Department of Biology and Center for Biological Timing, University of Virginia, Charlottesville, Virginia 22904-4328
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ABSTRACT |
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The Malpighian (renal) tubule of Drosophila melanogaster is a useful model for studying epithelial transport. The purpose of this study was to identify factors responsible for modulating transepithelial chloride conductance in isolated tubules. I have found that tyrosine and several of its metabolites cause an increase in chloride conductance. The most potent of these agonists is tyramine, which is active at low nanomolar concentrations; the pharmacology of this response matches that of the previously published cloned insect tyramine receptor. In addition, the tubule appears capable of synthesizing tyramine from applied tyrosine, as shown by direct measurement of tyrosine decarboxylase activity. Immunohistochemical staining of tubules with an antibody against tyramine indicates that the principal cells are the sites of tyramine production, whereas previous characterization of the regulation of chloride conductance suggests that tyramine acts on the stellate cells. This is the first demonstration of a physiological role for an insect tyramine receptor.
tyrosine decarboxylase; yohimbine; tyramine receptor; biogenic amines
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INTRODUCTION |
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THE MALPIGHIAN TUBULES comprise the initial component of the insect excretory system and have been a useful model system for studying the regulation of epithelial ion transport (4, 17, 37). The fruit fly Drosophila melanogaster possesses two pairs of tubules; these are blind-ended simple tubular epithelia that empty into the digestive tract (17). The main segment of the tubule, in which the secretion of a potassium-rich primary urine occurs, contains two morphologically defined cell types, principal cells and stellate cells (52). Recent work has shown that the principal cells are both genetically and functionally heterogeneous (43, 49), although neither the molecular nor the physiological bases for this heterogeneity have been determined.
In Drosophila, as in other insects, cations move into the tubule lumen by active transport through the principal cells (4, 16, 25, 32). The electromotive force for this transport is generated by a vacuolar-type proton pump located in the apical membrane. Chloride moves passively into the lumen down its electrochemical gradient through a pathway that lies outside of the principal cells (32, 34). The anatomical site of this chloride shunt conductance, paracellular or transcellular through the stellate cells, has been controversial; however, regulation of chloride conductance in the Drosophila tubule is linked to intracellular calcium levels in the stellate cells (33). The net effect of ion transport in the Drosophila tubule is the establishment of a lumen-positive transepithelial potential (TEP) with an amplitude of ~50-60 mV.
Ion transport across the Drosophila tubule is a highly regulated process, falling under the control of at least three different second messenger systems. Cation transport through the principal cells can be stimulated by both cGMP and cAMP (32). The former is produced after treatment of tubules with the diuretic hormone cardioacceleratory peptide 2b (CAP2b) (15); production of cAMP is stimulated by a calcitonin-related peptide (14). In many other insect species, a corticotropin-releasing factor (CRF)-like diuretic peptide also acts through cAMP (13). In addition, the diuretic hormone leucokinin stimulates urine secretion by raising calcium levels in the stellate cells and increasing the amplitude of the chloride shunt conductance (33). Recently, a receptor for leucokinin has been cloned and found to be expressed exclusively in stellate cells (41). The biogenic amines serotonin and octopamine have diuretic activity on the tubules of multiple insect species (26, 31, 48); however, no aminergic modulation of Drosophila renal function has previously been reported.
In the tubules of many insects, including Drosophila, the TEP measured in vitro is not constant but instead undergoes large oscillations in amplitude (15, 30, 39, 53). Recent studies in the mosquito Aedes aegypti and in Drosophila have shown that these voltage oscillations are caused by fluctuations in transepithelial chloride conductance (5, 7), and in Drosophila they are dependent on increased intracellular calcium levels in the stellate cells (7). No function has yet been determined for these oscillations; neither have they been found to be caused by any particular stimulus. However, the occurrence of such oscillations in the tubules of several different insect species raises the possibility that they may be important for the regulation of normal ion transport. The initial purpose of this study was the identification of the factor responsible for the triggering of these oscillations; the result, as described below, was the identification and characterization of tyramine as a novel insect diuretic factor.
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METHODS |
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Materials. All chemicals were obtained from Sigma (St. Louis, MO).
Drosophila maintenance. Drosophila melanogaster (Canton-S) were maintained on a 12:12-h light-dark cycle at 23-25°C on cornmeal-molasses-yeast medium.
Tubule isolation.
Posterior Malpighian tubules were dissected under dissecting saline
from adults females 6-8 days posteclosion and placed in a tissue
culture dish in which a 100-µl drop of 0.125 mg/ml
poly-L-lysine had been dried to promote adhesion of the
tubule to the dish, and the solution was replaced with recording medium
(32). The dissecting/recording saline contained (in mM) 85 NaCl, 20 KCl, 3 CaCl2, 12 MgSO4, 7.5 NaHCO3, 4 NaH2PO4, 15 glucose, and
10 HEPES, pH 6.75 [osmolality = 255-270
mosmol/kgH2O as measured with a vapor pressure osmometer
(Wescor, Logan, UT)]. Standard bathing medium (SBM) consisted of a 1:1
mixture of Schneider's Drosophila medium (Invitrogen,
Carlsbad, CA) and a "diluting saline" containing 36 NaCl, 21 KCl,
15 MgCl2, 5 CaCl2, 4.8 NaHCO3, 2 NaH2PO4, 11.1 glucose, and 15 HEPES, pH 6.75. The composition of SBM, based on the compositions of the diluting
saline and Schneider's medium (Invitrogen) is (in mM) 36 NaCl, 21 KCl,
5.2 CaCl2, 7.5 MgSO4, 7.5 MgCl2,
4.8 NaHCO3, 1.3 KH2PO4, 3.4 sodium
phosphate, 7.5 HEPES, 0.7 -ketoglutaric acid, 11.1 D-glucose, 0.43 fumaric acid, 0.38 malic acid, 0.42 succinic acid, 2.9 trehalose, 2.8
-alanine, 1.15 L-arginine, 1.5 L-aspartic acid, 0.25 L-cysteine, 0.21 L-cystine, 2.7 L-glutamic acid, 6.15 L-glutamine, 1.67 glycine, 1.29 L-histidine, 0.58 L-isoleucine,
0.58 L-leucine, 4.5 L-lysine HCl, 2.7 L-methionine, 0.45 L-phenylalanine, 7.4 L-proline, 1.19 L-serine, 1.47 L-threonine, 0.25 L-tryptophan, 1.38 L-tyrosine, 1.32 L-valine, and 1,000 mg/l yeastolate. The osmolality of SBM was 255-270
mosmol/kgH2O. The osmolality of these solutions is equal to
that reported for adult Drosophila hemolymph
(47) but is much lower than that used in other studies of
Drosophila tubule physiology. This difference in osmolality
explains the higher urine secretion rates and more prominent TEP
oscillations than those reported by others (8). For the
chloride replacement experiment, the low chloride media consisted of
10% normal dissecting/recording saline and 90% dissecting/recording saline in which NaCl, KCl, and CaCl2 had been replaced with
85 mM sodium isethionate, 10 mM K2SO4, and 3 mM
CaSO4. Experiments were conducted within 2 h of tubule dissection.
Recording. The tubule lumen and principal cells were impaled with a sharp electrode (R > 25 MOhms) pulled from theta-glass (Sutter Instruments, Novato, CA) and filled with 3 M KCl. Potentials were amplified (Axopatch 200B; Axon Instruments, Foster City, CA), digitized at 100 Hz, and stored online. Recording and analysis were conducted using pCLAMP software (Axon Instruments). The peritubular bath was continuously perfused during recording. Drugs were applied to and removed from the bath during recording by switching perfusion lines.
Quantitation of electrophysiological responses.
To compare the responses of tubules to various agonists, a response
index was calculated as follows. A straight line was fitted to the TEP
record for the 30 s preceding the drug application and then
extrapolated through the time of the application. The area under the
TEP trace and under the extrapolated line was measured for a period
beginning 15 s after the beginning of the drug application and
ending 15 s after the end of the drug application. The baseline for the integral was 10 mV, based on the observation that during maximal responses, the TEP drops to approximately this value. The
response index was calculated as 1
(area under the TEP
trace/area under the extrapolated line). In occasional traces,
small instabilities in the TEP led to extrapolated lines that did not
accurately reflect the behavior of the TEP; in those cases, the 30-s
fitting interval was adjusted to give a more accurate line.
RT-PCR. RT-PCR from whole fly and Malpighian tubule cDNA was performed as previously described (7). Primer sequences for the tyramine receptor (accession no. AB-073914) were ACAAGGACTCAGCGGGAGAATG and CGATGATGGTCAGCACGATAATG.
Urine secretion assay. The rate of urine secretion from isolated tubules was measured as described (16). Posterior tubules were dissected from adult females under SBM or saline (depending on the experiment) and then moved into a 15 µl of droplet of SBM or saline under mineral oil. One tubule was pulled out of the droplet and wrapped around a dissecting pin such that the cut end of the ureter and the lower section of the other tubule branch were pulled into the oil. Periodically, the secreted urine droplet was removed from the ureter with a fine glass rod, and its diameter was measured with an ocular micrometer. The volume of the droplet was calculated assuming spherical geometry. Collection intervals ranged from 12-20 min. Secretion rate was calculated as urine volume/collection time. For drug application, 12 µl of the bathing droplet was replaced twice with 12 µl of bathing solution + drug. The bathing droplet of control tubules was replaced twice with bathing solution alone.
Determination of chloride concentration in urine. The urine secretion assay was performed as described above. After the diameter of each urine droplet was measured, the droplet was taken up into a 250 nl of capillary tube (Drummond Scientific, Broomall, PA). The volume of the urine was again determined by measuring the length of the aqueous phase within the capillary. Any samples for which the two volume measurements disagreed by >10% were discarded from further analysis. The urine droplet was expelled into a tube containing 20 µl of H2O. Capillary electrophoresis was performed on the samples as described using a Waters Quanta 4000 (23). The elution time and area of the ultraviolet absorbance peaks were compared with standards of known composition and concentration, and the ion concentrations in the original urine droplet were then calculated.
Tyramine immunohistochemistry. Tubules were dissected from 7- to 9-day-old females under PBS and then incubated for 20 min in PBS containing 1 mM L-tyrosine before fixation. Immunostaining was performed as previously described (11), with the following modifications: the concentration of NaBH4 was 1%, and 0.1% BSA was added to the PMT solution. The primary antibody was a rabbit anti-tyramine antibody (Chemicon International, Temecula, CA), diluted 1:2,500. Immunostaining was detected with a biotinylated anti-rabbit secondary antibody, diluted 1:250 (Jackson ImmunoResearch, West Grove, PA), followed by the Vectastain peroxidase ABC kit (Vector Laboratories, Burlingame, CA) and incubation with DAB/peroxide/nickel.
Tyrosine decarboxylase enzyme assays.
Assays for tyrosine decarboxylase (TDC) activity were performed on
tubule extracts as described (28). Tubules from 18 adult females (4-7 days posteclosion) were homogenized on ice in 90 µl
of buffer (50 mM Tris, pH 7.5, and 1 mM phenylthiourea). Protein content was measured in an aliquot of homogenate using the Bio-Rad protein assay reagent (Bio-Rad, Hercules, CA). Tubule homogenate (12 µl) was mixed with 48 µl of assay mix [100 mM sodium phosphate, pH
6.8, 0.1 mM pyridoxal phosphate, 1 mM -mercaptoethanol, 0.1 mM EDTA,
and 20 µCi/ml L-[3,5-3H]tyrosine (Amersham,
Piscataway, NJ)] and either incubated for 60 min at 30°C or placed
on ice and processed immediately. Reactions were stopped by the
addition of 300 µl of sodium phosphate, pH 8.0, and extracted with
100 µl of chloroform containing 100 mM diethylhexylphosphoric
acid. The organic phase was reextracted with an additional 300 µl of
sodium phosphate, and then 80 µl of the organic phase was placed into
a scintillation vial and allowed to dry before the addition of
scintillant and counting. Each experiment consisted of two
zero-timepoint reactions (background) and three 60-min reactions; TDC
activity was calculated by subtracting the averaged background counts
from the averaged 60-min counts.
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RESULTS |
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Isolated Drosophila Malpighian tubules exhibit a
lumen-positive TEP that undergoes large oscillations in amplitude, but
the cause of these oscillations has not yet been determined (7, 15). During the characterization of these oscillations, it
became clear that although they were always present in tubules bathed in SBM, which contains tissue culture medium, oscillations were rarely
seen when tubules were bathed in a simple saline. This effect of SBM is
demonstrated most clearly by switching the bathing medium from saline
to SBM; a typical response is shown in Fig. 1A. As is clear from this
trace, SBM has two effects on the TEP. The first is a rapid and
reversible depolarization and induction of oscillations. The appearance
of oscillations is quantified by an increase in the coefficient of
variation of the TEP (Fig. 1B, left). The second
effect of SBM is an increase in the amplitude of the TEP that persists
after the SBM is washed out (Fig. 1B, right).
This study focuses on the depolarizing activity of SBM (however, see
Fig. 7).
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SBM contains 50% Schneider's Drosophila medium, which is a
mixture of yeast extract, salts, sugars, metabolic intermediates, and
20 amino acids (see METHODS for precise composition). To
determine which component of the medium is responsible for the
induction of oscillations, tubules were exposed to solutions containing different constituents of the complete medium. TEP oscillations were
observed in the presence of either yeast extract or a mixture of all of
the amino acids (data not shown). Individual testing of 10 of these
amino acids revealed that tubules respond only to
L-tyrosine. As shown in Fig.
2A, application of
L-tyrosine causes the TEP to depolarize and oscillate in a
manner indistinguishable to that induced by complete SBM.
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The amino acid tyrosine serves as a precursor for the synthesis of
several biogenic amines (29, 45); tyrosine metabolism in
insects is diagrammed in Fig. 3. Unlike
in mammals, where a single aromatic amino acid decarboxylase acts upon
both tyrosine and DOPA, insects possess separable TDC and DOPA
decarboxylase (DDC) enzymes (54). Because it seemed
possible that one of these metabolites could be mediating the effect of
tyrosine, I tested three biogenic amines-dopamine, octopamine, and
tyramine-on tubules. All three compounds elicited qualitatively
similar electrical responses that also resembled those caused by
tyrosine (Fig. 2B). At low doses, these amines caused the
TEP to oscillate. Intermediate doses elicited a transient
depolarization followed by oscillations, and high doses resulted in a
sustained depolarization. This pattern of responses is quite similar to
that observed in Aedes tubules to increasing doses of
leucokinin (51). Strikingly, tyramine was several orders
of magnitude more potent than any other compound tested, giving a
measurable response at a concentration of 1 nM and a maximal response
at ~100 nM (Fig. 2C). In contrast, octopamine caused
measurable responses only at concentrations above 1 µM. The
preferential response of tubules to tyramine over octopamine is
particularly interesting, given that for many years, tyramine was
thought simply to be a metabolic intermediate on the octopamine synthesis pathway. Only relatively recently have several lines of
evidence suggested that tyramine can function as an independent signaling molecule; chief among these has been the cloning from Drosophila and other insect species of a receptor that is
preferentially activated by tyramine over octopamine (2, 6, 40,
42, 46, 50).
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To determine whether the antagonist pharmacology of the tubule response
also matched that of the cloned tyramine receptor, I assayed the
ability of several tyramine/octopamine receptor antagonists to block
the depolarization elicited by an application of 50 nM tyramine. A full
dose-response profile was generated for yohimbine, which is the most
potent inhibitor of tyramine binding in the locust brain and to the
cloned receptor (20, 46, 50); yohimbine blocked the
response of tubules to 50 nM tyramine with a half-maximal concentration
of ~300 nM. The rank-order potency of the three antagonists tested
was yohimbine phentolamine > metoclopramide (Fig.
4). This profile is very similar to that of an insect tyramine receptor stably expressed in a
Drosophila cell line (50) and of the tyramine
binding site in locust brain (20) but does not resemble
that of any of the octopamine receptor subtypes (21).
Consistent with the pharmacology, RT-PCR analysis shows that the gene
encoding the tyramine receptor is expressed in the Malpighian tubules
(Fig. 5).
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I next sought to characterize the ionic basis of the tyramine-induced
depolarization. Previous work has shown that the TEP oscillations seen
in the presence of SBM are due to fluctuations in the transepithelial
chloride conductance (7). Because the Nernst potential for
chloride between the peritubular bath and the lumen is near 0 mV, an
increase in chloride conductance will cause the TEP to depolarize. It
was therefore of interest to determine whether the tyramine-induced
depolarizations were also associated with an increase in chloride
conductance. Such was likely to be the case, both because high
concentrations of tyramine cause the TEP to depolarize to near 0 mV and
also because the electrical response of the tubule to tyramine
resembles the response to leucokinin, which is known to increase
chloride conductance (32). The chloride substitution
experiment shown in Fig. 6 shows that
tyramine does indeed cause a large increase in transepithelial chloride
conductance, as indicated by the increased chloride diffusion potential
in the presence of tyramine.
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It is evident that the treatment of tubules with tyramine receptor
agonists causes an electrical response indistinguishable from that
induced by SBM. To determine whether the components of SBM act solely
through the tyramine receptor, I measured the ability of yohimbine to
block the response of the tubule to SBM. Figure
7A shows the TEP coefficient
of variation measurements from tubules recorded in SBM in the presence
of yohimbine; oscillations are blocked with a dose dependence very
similar to that shown in Fig. 4 for the antagonism of tyramine
responses. At the highest concentrations of yohimbine, TEP oscillations
are completely absent, although the coefficient of variation is nonzero
due to small fluctuations and drift in the TEP. Importantly, long-term
(1-2 h) exposure to yohimbine did not appear to be toxic to the
tubules, because there was no reduction in TEP amplitude with
increasing concentrations of yohimbine. These results demonstrate that
all of the oscillation-inducing activity of SBM is due to the
activation of yohimbine-sensitive tyramine receptors.
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Interestingly, the ability of SBM to increase the amplitude of the TEP, shown in Fig. 1, is not blocked by yohimbine. Figure 7, B and C, shows a similar experiment to that in Fig. 1, but in the presence of 100 µM yohimbine. Clearly, the blockade of the tyramine receptor prevents the depolarization and oscillations seen previously; however, the sustained increase in TEP amplitude is identical to that seen in Fig. 1. The time course of this increase is much more apparent in the absence of the depolarization and oscillations. This augmentation of the TEP could be due to a specific component of the SBM acting on a yohimbine-insensitive receptor; alternatively, exposure of the tubule to a rich medium such as SBM may result in a more general stimulation of epithelial transport. Also seen in the trace in Fig. 7B are a rapid hyperpolarization and depolarization when SBM is added and removed. These rapid effects are most likely due to differences in the ionic composition of saline and SBM (see METHODS). Most notably, SBM contains significantly less sodium than the saline, and removal of sodium from the peritubular bath leads to a hyperpolarization of the TEP (unpublished results).
Because yohimbine appears to be a selective blocker of tyraminergic
signaling in the tubule, it is possible to use this antagonist to
examine the role of tyramine in the function of the intact tubule.
Urine secretion rates were measured in isolated tubules bathed in SBM;
the addition of 100 µM yohimbine caused a 60% reduction in the rate
of urine production (Fig. 8A).
Measurements of the chloride concentration in the secreted urine
droplets showed no effect of yohimbine (Fig. 8B). Thus the
60% reduction in secretion rate indicates a reduction in
transepithelial chloride flux of the same magnitude. In isolated
tubules bathed in SBM, therefore, approximately half of the
transepithelial chloride transport occurs through a yohimbine-sensitive
pathway, whereas the other half of the chloride transport persists in
the absence of tyramine signaling.
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If tyramine acts to stimulate transepithelial chloride flux, it should trigger diuresis when added to isolated tubules. Figure 8C shows that this is indeed the case. When added to tubules that were bathed in saline, 1 µM tyramine caused a 45% increase in urine secretion (P < 0.005, paired t-test).
The data shown in Fig. 2 demonstrate that tubules respond to tyrosine
as well as tyramine, albeit at far greater concentrations. Tyrosine
could be acting in one of two ways to elicit this response; it could be
a weak agonist of the tyramine receptor, or it could be converted into
tyramine by a TDC activity endogenous to the tubule. Two lines of
evidence suggest that the latter is true. First, tubules contain a
significant level of TDC activity. Assays of tubule extracts show a TDC
activity of 18.8 ± 2.6 fmol
tyramine · min1 · mg
1
protein (n = 2 experiments). The only basis for
comparison of this value with another Drosophila tissue is a
report of TDC activity in extracts of adult brain of ~4
fmol · 10 min
1 · brain
1
(28). Converting the tubule activity yields a value for
the activity from one fly's set of tubules of 0.21 ± 0.05 fmol/10 min; thus the tubules in a fly contain ~5% of the TDC
activity of the brain. Given that the tubules consist of just over 300 cells (49), the amount of activity per cell is far higher
in the tubules than in the brain.
The second line of evidence for endogenous conversion of tyrosine to
tyramine is the observation that D-tyrosine, the inactive enantiomer of L-tyrosine, inhibits the response of
wild-type tubules to L-tyrosine but not to tyramine. Figure
9 shows traces from tubules challenged
with two applications of either L-tyrosine or tyramine;
between the two applications, the tubule is treated with either
D-tyrosine or saline. As quantified in Fig. 9C,
the second response to L-tyrosine is nearly eliminated by
the D-tyrosine application, whereas the tyramine response
is unaffected. This selective antagonism would be virtually impossible
to explain if L-tyrosine were a direct agonist of the
tyramine receptor. If, however, the tubule is converting tyrosine to
tyramine, there are many steps that could potentially be inhibited by
D-tyrosine (see DISCUSSION). Taken together
therefore, the D-tyrosine and TDC assay results provide
strong evidence for endogenous production of tyramine by the
Drosophila tubule.
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The tubule is a heterogeneous tissue, and TEP measurements do not give
any information as to cellular localization of function; therefore, it
is important to determine which cell type is responsible for the
production of tyramine. Because TDC is thought to be an intracellular
enzyme (19), I reasoned that the tyramine-producing cells
should contain relatively high levels of tyramine. Figure 10 shows the pattern of immunostaining
seen with an antibody against tyramine. The principal cells are
labeled, whereas stellate cells are stained either lightly or not at
all. Thus the principal cells appear to be the major site of tyramine
production.
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DISCUSSION |
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I have shown that tyramine causes an increase in chloride conductance across the Drosophila Malpighian tubule. The pharmacology of this response matches very closely that of the cloned insect tyramine receptor, both in the selectivity of tyramine over other biogenic amines, particularly octopamine, and in the rank-order potency of several tyramine/octopamine antagonists (40, 42, 46, 50). This represents the first report of a physiological response with the same pharmacology as the tyramine receptor and the first well-defined role for tyramine in an insect tissue. Three previous studies have implicated tyramine in other aspects of insect physiology. First, tyramine has been reported to be necessary for the sensitization of Drosophila to repeated applications of cocaine (28), although this function has not yet been localized anatomically. In addition, two brief reports showed differential effects of tyramine, compared with octopamine, in stimulating trehalose metabolism in isolated cockroach fat bodies (18) and in inhibiting the contraction of locust visceral muscle (22). No additional pharmacological characterization was performed in either system. Although I have not shown conclusively in the Drosophila tubule that the cloned tyramine receptor is responsible for the increased chloride conductance, the expression of the receptor in the tubule and the close match in the pharmacology of the response with that of the cloned receptor make this highly likely to be the case.
A recent paper has reported the identification of flies carrying a mutation (hono) in the tyramine receptor gene (24). These flies contain a transposon inserted upstream of the first noncoding exon of the receptor gene; this exon is ~25 kb upstream of the rest of the gene. I have found that, although the tubules of hono flies are hyposensitive to tyramine, tubules from the parental stock, which does not contain the transposon insertion, are similarly hyposensitive (unpublished results). Indeed, tubules from many common laboratory stocks are hyposensitive to tyramine; the genetic basis for this phenotype is currently under investigation. It is possible that the hono insertion does not affect the expression of the tyramine receptor in the tubule, perhaps due to the utilization of an alternative transcriptional start site or other tissue-specific message processing.
A model for the synthesis and action of tyramine in the tubule is
presented in Fig. 11. Three features of
this model merit discussion. The first is the synthesis and release of
tyramine by the tubule. The evidence for this is as follows: first,
tyrosine and tyramine both increase chloride conductance, and the
actions of both are blocked by yohimbine with a very similar potency
(compare Figs. 4C and 7A), suggesting that both
stimulate the same receptor. Second, the D-tyrosine effect
presented in Fig. 9 argues against a direct action of tyrosine at the
tyramine receptor. Finally, the tubule contains significant TDC
activity and so is capable of converting tyrosine to tyramine. Thus
tyrosine must be taken up from the peritubular bath and decarboxylated
into tyramine. This tyramine must then be released; release is most
likely across the basolateral membrane because the tyramine receptor is
accessible to agents applied to the peritubular bath, although I cannot
exclude the possibility that tyramine can also act within the tubule
lumen. The molecular mechanisms governing the uptake of tyrosine and subsequent release of tyramine are unknown, although the data shown in
Fig. 9 suggest that one of these steps is inhibited by D-tyrosine.
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The second feature of the model is that the principal cells are the site of tyramine production and release. This is a likely but not definitive conclusion from the data. Figure 10 clearly shows that principal cells contain tyramine and that stellate cells do not stain strongly with the antibody, but it remains possible that stellate cells are also capable of synthesizing tyramine. Interestingly, incubation of tubules in the absence of tyrosine, which the electrophysiology suggests should deplete the "releasable" pool of tyramine, does not lead to a noticeable reduction in tyramine immunoreactivity (data not shown), demonstrating that most of the immunostaining does not represent the releasable pool of tyramine. Clearly then, the production of tyramine by the tubule is likely to be compartmentalized, and further studies are needed for its conclusive localization.
The third feature of the model is that tyramine binds to receptors on the stellate cells. Previously, I showed that the TEP oscillations seen in SBM were governed by calcium levels in the stellate cells; the oscillations are eliminated by chelation of intracellular calcium, and treatment of tubules with leucokinin, which increases calcium levels specifically in stellate cells (33, 41), causes a long-lasting suppression of the oscillations (7). The current work has demonstrated that the TEP oscillations are the result of the activation of tyramine receptors, thereby linking this signaling pathway, albeit indirectly, with intracellular calcium levels in the stellate cells. Consistent with this hypothesis, activation of the cloned insect tyramine receptor can cause increases in intracellular calcium levels in several heterologous systems (40, 42, 44). It now appears likely that the leucokinin-induced suppression of oscillations resulted from a cross-desensitization of the leucokinin and tyramine signaling pathways and that both pathways converge at some point downstream of receptor activation to stimulate chloride transport. The pathway for this chloride transport, paracellular or transcellular, is still unknown, and both possibilities are shown on the model.
The action of tyramine by the tubule bears a striking resemblance to
dopaminergic signaling in the mammalian kidney. In the latter case,
dopamine is employed in the proximal tubule as a paracrine or autocrine
hormone to regulate sodium transport (reviewed in Ref. 1).
The dopamine is produced from circulating DOPA by an aromatic amino
acid decarboxylase (AADC). Interestingly, AADC is also able to
decarboxylate tyrosine to produce tyramine. Although DOPA is the
preferred substrate, it has been calculated that due to the much higher
levels of tyrosine than DOPA in the circulation, tyramine and dopamine
should be produced in vivo at comparable rates (10).
Indeed, the kidney contains high levels of tyramine (36),
and a recently cloned mammalian tyramine receptor is expressed in the
kidney (9, 12). There are currently no specific
pharmacological tools for manipulating tyraminergic signaling in
vertebrates-yohimbine is also an -adrenergic
antagonist-but it is possible that tyramine will prove to be
important in vertebrate renal function as well.
The current work raises the possibility that chloride transport in the tubule may be more complicated than was previously appreciated. Until now, there has been no evidence for heterogeneity of chloride transport pathways; hence, agents that increase chloride conductance, such as leucokinin, are thought to increase the amplitude of the conductance pathway that is active in the unstimulated tubule (33, 34). The data shown in Fig. 8 suggest an alternative possibility; the tubule may possess multiple, pharmacologically separable pathways for chloride transport. At least one pathway is calcium sensitive and can be stimulated by leucokinin and tyramine. When tyraminergic signaling is blocked by yohimbine, however, there is still a significant level of chloride transport and urine secretion. This could represent a basal level of activation of the calcium-sensitive pathway, but it could also represent an additional, calcium-independent pathway. Unfortunately, the lack of specific pharmacological agents for blocking chloride conductance makes it difficult to distinguish between these two possibilities. It is hoped that the techniques available in Drosophila for the manipulation of gene expression may allow for the resolution of this issue.
Bathing solutions containing Schneider's medium have been used as the control medium in many studies involving Drosophila tubule function. It is clear from the current study that tubules bathed in SBM are not unstimulated relative to saline. Rather, SBM influences tubule physiology in at least two different ways; first, by increasing chloride conductance through the activation of tyramine receptors, and second, by increasing the amplitude of the TEP through an as-yet uncharacterized mechanism.
The stimulation of chloride conductance and urine secretion by tyramine is likely to be physiologically important in the intact fly and not simply an artifact of the recording conditions in vitro. Drosophila hemolymph contains a relatively high level of tyrosine (38), similar to that found in SBM, so the tyraminergic signaling pathway is almost certainly active in vivo. Moreover, because tyramine is a product of bacterial metabolism, it is likely to be present in the decomposing fruit on which the fly feeds. Food containing tyramine would therefore be expected to trigger a postfeeding diuretic response. In the locust, postfeeding diuresis is triggered by a CRF-like diuretic peptide hormone (3, 35); the response to tyramine in Drosophila would be different in that it would be mediated by a component of the food itself. Because the high rate of urine production by the tubules is thought to important for clearing toxins from the hemolymph (27, 37), it is tempting to speculate that the response to tyramine could have evolved to protect the fly from toxic substances that might be present in its food. Whatever the purpose, it is clear that tyramine represents an addition to the multiple signals already known to control renal function in insects.
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ACKNOWLEDGEMENTS |
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I thank Dr. Gene Block for support, Dr. Jay Hirsh and the members of his lab for helpful discussions, fly lines, equipment, and reagents, Dr. James Burnette for the tyramine receptor primers, Shannon Cole for assistance with the TDC assay, and Drs. Oliver Schneider and Robert Kelly for assistance with the capillary electrophoresis.
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FOOTNOTES |
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This work was supported by the University of Virginia and National Institute of Diabetes and Digestive and Kidney Diseases R21-DK-060860 to E. M. Blumenthal.
Address for reprint requests and other correspondence: E. M. Blumenthal, Dept. of Biology, Gilmer Hall, Univ. of Virginia, P.O. Box 400328, Charlottesville, VA 22904-4328 (E-mail: eb5f{at}virginia.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.
First published November 20, 2002;10.1152/ajpcell.00359.2002
Received 2 August 2002; accepted in final form 18 November 2002.
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