch. 407,657 -663.[Medline]