Voltages and resistances of the anterior Malpighian tubule of Drosophila melanogaster

The transepithelial electrical resistance is easily measured in a planar epithelium as the ratio of the transepithelial voltage deflection consequent to a transepithelial current pulse at uniform current density (Ohm's law). The current density cannot be uniform in a tubular epithelium unless a wire is passed through the tubule lumen, which so far has been accomplished only once in a renal epithelium. Taking advantage of the large size of the renal proximal tubule of the aquatic salamander Necturus maculosus, Spring and Paganelli (1972) were able to pass a platinized tungsten wire through the tubule lumen to measure a specific transepithelial resistance (R_t) of 43 Ω cm². All other measurements of R_t in tubular epithelia are grounded in cable theory (Taylor, 1963), which models the tubule as an electrical cable. The fluid in the tubule lumen is considered the conductive cable core and the surrounding epithelial cells are considered the insulator. Current is passed into the tubule lumen from a point source (microelectrode), and the transepithelial voltage deflections are recorded with microelectrodes at other points in the tubule lumen. The decay of the injected current along the length of the tubule depends on (1) the tubule geometry (length and lumen diameter), (2) the resistivity of the fluid occupying the tubule lumen and (3) the electrical properties of the epithelial cells, i.e. the transepithelial resistance.

The tubule may be modelled as a cable of finite length or infinite length. In the finite-length model, the two ends of the tubule are electrically isolated from the peritubular medium, defining the tubule length as in isolated perfused renal tubules (Burg et al., 1966; Helman, 1972). When the electrical isolation of the tubule ends is not possible, the tubule is considered a cable of infinite length (Boulpaep and Giebisch, 1978; Frömter, 1986; Hegel et al., 1967). For both models, variations of single- and double-barreled microelectrodes and measuring circuits have been employed to determine the cable parameters. In the present study, I introduce the use of three conventional microelectrodes in the tubule lumen to determine the tubule length constant, the transepithelial resistance and the core resistances in the main secretory segment of the anterior Malpighian tubule of Drosophila melanogaster Meigen 1830. Intact pairs of anterior Malpighian tubules still attached to a small piece of gut are isolated in Ringer’s solution and transferred to rest on the bottom of a perfusion bath (see Fig. 1). One microelectrode injects current into the tubule lumen of the main segment and two additional microelectrodes in the tubule lumen measure the transepithelial voltage deflections at different sites downstream. The data are then analyzed on the basis of the infinite-length cable model to yield specific and effective transepithelial and core resistances. The use of only two microelectrodes in the tubule lumen, one to inject current and the other to record voltage, yields relative resistances that are useful in qualitative before and after comparisons in a single tubule.

The usefulness of cable analysis in renal and Malpighian tubules is manifold. Cable analysis yields measures of the transepithelial electrical resistance that spans the range from leaky to tight epithelia. Leaky epithelia usually transport large volumes of fluid in isosmotic proportions, whereas tight epithelia are capable of separating solute from water, thereby forming either dilute or concentrated fluids on one side of the epithelium. Thus, cable analysis gives first clues about the general transport properties of the tubule. Supplemented with ion substitution in the bathing media, cable analysis yields measures of ionic permselectivity, and together with measurements of the intracellular voltages, cable analysis reveals the electromotive forces and resistances of basolateral and apical cell membranes. In brief, cable analysis is the necessary electrical experimental method for unravelling the mechanism and regulation of transepithelial electrolyte transport in tubular epithelia. In the present study, cable analysis has revealed a moderately tight epithelium in the main segment of the anterior Malpighian tubule of D. melanogaster and yielded an electrical model of transepithelial electrolyte secretion that is remarkably similar to the model of electrolyte secretion in distal tubules of Aedes aegypti when both tubules are studied under similar experimental conditions.

We considered white¹¹¹⁸ (RRID:BDSC 5905) as the wild type. The flies were reared under standard conditions at 22°C on cornmeal agar and a 12 h:12 h light:dark light cycle. Malpighian tubules of D. melanogaster are formed during the first 21 h of embryogenesis at 25°C, i.e. before the hatch of the first instar larva. Thereafter, the tubules are not rebuilt during metamorphosis and remain almost unmodified to the adult stage (Denholm et al., 2003; Singh et al., 2007; Sozen et al., 1997). The development of the tubule stops as the number of cells in the upper tubule (initial, transitional and main segments of the tubule) has reached approximately 150 cells (Singh et al., 2007). Of these, 80% are principal cells with ectodermal origin and 20% are stellate cells with mesodermal origin (Jung et al., 2005; Pugacheva and Mamon, 2003; Singh et al., 2007; Sozen et al., 1997).

Adult female flies at least 5 days old were cold-anesthetized on ice. Pairs of anterior Malpighian tubules were isolated with the cold-anesthetized fly submerged in Ringer’s solution as shown in the video produced in the laboratory of Rodan (Schellinger and Rodan, 2015). The main segment of the anterior tubule, which secretes most of the fluid arriving in the gut (Dow et al., 1994), was the focus of the present study.

The basic Ringer’s solution (BRS) contained (in mmol l⁻¹): 117.5 NaCl, 20 KCl, 2 CaCl₂, 8.5 MgCl₂, 10.2 NaHCO₃, 15 Hepes, 4.3 NaH₂PO₄ and 20 glucose. For measurements of transepithelial Cl⁻ diffusion potentials, the Cl⁻ concentration in BRS was reduced 10-fold, and included (in mmol l⁻¹): 117.5 Na-gluconate, 20 K-gluconate, 2 CaCl₂, 5.95 MgCl₂, 3.83 MgSO₄, 10.2 NaHCO₃, 15 Hepes, 4.3 NaH₂PO₄ and 20 glucose. The pH was adjusted to 7.0. The average [H⁺] of all solutions was (9.6±0.07)×10⁻⁸ mol l⁻¹ (n=50), pH 7.02. The osmotic pressures of the Ringer’s solutions containing 100% Cl⁻ and 10% Cl⁻ solution were, respectively, 335.6±1.2 (n=43) and 332.3±1.3 mOsm kg⁻¹ (n=6) H₂O. Drosokinin (JPT Peptides, Berlin, Germany) was used at a concentration of 1 µmol l⁻¹. To estimate the epithelial shunt resistance, R_sh, BRS was initially exchanged with BRS containing 2,4-dinitrophenol (DNP), KCN and NaN₃ (each 1 mmol l⁻¹). However, this inhibitory cocktail of ATP synthesis (and transcellular active transport) proved insufficient for estimates of R_sh owing to the high K⁺ conductance of basolateral membranes. For this reason, a low-K⁺ phosphate-free BRS containing 2 mmol l⁻¹ K⁺, 5 mmol l⁻¹ BaCl₂ and the above inhibitors of ATP synthesis was used to raise the transcellular resistance to such a degree that measures of the transepithelial resistance approached the resistance of the transepithelial shunt. The specific resistance of BRS was 59.2±0.2 Ω cm (n=20) measured with the WTW Multimeter 3430 and the TetraCon 925 conductivity probe (Xylem Analytics, Weilheim, Germany).

Although fluid secretion rates are highest in a 1:1 mixture of BRS and Schneider's medium, D. melanogaster Malpighian tubules are rather insensitive to the choice of saline (Dow et al., 1994). Because the use of Schneider's medium complicated the studies of Cl⁻ diffusion potentials, it was omitted in the present study.

Stable microelectrode impalements of epithelial cells and the tubule lumen required good immobilization of the Malpighian tubule in the perfusion bath. Glass cover slips were coated in three or more drying cycles with three short puffs of 0.1 mmol l⁻¹ poly-lysine (JPT Peptides Technologies GmbH, Berlin, Germany) delivered from a vaporizer and stored at 4°C. On the day of the experiment, 50 µl of BRS was deposited on the poly-lysine-coated cover slip to receive a pair of anterior Malpighian tubules still attached to a small piece of gut. Although the tubules stick immediately here and there to the glass, they can still be straightened out by handling the piece of gut (tubules and ureter were never handled with forceps). As shown in Fig. 1, slight stretching of the tubules forms a straight line of the main segment, which facilitates the impalement with a microelectrode approaching a principal cell or the tubule lumen at an angle of 45 deg or less in line with the tubule. The bath volume was brought up to 150 µl or 3 ml depending on the need to conserve reagent. Inflow and outflow lines allowed the change of the perfusion bath. The tubules were viewed under a Leica Stereoscope MZ95 at magnification ranging from 16× to 150×. The distances between microelectrode tips in the tubule lumen were measured with an ocular micrometer.

Microelectrodes were pulled with a Sutter P-97 Flaming/Brown microelectrode puller (Sutter Instruments, Novato, CA, USA) and filled with 3 mol l⁻¹ KCl. The electrical resistance for both current-injection and voltage electrodes was on average 48.9±0.9 MΩ (n=264). After each pull, the microelectrode with the lower resistance was used as the current/voltage electrode. For the measurement of voltage, the preamplifiers HS-2A Headstage Gain x1LU and HS-2A Headstage Gain x1MGU (Axon Instruments, Molecular Devices, San Jose, CA, USA) were used. The latter preamplifier was also used to pass current for the measurement of resistance. Ag/AgCl bridges in the microelectrodes and a 4% BRS agar bridge in the perfusion bath completed the measuring circuits. When both current and voltage electrodes were lodged in a principal cell (or in the lumen), the basolateral membrane resistance R_bl or the transepithelial resistance R_t, respectively, were measured in six 200 ms voltage clamp steps of 5 mV bracketing the basolateral membrane voltage V_bl or the transepithelial V_t, respectively. The current–voltage plots (I–V plots) were usually linear in both measurements of R_bl and R_t. A GeneClamp 500B along with the Digidata 1322A were used to record electrophysiological data (Molecular Devices). To monitor the quality of microelectrode impalements, voltages were recorded continuously at a frequency of 10⁷ Hz except for the brief periods of I–V plots, during which the voltage clamp was set at a gain of 1 K and a stability of 200 μs.

The measurement of the transepithelial resistance in a tubular epithelium requires cable analysis. The specific cable equations depend on the experimental condition. When the two ends of the tubule can be electrically isolated from the bathing medium, as in isolated tubules perfused by the method of Burg et al. (1966), the cable length is defined and cable analysis yields direct measurements of the length constant and the transepithelial and core resistances in renal tubules (Helman, 1972) and Malpighian tubules (Pannabecker et al., 1993; Williams and Beyenbach, 1984). When the tubule ends are not electrically isolated, as in studies of renal tubules in vivo or in isolated Malpighian tubules in vitro as in the present study, the tubule is considered a cable of infinite length (Boulpaep and Giebisch, 1978; Frömter, 1986). The infinite cable model requires that three microelectrodes are placed in the tubule lumen for the measurement of the tubule length constant and the transepithelial and core resistances (Fig. 2).

where r_e is the electrical radius of the tubule lumen. Because the optical radius of the tubule lumen measured through the microscope varies considerably with length along the tubule (see Fig. 1), sR_t was determined in the present study with known values λ, R_input, and ρ, the resistivity of secreted fluid in the tubule lumen; i.e. the second equivalence of Eqn 4 does not require knowledge of the lumen radius. The resistivity of secreted fluid in the tubule lumen was assumed to be similar to that of the peritubular Ringer’s solution in view of (1) the secretion of fluid isosmotic to the peritubular Ringer’s bath, and (2) the largely electrolyte composition of both peritubular and luminal fluids.

The use of only two microelectrodes in the tubule lumen yields the relative transepithelial conductance rg_t, as the ratio of I_o and V_t2, or its inverse, the relative transepithelial resistance rR_t (Fig. 2). The relative transepithelial resistance decreases with increasing distance between current and voltage electrodes as epithelial mass between the two electrodes increases. For this reason, measures of rR_t are relative and useful only in before/after comparisons in the same tubule (paired comparisons using each tubule as its own control).

Non-linear or distorted I–V plots resulted when the tip of one electrode is not free in the tubule lumen or in the cell as in touching a membrane. Currents as high as 300 nA were injected into the tubule lumen without ill effects on the tubule.

where V is voltage, R is resistance and the subscripts bl and a denote the basolateral and apical membrane, respectively. In the typical measurement of fR_bl, the current electrode E₀ and the voltage electrode E₂ were lodged in the tubule lumen, and the voltage electrode E₁ was in the cytoplasm of a principal cell (Fig. 2). The I–V plot yielded the voltage deflection across the basolateral membrane of the impaled principal cell (ΔE₁=ΔV_bl). Electrode E₁ was subsequently advanced across apical membrane into the tubule lumen. With all three microelectrodes located in the tubule lumen, the transepithelial I/V plot yielded λ and consequently ΔV_t at the site of the previously impaled principal cell for the determination fR_bl (Eqn 12).

Significant differences were evaluated using the Student's t-test for either unpaired or paired samples.

Studies with three microelectrodes yield resistances normalized to luminal surface area or tubule length for comparisons with Malpighian tubules from other species and other epithelia. The tubule lumen of Malpighian tubules is easily reached by passing the microelectrode through thin stellate cells (<5 μm). Passing a microelectrode through principal cells (>15 μm) usually yields the basolateral membrane voltage before reaching the tubule lumen (as in Fig. 5). Under control conditions when the tubules were bathed in BRS, the transepithelial voltage was 31.6±4.0 mV, the basolateral membrane voltage of principal cells was −39.3±3.8 mV, and the apical membrane voltage was 70.9±4.0 mV in 11 main segments of the anterior Malpighian tubule studied in Table 1.

According to the Ussing–Windhager conception of transepithelial transport, active and passive transport pathways are parallel across the epithelium (see Fig. 4A). The passive transport pathway is considered the shunt. In initial estimates of the shunt resistance, the cocktail of DNP, NaN₃ and KCN was added to the peritubular bath to inhibit ATP synthesis and hence active transcellular transport. The inhibition was expected to increase the transcellular resistance to such a degree that measures of the transepithelial resistance approach the resistance of the shunt as in in A. aegypti Malpighian tubules (Pannabecker et al., 1992). Not so in D. melanogaster Malpighian tubules: the inhibitor cocktail failed to significantly increase the effective transepithelial resistance (control, 14.2±2.0 kΩ cm; cocktail 16.1±3.0 kΩ cm; P<0.34, 13 anterior tubules). However, the inhibitory cocktail of ATP synthesis significantly brought all voltages towards zero with two kinetics of voltage depolarization (Fig. 3A). The first fast depolarization (phase 1, Fig. 3A) took off as soon as the cocktail was added to the peritubular medium and lasted about the time it took to change the bath, ∼25 s. In phase 1, V_a depolarized from 80 to 24 mV (lumen-positive), V_t depolarized from 33 mV and reversed polarity to −12 mV, and V_bl depolarized from −47 to −38 mV. Thereafter, in phase 2 of slow depolarization, voltages gradually depolarized and converged at values near 0 mV in the time of 196 s (Fig. 3A).

To elicit a sharp drop of all voltages together with a sharp increase in the transepithelial resistance, it was necessary to reduce the K⁺ conductance of basolateral membranes in addition to the inhibition of ATP synthesis. As shown in Fig. 3B, the inhibitory cocktail together with 5 mmol l⁻¹ Ba²⁺ in low K⁺ (2 mmol l⁻¹) BRS promptly depolarized all voltages in the time it took to replace the peritubular bath. Moreover, the K⁺ channel blocker Ba²⁺ in low K⁺ peritubular Ringer’s solution obliterated phase 2 of depolarization as well as the reversal of the transepithelial voltage to lumen-negative values. In subsequent studies of 11 main segment of anterior Malpighian tubules, the combined effects of metabolic inhibition and reduced K⁺ conductance significantly increased the tubule length constant from 540.0 to 772.4 µm and significantly increased the effective transepithelial resistance eR_t from 17.1 to 26.9 kΩ cm (Table 1). The lack of a significant effect on the effective core resistance eR_c and the electrical radius of the tubule lumen point to the successful inhibition of, specifically, the transcellular active transport pathway (Table 1).

The transepithelial secretion (or absorption) of electrolytes such as Na⁺, K⁺ and Cl⁻ carries current and generates voltages. Accordingly, the transepithelial secretion of electrolytes in the main segment of the anterior Malpighian tubule can be modelled with an electrical circuit consisting of two transepithelial routes for current flow: an active transport pathway leading through cells where active transport finds ATP, parallel to a passive transport pathway, the shunt that does not require ATP (Fig. 4A). In general, Na⁺ and K⁺ are actively transported through principal cells, and Cl⁻ is passively transported through stellate cells and/or the paracellular pathway (Beyenbach and Piermarini, 2011; O'Donnell et al., 1996; Pannabecker et al., 1993). The analysis of electrical equivalent circuits is useful in that it reveals (1) the driving forces for electrolyte secretion across the basolateral and apical membranes, and (2) the electrical resistances of all three barriers: basolateral membrane, apical membrane and shunt. Knowledge of these variables is useful for further dissection of transport pathways. For example, the analysis of an electrical equivalent circuit of Malpighian tubules in A. aegypti has shown inter alia that (1) the conductance of the basolateral membrane of principal cells is dominated by K⁺, (2) the conductance of the apical membrane is dominated by the V-type ATPase, and (3) the diuretic hormone leucokinin increases the Cl⁻ conductance of the shunt pathway (Beyenbach, 2012; Beyenbach and Masia, 2002; Beyenbach et al., 2000b; Beyenbach and Wieczorek, 2006; Pannabecker et al., 1993).

In the case of the electrical equivalent circuit of transepithelial electrolyte transport in anterior Malpighian tubules of D. melanogaster, the estimate of the shunt resistance allows estimates of the electromotive forces and resistances of the transcellular pathway under control conditions (Fig. 4). Effective rather than specific resistances are used from here on for comparison with Malpighian tubules of A. aegypti (Pannabecker et al., 1992). However, the purpose of this paper is not this comparison but to introduce a new experimental method: the measurement of resistance in tubular epithelia as small as Malpighian tubules of D. melanogaster using cable analysis and conventional microelectrodes rather than the method of in vitro microperfusion of tubules.

Because eR_t is 17.1 kΩ cm and eR_sh is 26.9 kΩ cm (Table 1), the effective transcellular resistance eR_cell is 46.9 kΩ cm (Fig. 4A).

Because V_t is 31.6 mV and eR_t is 17.1±2.4 kΩ cm (Table 1), I_sc is 1.85 μA cm⁻¹ and E_cell is 86.7 mV for a transcellular resistance of 46.9 kΩ cm (Fig. 4A). The fractional resistance of the basolateral membrane of principal cells was 0.33±0.05 in nine main segments of the anterior tubule (Eqn 12). Accordingly, the effective resistance of the basolateral membrane eR_bl is 15.5 kΩ cm, and the effective apical membrane resistance eR_a is 31.4 kΩ cm (Fig. 4B). Knowledge of these resistances yields the voltage drops across eR_bl and eR_a under open-circuit conditions as well as the electromotive force E of the basolateral and apical membrane when the tubule secretes electrolytes and fluid.

According to Eqn 13, the open-circuit current I_oc is 1.17 μA cm⁻¹ (Fig. 4A). Hence, the voltage drop across eR_bl is 18.2 mV (cell negative) and E of the basolateral membrane (E_bl) is cell-negative 21.1 mV (Fig. 4B). The voltage drop across eR_a is 36.9 mV and E of the apical membrane (E_a) is also cell-negative 107.8 mV (Fig. 4B).

Studies with two microelectrodes, one to inject current and the other to record voltage, yield relative resistances that are useful for comparing before/after effects using each tubule as its own control. As relative resistances are not normalized to tubule length or area, they should not be used for comparison with other tubules or epithelia.

Fig. 5 illustrates a representative experiment using two microelectrodes. The two microelectrodes are located within 600 μm of each other in the main segment of an anterior tubule; on average, the distance between the two electrodes was 357.4±30.1 μm in 25 tubules. The tip of the current–voltage microelectrode has entered the tubule lumen at point A to record the lumen-positive transepithelial voltage (V_t). V_t is usually constant with time, but oscillating voltages were observed occasionally. The tip of the other microelectrode has penetrated the basolateral membrane of a principal cell at point B to record the cell-negative basolateral membrane voltage (V_bl). Significantly, V_bl does not oscillate but remains constant near −50 mV. It follows that oscillations of V_t parallel oscillations of the apical membrane voltage (V_a). V_a was not measured directly but was determined as the difference between V_t and V_bl in accordance with Kirchhoff's law. At point C, the voltage microelectrode has been advanced across the apical membrane of the impaled principal cell into the tubule lumen. Both microelectrodes now measure V_t. With both microelectrode tips lodged in the tubule lumen, the relative transepithelial resistance (rR_t) is measured with a current–voltage plot. When V_t had hyperpolarized to 25 mV, rR_t was 122 kΩ; when V_t had depolarized to 10 mV, rR_t was 91 kΩ (Fig. 5).

In the experiment shown in Fig. 6, the tips of both microelectrodes were lodged in the tubule lumen of the main segment of an anterior Malpighian tubule and recorded similar values of the transepithelial voltage. Shortly before adding drosokinin (1 μmol l⁻¹) to the peritubular bath, V_t was 26 mV and rR_t was 109 kΩ. Upon the addition of drosokinin, V_t promptly dropped towards 0 mV; in parallel, rR_t dropped to 87 kΩ.

In paired t-tests, using each tubule as its own control, the addition of 1 μmol l⁻¹ drosokinin to the peritubular bath significantly hyperpolarized V_bl to −53.5±2.5 mV (n=4) (P<0.04), significantly depolarized V_t to 4.7±2.1 mV (n=11) (P<10⁻⁶), but had no significant effect on V_a 64.0±2.5 mV (n=3). The relative transepithelial resistance significantly decreased from 111.5 to 55.0±11.8 kΩ (n=25) (P<0.02).

Because the kinins – leucokinin in A. aegypti Malpighian tubules and drosokinin in D. melanogaster Malpighian tubules – are known to increase the permeability of Malpighian tubules to Cl⁻, the effect of drosokinin on transepithelial Cl⁻ diffusion potentials was of interest (Cabrero et al., 2014; Halberg et al., 2015; Lu et al., 2011; Pannabecker et al., 1993; Yu and Beyenbach, 2001; Yu and Beyenbach, 2002). A representative experiment is shown in Fig. 7. In BRS containing 158.5 mmol l⁻¹ Cl⁻, V_t was 25 mV and the rR_t 149 kΩ (Fig. 7, 9 min). The basolateral membrane voltage of a principal cell was −40 mV and the apical membrane voltage was 65 mV (data not shown). The 10-fold reduction of the peritubular Cl⁻ concentration had negligible effects on V_t and rR_t in the absence of drosokinin (Fig. 7, 9–31 min). The addition of drosokinin (1 µmol l⁻¹) to the peritubular bath had major effects on the transepithelial voltage and resistance. Drosokinin depolarized V_t from 25 to 4 mV and reduced rR_t from 161 to 49 kΩ (Fig. 7, 43 min). In the presence of drosokinin, the 10-fold reduction of the peritubular Cl⁻ concentration induced a large transepithelial Cl⁻ diffusion potential of 40 mV (Fig. 7, 67 min) and rR_t increased from 49 to 128 kΩ. After 44 min in the presence of 10% peritubular Cl⁻ concentration, the return to 100% Cl⁻ concentration in the peritubular bath returned V_t to 2 mV, consistent with the transepithelial electrical short circuit induced by drosokinin as rR_t decreased to 92 kΩ (Fig. 7, 97 min).

Measurements of the transepithelial electrical resistance in Malpighian tubules of D. melanogaster have not been thought feasible because of the small size of the tubules (Blumenthal, 2001). Since the time of this assessment, D. melanogaster Malpighian tubules have been successfully perfused in vitro, which renders them suitable for measurements of the transepithelial resistance (Wu et al., 2015). The present paper presents less demanding methods using conventional microelectrodes and commercially available electronic hardware. Resistance measurements normalized to luminal surface area or tubule length require the use of three microelectrodes. The use of only two microelectrodes yields relative resistances for before/after comparisons in a single tubule.

The specific transepithelial resistance of the main segment of the anterior tubule of D. melanogaster is 229.3 Ω cm² (Table 1). By comparison, sR_t of the renal proximal tubule, a leaky epithelium, is only 6 and 7 Ω cm² in the rat and dog, respectively, 30 Ω cm² in the rabbit gall bladder, 75 Ω cm² in frog choroid plexus, 100 Ω cm² in the rabbit ileum, between 150 and 300 Ω cm² in the rat distal tubule, 867 Ω cm² in rabbit cortical collecting tubules, 1500 Ω cm² in the toad urinary bladder, and 3600 Ω cm² in the frog skin (Boulpaep and Seely, 1971; Dor'o, 1968; Erlij, 1976; Frizzell and Schultz, 1972; Hegel et al., 1967; Helman et al., 1971; Malnic and Giebisch, 1972; Reuss and Finn, 1974; Ussing and Windhager, 1964; Wright, 1972). Accordingly, the specific transepithelial resistance of the main segment of the anterior Malpighian tubule of D. melanogaster falls in the range of renal distal tubules, namely moderately tight epithelia, reflecting selective barrier and transport properties consistent with a wide quantitative and qualitative range of transepithelial transport (Beyenbach, 1993a, 2016). Resistances can also be expressed in terms of tubule length (Eqns 8 and 9). Accordingly, the specific transepithelial resistance of 229.3 Ω cm² is equivalent to the effective transepithelial resistance of 17.1 kΩ cm (Tables 1 and 2, Fig. 4).

Table 2 lists the voltages and effective resistances of the main segment of the anterior Malpighian tubule of D. melanogaster measured in the present study. The transepithelial voltage, 31.6 mV, measured with microelectrodes is similar to that measured in the laboratory of Rodan using the method of in vitro microperfusion of Malpighian tubules (Wu et al., 2015), and the basolateral membrane voltage of principal cells, −45.3 mV, is similar to that measured in the laboratory of O'Donnell using double-barreled microelectrodes (Ianowski and O'Donnell, 2004). The ratio of the transepithelial voltage and the effective shunt resistance yields the open-circuit current I_oc (Eqn 13), namely the intraepithelial current when tubules are secreting solutes and water into the lumen. The I_oc of 1.17 μA cm⁻¹ is equivalent to a transepithelial flux of 12.1 pmol s⁻¹ cm⁻¹ monovalent ions. Accordingly, 12.1 pmol of cations pass per second through principal cells from bath to tubule lumen in a tubule segment 1 cm long, and 12.1 pmol of anions per second pass via the transepithelial shunt (paracellular pathway and/or stellate cells). Rates of the secretory cation flux measured in the laboratories of Dow, O'Donnell and Rodan are, on average, 89 pmol min⁻¹ and stem largely from the main segment that comprises approximately 60% of the tubule length (Dow et al., 1998; Linton and O'Donnell, 1999; O'Donnell and Maddrell, 1995; Rodan et al., 2012; Wu et al., 2015). Because the average D. melanogaster Malpighian tubule is 2.2 mm long (Rheault and O'Donnell, 2004), the flux of 89 pmol min⁻¹ for the whole tubule is equivalent to 11.2 pmol s⁻¹ cm⁻¹ tubule length. The latter is in good agreement with 12.1 pmol s⁻¹ cm⁻¹ measured by electrophysiological methods in the present study, and confirms (1) the cable analysis and the model of the tubule as a cable of infinite length, and (2) the largely electrogenic nature of transepithelial cation secretion.

According to the Ussing–Windhager conception of transepithelial NaCl transport by the frog skin, Na⁺ takes the active transport pathway in parallel with Cl⁻ through the shunt (Ussing and Windhager, 1964). The separation of transepithelial cation and anion transport applies widely to absorptive and secretory epithelia. In the case of insect Malpighian tubules in general and D. melanogaster Malpighian tubules in particular, the cations Na⁺ and K⁺ are secreted through principal cells, and Cl⁻, the counter ion of Na⁺ and K⁺, is secreted through the shunt (Beyenbach and Piermarini, 2011; O'Donnell et al., 1996). Thus, the transepithelial secretion of Na⁺, K⁺ and Cl⁻ in D. melanogaster Malpighian tubules can be modelled with an electrical circuit consisting of the electromotive force E_cell for the transcellular secretion Na⁺ and K⁺ in series with the transcellular resistance eR_cell, both parallel to the transepithelial shunt R_sh for Cl⁻ (Fig. 4A). An E_cell of 86.7 mV polarized to move positive charge (Na⁺, K⁺) into the tubule lumen is the sum of the electromotive forces of the basolateral and apical membranes for transcellular Na⁺ and K⁺ secretion (Fig. 4A). The distributed circuit reveals an electromotive force of 21.1 mV across the basolateral membrane of principal cells that reflects in part the K⁺ equilibrium potential across that membrane (Fig. 4B). The E_a of 107.8 mV across the apical membrane likely reflects the electromotive force of the V-type H⁺ ATPase located at that membrane.

In A. aegypti Malpighian tubules, the inhibition of ATP synthesis brings transepithelial electrolyte and fluid secretion to a halt and all voltages to zero in a matter of seconds (Beyenbach et al., 2000b; Pannabecker et al., 1992; Wu and Beyenbach, 2003). With transcellular cation secretion inhibited and eR_cell at a maximum, the transepithelial resistance approaches the resistance of the shunt, 16.8 kΩ cm (Table 2). This approach for estimating the shunt resistance is successful in epithelia powered largely, if not exclusively, by active transport pumps. The approach does not work in epithelia powered in addition by secondary active transport mechanisms and diffusion potentials. For example, the inhibition of ATP synthesis in the gall bladder has no effect on the transepithelial resistance, and it hyperpolarizes both basolateral and apical membrane voltages on account of ATP-sensitive K⁺ channels (Bello-Reuss et al., 1981). In the present study of D. melanogaster Malpighian tubules, the inhibition of ATP synthesis failed also to significantly increase the transepithelial resistance during the initial phase 1 of voltage depolarization because of the large K⁺ conductance of the basolateral membrane (Fig. 3A). The large K⁺ conductance accounts for nearly 75% of the basolateral membrane voltage and the lumen-negative transepithelial voltage during phase 1 (Fig. 3A). Subsequently, in phase 2 of slow depolarization, all voltages decay gradually to zero as K⁺ leaks to the peritubular bath in the absence of ATP (Fig. 3A). In order to reduce the K⁺ conductance of the basolateral membrane, the inhibitory cocktail was fortified with the K⁺ channel blocker barium and the peritubular K⁺ concentration was reduced to 2 mmol l⁻¹. As shown in Fig. 3B, this manoeuvre brought all voltages to zero in the time it took to replace the peritubular medium. At the same time, the transcellular resistance increased to values that allowed the estimate of the effective shunt resistance, 26.9 kΩ cm (Fig. 4, Tables 1 and 2).

Oscillations of the transepithelial voltage (V_t) are widely observed in Malpighian tubules of insects, among them Carausius (Pilcher, 1970), Locusta (Morgan and Mordue, 1981), Aedes (Williams and Beyenbach, 1984) and Drosophila (Davies et al., 1995). V_t oscillations are also observed in the present study, which paralleled oscillations of the transepithelial resistance as in A. aegypti Malpighian tubules (R_t) (Fig. 5). As V_t depolarized towards zero, R_t decreased, and as V_t hyperpolarized, R_t increased, as in A. aegypti Malpighian tubules, where the oscillations are dependent on the peritubular Cl⁻ concentration (Beyenbach et al., 2000a). In particular, as R_t decreases, V_t decreases towards the transepithelial Cl⁻ equilibrium potential in A. aegypti Malpighian tubules, revealing cyclical changes in transepithelial Cl⁻ conductance.

Whereas oscillations of V_t and R_t reflect transient changes in the shunt Cl⁻ conductance, drosokinin brings both voltage and resistance to lower constant values in D. melanogaster Malpighian tubules, as in A. aegypti Malpighian tubules in the presence of leucokinin (Fig. 6) (Miyauchi et al., 2011; Pannabecker et al., 1993; Schepel et al., 2010; Yu and Beyenbach, 2001). Drosokinin is the leucokinin of D. melanogaster. It has a single gene that encodes the longest known leucokinin with 15 amino acid residues (Radford et al., 2002). So far, only one drosokinin G protein-coupled receptor (GPCR; CG10626) has been identified in specifically stellate cells of Malpighian tubules of Drosophila melanogaster, Anopheles stephensi and Aedes aegypti (Lu et al., 2011; Radford et al., 2002; Radford et al., 2004). Binding to its GPCR at the basolateral membrane of stellate cells, drosokinin increases the intracellular [Ca²⁺] in selectively stellate cells in D. melanogaster Malpighian tubules (Cabrero et al., 2013; Cabrero et al., 2014; O'Donnell et al., 1998; Terhzaz et al., 1999). In a comprehensive study of the mechanism of action of drosokinin, Cabrero et al. (2014) offer strong evidence for localizing the kinin-activated transepithelial Cl⁻ shunt to stellate cells of D. melanogaster Malpighian tubules. Ca²⁺ imaging, Ramsay fluid secretion assays, measurements of transepithelial voltage, and transgenic chloride reporter technology are consistent with the chloride channel CLC-a at the apical membrane of stellate cells as part of the kinin-activated transepithelial shunt (Cabrero et al., 2014).

Table 2 compares the electrophysiological variables of the main secretory segments of Malpighian tubules in two dipterans, D. melanogaster and A. aegypti, separated by 189 million years of evolution (Chen et al., 2015). Every variable is significantly different. Though time for evolution could account for the differences, they can largely be attributed to the use of different peritubular K⁺ concentrations. Drosophila melanogaster Malpighian tubules were bathed in 20 mmol l⁻¹ K⁺ and A. aegypti Malpighian tubules in 3.4 mmol l⁻¹ K⁺ Ringer’s solution (Table 2). The physiological differences between D. melanogaster and A. aegypti Malpighian tubules disappear when both tubules are bathed in either low K⁺ or high K⁺ Ringer’s solutions.

In the presence of the usual 20 mmol l⁻¹ K⁺ in the Ringer’s solution, Malpighian tubules of D. melanogaster secrete KCl-rich fluid with a K⁺ concentration (180 mmol l⁻¹) 9-fold higher than the concentrations of Na⁺ (Dow et al., 1994; Dow et al., 1998; Linton and O'Donnell, 1999; O'Donnell and Maddrell, 1995; Rheault and O'Donnell, 2001; Rodan et al., 2012). However, in K⁺-free Ringer’s solution, the tubules secrete a NaCl-rich fluid containing 150 mmol l⁻¹ Na⁺, like A. aegypti Malpighian tubules (Linton and O'Donnell, 1999). What is more, in K⁺-free Ringer’s solution, cAMP stimulates fluid secretion by 48%, presumably via the stimulation of transepithelial Na⁺ secretion, as in A. aegypti Malpighian tubules (Beyenbach, 1993b; Beyenbach, 2003; Beyenbach and Petzel, 1987; Linton and O'Donnell, 1999; Williams and Beyenbach, 1983). The corollary is observed in A. aegypti Malpighian tubules.

As shown in Fig. 8A, Malpighian tubules of A. aegypti secrete NaCl-rich fluid (151 mmol l⁻¹) when bathed in Ringer’s solution containing 3.4 mmol l⁻¹ K⁺ (Hine et al., 2014). However, in the presence of 34 mmol l⁻¹ K⁺ (Fig. 8B), the tubules secrete a KCl-rich fluid like D. melanogaster Malpighian tubules (Hine et al., 2014). In parallel, the basolateral membrane voltage of 12 principal cells significantly depolarizes from −76.5±4.4 to −41.2±1.7 mV (P<10⁻⁶), similar to the depolarization of V_bl in D. melanogaster Malpighian tubules (Table 2), and the basolateral membrane resistance of principal cells significantly decreases from 258.6±13.6 to 165.2±9.2 kΩ (P<10⁻⁵) in the same cells. Thus, Malpighian tubules of both A. aegypti and D. melanogaster respond to peritubular Na⁺ by increasing transepithelial Na⁺ secretion at the expense of K⁺ secretion, and respond to peritubular K⁺ by increasing transepithelial K⁺ secretion at the expense of Na⁺ secretion (Fig. 8). The tubules do so spontaneously in vitro without signalling by extracellular natriuretic or kaliuretic agents. Aedes aegypti evolved 189 million years after D. melanogaster. Thus it appears that the capacity for secreting K⁺ and Na⁺ by D. melanogaster Malpighian tubules has allowed females of A. aegypti to adopt hematophagous habits and to excrete the Na⁺ load of blood meals because their Malpighian tubules have retained ancestral mechanisms.

Thanks are due to Frederike Schöne and Laura Momann for the study of Aedes aegypti Malpighian tubules in 34 mmol l⁻¹ K⁺ Ringer’s solution; Leonard Breitsprecher for providing the image in Fig. 1; Dr Heiko Harten and Prof. Dr Achim Paululat for providing Drosophila melanogaster; and the University of Osnabrück for providing space and equipment under the auspices of my guest professorship in the Department of Biology/Chemistry.