Thiazolidinedione-Induced Fluid Retention Is Independent of Collecting Duct αENaC Activity

Abstract

Thiazolidinediones are agonists of peroxisome proliferator–activated receptor γ (PPARγ) that can induce fluid retention and weight gain through unclear mechanisms. To test a proposed role for the epithelial sodium channel ENaC in thiazolidinedione-induced fluid retention, we used mice with conditionally inactivated αENaC in the collecting duct (Scnn1a^loxloxCre mice). In control mice, rosiglitazone did not alter plasma aldosterone levels or protein expression of ENaC subunits in the kidney, but did increase body weight, plasma volume, and the fluid content of abdominal fat pads, and decreased hematocrit. Scnn1a^loxloxCre mice provided functional evidence for blunted Na⁺ uptake in the collecting duct, but still exhibited rosiglitazone-induced fluid retention. Moreover, treatment with rosiglitazone or pioglitazone did not significantly alter the open probability or number of ENaC channels per patch in isolated, split-open cortical collecting ducts of wild-type mice. Finally, patch-clamp studies in primary mouse inner medullary collecting duct cells did not detect ENaC activity but did detect a nonselective cation channel upregulated by pioglitazone. These data argue against a primary and critical role of ENaC in thiazolidinedione-induced fluid retention.

Peroxisome proliferator–activated receptors (PPARs) are ligand-activated transcription factors that regulate a large number of diverse genes by transcriptional activation and repression.¹ Thiazolidinediones (TZDs) are potent ligands and activators of PPAR-γ and are insulin sensitizers used extensively to treat type 2 diabetes.² PPAR-γ is also expressed in other tissues, such as the vasculature, where TZDs affect the inflammatory response and cholesterol homeostasis.^3–5 An important clinical limitation for this class of drugs is fluid retention and edema formation, which occurs in up to 5% of patients who have diabetes and are treated with TZDs; as a consequence, these agents are contraindicated in patients with New York Heart Association classes III and IV congestive heart failure.^6,7

Investigators have proposed that TZD-induced edema formation relates to increases in vascular permeability^7–10 and vasodilation.^11,12 Lower BP could reduce renal perfusion pressure, thereby increasing renal reabsorption of Na⁺ and fluid.^13–15 Conversely, two articles reported that, in mice, genetic knockdown of PPAR-γ in collecting ducts (CDs) abrogated the ability of the TZDs pioglitazone and rosiglitazone to increase plasma volume and cause weight gain.^16,17 Moreover, TZD-induced weight gain was blocked by amiloride,¹⁷ which is an inhibitor of the epithelial sodium channel (ENaC). In addition, treating cultured wild-type mouse inner medullary CD (IMCD) cells in vitro with TZDs increased amiloride-sensitive Na⁺ absorption as well as γENaC expression.¹⁷ The investigators showed that PPAR-γ bound to intron 1 of the γENaC gene, and they suggested that TZD-induced upregulation of γENaC was associated with derepression of the γENaC gene. These effects were inhibited by the PPAR-γ antagonist GW9662 and in IMCD cells derived from floxed PPAR-γ mice transduced in vitro with adenoviral Cre. On the basis of these findings, the authors concluded that TZDs expand body fluid volume through PPAR-γ stimulation of ENaC-mediated renal salt absorption in the CD.¹⁷

The PPAR-γ agonist farglitazar also increased blood volume, and this effect was associated with increased renal expression of the α subunit of Na-K-ATPase and aquaporin 2 (AQP2), whereas the renal expression of the ENaC subunits was not significantly altered, and amiloride did not prevent blood volume expansion.¹⁸ Similarly, rosiglitazone was reported to increase whole-kidney protein abundance of the α subunit of Na-K-ATPase as well as AQP2, whereas ENaC subunits were not affected; in addition, rosiglitazone increased the expression of the Na⁺-H⁺ exchanger NHE3, the Na-K-2Cl co-transporter NKCC2, and AQP3.¹⁵ These data indicate that activation of renal Na⁺ reabsorption pathways other than ENaC and of water reabsorption through water channels may also contribute to TZD-induced Na⁺ and fluid retention.

Considering the diversity of results, an editorial concluded that the molecular mechanisms and the renal sites of action involved in TZD-induced fluid retention, including a role for ENaC in the CD, have not been clearly established.¹⁹ In this study, we therefore assessed the role of ENaC in the CD in TZD-induced Na⁺ and fluid retention by performing studies of mice that lack αENaC (Scnn1a) selectively in the CD.^20,21 Cross-breeding mice with a floxed Scnn1a allele (Scnn1a^loxlox) to a Hox7:Cre-expressing mouse line generated Scnn1a^loxloxCre mice. In these mice, αENaC was selectively inactivated in the CD but not in the early segments of the aldosterone-sensitive distal nephron, namely the late distal convoluted tubule (DCT) and connecting tubule (CNT).²⁰ These mice apparently lack the expression of α, β, and γ subunits of ENaC in apical membranes of the CD, consistent with a critical role of αENaC for exporting the ENaC complex to the apical membrane. As a consequence, ENaC activity, as measured by patch-clamp experiments, was not detectable in principal cells of the CD in Scnn1a^loxloxCre mice. Furthermore, this inactivation in the CD did not impair Na⁺ and K⁺ balance,²⁰ consistent with the concept that the DCT and CNT, rather than the CD, are the dominant physiologic sites of aldosterone-regulated renal Na⁺ and K⁺ excretion.^22–25 In addition to using this model, we performed patch-clamp experiments to assess directly the effect of pretreating wild-type mice (C57BL/6; the overall genetic background strain of the mutants used in this study) with rosiglitazone and pioglitazone on ENaC activity in isolated CDs. Finally, we investigated the effects of pioglitazone on Na⁺-conducting channels in primary mouse IMCD cells.

RESULTS

Rosiglitazone Does not Alter Renal ENaC Protein Expression

Western blot analysis under control diet showed that the expression of αENaC in the inner medulla was significantly reduced in Scnn1a^loxloxCre compared with Scnn1a^loxlox mice (Figure 1A), in accordance with efficient knockdown in the CD. In comparison, whole-kidney expression of αENaC was not altered in Scnn1a^loxloxCre compared with Scnn1a^loxlox mice (Figure 1A), which is consistent with the dominant expression of this isoform in the early aldosterone-sensitive distal nephron upstream of the CD. Also, a modest compensatory upregulation of αENaC in Scnn1a^loxloxCre in the early aldosterone-sensitive distal nephron cannot be excluded. A significant upregulation of the renal expression of both βENaC and γENaC was detected in Scnn1a^loxloxCre compared with Scnn1a^loxlox mice. On the basis of a previous immunohistologic analysis,²⁰ this could be the consequence of accumulation of these two subunits in the intracellular compartment of the CD as a result of the absence of αENaC. In comparison, the renal expression of the Na-Cl co-transporter (NCC) was not different between genotypes.

Figure 1.

Rosiglitazone (RGZ) does not alter whole-kidney protein expression of ENaC or NCC. (A) Expression of αENaC in the inner medulla was significantly reduced in Scnn1a^loxloxCre compared with Scnn1a^loxlox mice, in accordance with efficient knockdown in the CD. In comparison, whole-kidney expression of αENaC and NCC was not different between genotypes under basal conditions, whereas βENaC and γENaC were greater in Scnn1a^loxloxCre mice; because of the low level of expression of the 70-kD subunit of γENaC, any conclusion on differences between groups must be considered with caution. *P < 0.05 versus Scnn1a^loxlox; #P = 0.06 versus Scnn1a^loxlox; n = 3. (B) RGZ treatment for 11 d did not significantly alter the renal expression of these proteins in either genotype. For each genotype, mean values of mice on control diet were set at 100%. A value of 1 means no change in response to RGZ compared with control diet.

Treatment for 11 d with rosiglitazone (320-mg/kg diet¹⁶) did not significantly alter the renal expression of ENaC subunits or NCC in Scnn1a^loxlox or Scnn1a^loxloxCre mice (Figure 1B). Plasma levels of Na⁺, K⁺, and aldosterone were not significantly different between Scnn1a^loxloxCre and Scnn1a^loxlox mice fed control diet (143 ± 2 versus 143 ± 2 mM; 4.77 ± 0.13 versus 4.54 ± 0.11 mM; 406 ± 66 versus 344 ± 27 pg/ml; n = 6 to 8; NS) and were not significantly affected by rosiglitazone treatment or different between the genotypes under these conditions (147 ± 1 versus 145 ± 1 mM; 4.89 ± 0.13 versus 4.74 ± 0.22 mM; 376 ± 38 versus 337 ± 45 pg/ml; n = 6 to 8; NS).

Lithium-Induced Diuresis Is Blunted in Scnn1a^loxloxCre Mice Independent of Rosiglitazone Treatment

Application of lithium downregulates AQP2 in the CD and induces diuresis.²⁶ Previous studies of Scnn1a^loxloxCre mice indicated that the uptake of lithium into the CD via apical ENaC is critical for the development of diuresis.²⁷ Whereas lithium increased fluid intake and reduced urine osmolality in Scnn1a^loxlox mice (0.17 ± 0.01 to 0.40 ± 0.09 ml/24 h per g body wt, 1684 ± 186 to 667 ± 105 mOsm/kg; n = 6; P < 0.01 for both comparisons), we confirmed that this response is lacking in our set of Scnn1a^loxloxCre mice (0.20 ± 0.02 to 0.21 ± 0.02 ml/24 h per g body wt, 1449 ± 187 to 1486 ± 152 mosm/kg; n = 8; NS) consistent with blunted ENaC and Li⁺/Na⁺ uptake activity in the CD of these mice. Moreover, another set of Scnn1a^loxloxCre mice was pretreated with rosiglitazone as described already. After 7 d of rosiglitazone, lithium treatment was added. Also under these conditions, lithium was without significant effect on fluid intake or urine osmolality in Scnn1a^loxloxCre mice (0.18 ± 0.02 to 0.18 ± 0.02 ml/24 h per g body wt 1742 ± 130 to 1769 ± 111 mosm/kg; n = 7; NS), indicating that rosiglitazone did not seem to activate a Li⁺/Na⁺ uptake pathway in the CD of Scnn1a^loxloxCre mice. Thus, we propose that this mouse model is suitable for testing the role of ENaC and Na⁺ reabsorption activity in the CD in rosiglitazone-induced fluid retention.

Rosiglitazone-Induced Fluid Retention and Weight Gain Is not Attenuated in Scnn1a^loxloxCre Mice

Basal body weight was not different between Scnn1a^loxloxCre and Scnn1a^loxlox mice (27 ± 1 versus 26 ± 1 g; n = 22; NS). In Scnn1a^loxlox mice, rosiglitazone increased body weight, plasma volume (assessed with intravenous application of Evans Blue) and the fluid content of abdominal fat pads, and lowered hematocrit (Figure 2). The observed increase in the fluid content of abdominal fat was proposed to relate an increase in vascular permeability in response to TZDs.⁹ All of these rosiglitazone-induced changes were not attenuated in Scnn1a^loxloxCre mice (Figure 2).

Figure 2.

RGZ-induced increases in body weight, plasma volume, and abdominal fat fluid content and the reduction in hematocrit are observed in Scnn1a^loxlox mice and not attenuated in Scnn1a^loxloxCre mice. Plasma volume, abdominal fat fluid content, and hematocrit were assessed after 11 d of treatment (n = 6 to 8/group each parameter); n = 10 to 12 per group for changes in body weight. *P < 0.05 versus control diet.

Rosiglitazone or Pioglitazone Treatment Does not Affect ENaC Activity in CD

Figure 3 shows the effects of 10-d treatments of two distinct TZDs, rosiglitazone (as described in the previous section) and pioglitazone (220-mg/kg diet¹⁷), on the activity of ENaC in native principal cells in freshly isolated, split-open cortical CD (CCD) of C57BL/6 mice. As indicated by the representative single-channel current traces in Figure 3A, the TZDs had no significant effect on ENaC. Indeed, ENaC open probability (P_o) in the two TZD test groups was not different compared with control (Figure 3B). Similarly, TZDs did not significantly affect the mean number of active ENaC (N) within patches (Figure 3B). Moreover, TZDs failed to change the chance of seeing active ENaC in membrane patches with 32 to 42% of the patches from all groups containing at least one active channel. These results demonstrate that TZDs do not change ENaC P_o and N in the CD and thus do not affect the activity (NP_o) of this channel, which is the product of these two variables.

Figure 3.

RGZ and pioglitazone (PGZ) have no effect on ENaC activity in native CCD. (A) Current traces from cell-attached patches formed on the apical plasma membrane of principal cells in split-open CD isolated from C57BL/6 mice maintained with control chow (0.4% Na⁺) and chow supplemented with RGZ or PGZ for 10 d. These representative patches contain at least two ENaC. Patches clamped at −V_p = −60 mV with inward Li⁺ current downward. Dashed lines note closed states. (B) Summary graphs showing the effects of RGZ or PGZ on ENaC P_o and the mean number of ENaC within patches (N) under each condition. Data collected from eight to 26 patches in three to four mice for each condition.

In contrast to TZDs and performed as a control experiment, manipulating dietary Na⁺ intake did affect ENaC activity as expected.^28–31 Figure 4A shows representative single-channel current traces for ENaC in membrane patches formed on native principal cells from CDs isolated from mice fed a nominally Na⁺-free diet and one enriched in Na⁺. As summarized in Figure 4B, ENaC P_o was significantly increased and decreased, respectively, in the low- and high-Na⁺ diet test groups as compared with control. Similarly, the mean number of active ENaCs within patches was significantly increased in the low-Na⁺ group and trended lower in the high-Na⁺ group compared with control. These results confirm that ENaC in the CD is sensitive to dietary Na⁺ and, thus, reinforces the previously mentioned findings that dietary supplementation with TZD does not change ENaC activity.

Figure 4.

Manipulating dietary salt intake affects ENaC activity in native CCD. (A) Current traces from cell-attached patches formed on the apical plasma membrane of principal cells in split-open CDs isolated from C57BL/6 mice maintained with nominally Na⁺ free chow (<0.01% Na⁺, low) and chow enriched in Na⁺ (2.0% Na⁺). All other conditions the same as in Figure 3. (B) Summary graphs showing the effects of manipulating dietary salt intake on ENaC P_o and N. Data collected from 79 patches in 15 mice on low-Na⁺ diet, 20 patches in five mice on control diet (0.32% Na⁺), and nine patches in four mice on high-Na⁺ diet. *P < 0.05 versus 0.32% Na⁺ diet.

Pioglitazone Treatment Increases the Activity of a Nonselective Cation Channel in Primary Mouse IMCD Cells

A previous study proposed that pioglitazone-induced increases in amiloride-sensitive Na⁺ reabsorption across primary mouse IMCD (mIMCD) cells are mediated by ENaC¹⁷; therefore, we investigated the effects of pioglitazone on Na⁺-conducting channels in primary wild-type mIMCD cultures. In the experiments in CCDs outlined in the previous section, the primary Na⁺-conducting ion channel in the apical plasma membrane was ENaC, which is highly Na⁺ selective and has slow gating kinetics and a small (approximately 6 to 8 pS) single-channel conductance. In contrast, the only Na⁺-conducting channel observed at a reliable frequency (in 47% of the patches made) in mIMCD cells was an approximately 23-pS nonselective cation (NSC) channel. Figure 5, A and B, shows representative current traces at hyperpolarizing and depolarizing holding potentials and the single-channel current-voltage relation, respectively, for the NSC channel observed in mIMCD cells. As is clear, this NSC channel is biophysically distinct from ENaC, having no preference for Na⁺ and Li⁺ over K⁺ and having a single-channel conductance that is approximately three times larger than ENaC. This NSC channel has been described before in rat IMCD and mIMCD by electrophysiology,^32,33 but the molecular correlates encoding NSC channels have not been identified.

Figure 5.

PGZ increases NSC channel activity in primary mIMCD cells. (A) Current trace from a representative cell-attached patch containing at least two NSC channels formed on the apical plasma membrane of a freshly isolated mIMCD cell within a polarized monolayer grown on permeable supports. The holding potential (−V_p) is indicated at the right with all other conditions identical to Figure 3. (B) Single-channel I-V relation for NSC channels in mIMCD cells (n = 7). (C) Current traces from cell-attached patches containing NSC channels formed on the apical plasma membrane of freshly isolated mIMCD cells within polarized monolayers grown on permeable supports in the absence (time control) and presence of 1 μM PGZ for 12 or 48 h. Patches clamped at −V_p = −60 mV. Summary graphs showing the effects of PGZ on NSC channel P_o (D) and the mean number of NSC channels within patches (N; E) under each condition.

Figure 5, C through E, shows the effects of treatment for 12 and 48 h with pioglitazone (1 μM) on the activity of the NSC channel in freshly isolated mIMCD cells. As indicated by the representative single-channel current traces in Figure 5C, pioglitazone markedly increased NSC channel activity after 48 h of treatment. Whereas NSC channel P_o in the pioglitazone test groups was not different compared with control (Figure 5D), the mean number of active NSC channels within a patch after 48 h of treatment was significantly increased (Figure 5E).

DISCUSSION

This study provides evidence that the rosiglitazone-induced fluid retention and body weight increase can occur independent of ENaC activation and Na⁺ reabsorption in the CD of the kidney. The evidence includes the observation that a mouse model with inactivation of ENaC activity in the CD responds to rosiglitazone with fluid retention and elevations in body weight just as well as control mice. Absence of lithium-induced diuresis in Scnn1a^loxloxCre mice in response to rosiglitazone argues against significant activation of ENaC-independent Na⁺ reabsorption in the CD. Moreover, rosiglitazone-induced fluid retention and increased body weight was not associated with significant changes in plasma aldosterone or renal expression of the ENaC subunits α, β, and γ, confirming previous reports.^15,18 Finally, treatment with rosiglitazone or pioglitazone did not significantly alter ENaC activity when directly assessed by patch-clamp analysis in split-open CDs of C57BL/6 mice.

These data are in accordance with previous in vitro studies in the A6, M-1, and mpkCCD(cl4) cell lines, which have been extensively used to elucidate the regulation of ENaC activity. Although immunodetection and quantitative PCR analyses showed that each of these three cell lines expresses viable and functional PPAR-γ, electrophysiologic studies revealed that both the basal and the insulin-stimulated Na⁺ flux via ENaC was insensitive to the PPAR-γ agonists pioglitazone and GW7845.³⁴

These results do not exclude the possibility that, under some conditions, aldosterone-induced ENaC activity may contribute to TZD-induced fluid retention. This may happen when TZD effects on vascular permeability and tone lower BP sufficiently to increase the activity of the renin-angiotensin-aldosterone system.³⁵ These data, however, argue against a primary and critical role for ENaC and Na⁺ reabsorption in the CD in TZD-induced fluid retention.

How can this study be reconciled with previous studies that proposed a critical role for ENaC in the CD in TZD-induced fluid retention.^16,17 In the previous studies investigating the mechanisms underlying TZD-induced fluid retention, the relevance of ENaC in vivo, in intact CDs or in cultured IMCD cells, was not conclusively assessed, but the analysis relied on amiloride as a pharmacologic tool rather than genetic or electrophysiologic approaches to identify a contribution of ENaC. This study confirms that pioglitazone can activate a Na⁺ conductance in primary mIMCD cells. This conductance, however, is mediated by a 23-pS NSC channel, which is biophysically very much distinct from ENaC. Previous studies described these NSC channels in rat IMCD cells in primary culture as well as in the mIMCD-3 cell with single-channel conductivities of approximately 27 pS³⁶ and approximately 24 pS,³³ respectively. Importantly, these channels are amiloride sensitive at concentrations ≤5 to 25 μM^33,36 (i.e., in the concentration range used in previous studies to inhibit pioglitazone-induced Na⁺ currents in primary IMCD cells¹⁷).

Studies in primary IMCD cells implicated regulation of NSC channels by atrial natriuretic peptide³² and G proteins³⁷ and proposed a role in electrogenic Na⁺ reabsorption; however, nonselective cation channels could not regulate Na⁺ and K⁺ reabsorption independently, and, therefore, the participation of these channels in Na⁺ reabsorption in the IMCD was called into question.³³ Because NSC channels in the mIMCD-3 cell line are activated by applied negative pressure and hypotonicity, they were proposed to contribute to volume regulation.³³ This study identified NSC channels as a new target of pioglitazone. Their physiologic significance in IMCD and their contribution to TZD-induced fluid retention remain to be determined. A previous study indicated that TZDs can induce mRNA expression of γENaC in IMCD cells.¹⁷ Whether TZDs induce channels that are biophysically different from the “classical” ENaC channel but involve γENaC and that are important for fluid retention remains to be determined.

Clear evidence was provided in previous studies that renal segment-specific knockdown of PPAR-γ prevented rosiglitazone- and pioglitazone-induced fluid retention and weight gain.^16,17 Because in this study PPAR-γ knockdown was driven by the AQP2 promoter, PPAR-γ was inactivated in the CD and possibly in parts of the upstream CNT; therefore, one could speculate that PPAR-γ–induced ENaC in the CNT is critical for TZD-induced fluid retention. Indirect evidence against this is the absence of effects of rosiglitazone or pioglitazone on ENaC activity in CCDs of C57BL/6 mice, assuming that ENaC responses are quantitatively but not qualitatively different in the CNT and the CCD. The changes in the CCD may be smaller compared with the CNT and late DCT, but the direction of the ENaC response, at least to aldosterone, is the same.^22,24,25 In accordance, also the CCD responds to changes in salt intake with adaptations in ENaC activity, as shown in this study.

Another possible explanation for the different findings includes the local formation of factors under the control of PPAR-γ activation, which may be released from CD or CNT cells that affect Na⁺ transport in other nephron segments. Notably, kidneys of PPAR-γ knockdown mice seem to have a reduced renal Na⁺ avidity that is compensated by increased plasma aldosterone levels.¹⁷ To which extent an impaired ability to retain salt and maintain body fluids and circulating volume per se (independent of the involved mechanism) will affect the ability of TZDs to induce and maintain fluid retention and increase body weight remains to be determined.

In summary, this study indicates that classical ENaC-mediated Na⁺ reabsorption in the CD is not critical for TZD-induced fluid retention and weight gain.

CONCISE METHODS

Animal use and welfare adhered to the National Institutes of Health Guide for the Care and Use of Laboratory Animals following a protocol reviewed and approved by the Institutional Laboratory Animal Care and Use Committee at the Veterans Affairs San Diego Healthcare System and the University of Texas Health Science Center at San Antonio.

Scnn1a^loxlox and Scnn1a^loxloxCre Mice

Female Scnn1a^loxlox and male Scnn1a^loxloxCre mice were bred to generate experimental Scnn1a^loxlox (control) and Scnn1a^loxloxCre mice.²⁰ Age- and gender-matched mice were used for the outlined experiments. Mice were housed with free access to food (1% K⁺, 0.4% Na⁺; Harlan TEKLAD TD.7001-repelleted [control diet] Madison, WI) and tap water. Rosiglitazone (320 mg/kg¹⁶) was added to this diet and the diet repelleted.

Western Blot Analysis

After receiving rosiglitazone-supplemented diet or control diet for 11 d, mice were anesthetized with isoflurane and whole kidneys were harvested and prepared for Western blot analysis as described previously.^38,39 Briefly, kidneys were homogenized in a buffer containing 250 mM sucrose, 10 mM triethanolamine (Sigma-Aldrich, St. Louis, MO), and Complete Protease Inhibitor cocktail (Roche, Indianapolis, IN) using a tissue homogenizer (Tissumizer; Tekmar, Cincinnati, OH). Homogenates were centrifuged at 16,000 × g for 30 min to obtain a membrane pellet. The pellet was washed once with homogenizing buffer and centrifuged again. The pellet was resuspended in homogenizing buffer, and protein concentration was determined using a DC Protein Assay (Bio-Rad, Hercules, CA) to ensure equal protein loading (20 μg per lane for αENaC, βENaC, and γENaC and 5 μg per lane for NCC). Samples were diluted in 4× LDS reducing sample buffer (Invitrogen, Carlsbad, CA), heated for 10 min at 70°C, and loaded on 4 to 12% Bis/Tris MOPS (ENaC) or 3 to 8% TA (NCC) precast SDS-PAGE gels (Invitrogen). After electrophoretic separation, proteins were blotted to a nitrocellulose membrane and stained with a Ponceau S solution (Sigma-Aldrich) to verify equal protein loading and transfer. After destaining, the membrane was blocked with 5% nonfat dry milk (Bio-Rad) in PBS (pH 7.4) containing 0.1% Tween 20 (PBS-T) for 1 h. Immunoblotting was performed at 4°C overnight with the primary antibody (rabbit anti-αENaC 1:3000,⁴⁰ rabbit anti-βENaC 1:750,⁴¹ rabbit anti-γENaC 1:750,⁴¹ and rabbit anti-NCC 1:2000 [Chemicon Int., Temecula, CA²⁴]) diluted in PBS Tween containing 1% BSA. The antisera to ENaC subunits were gifts from A. Doucet (Institut des Cordeliers, Paris, France; αENaC) and L. Palmer (Weill Medical College of Cornell University, Ithaca, NY; βENaC and γENaC). Chemiluminescent detection was performed using a 1:5000 dilution of ECL donkey anti-rabbit IgG linked to horseradish peroxidase and ECL detection reagent (GE Healthcare, Buckinghamshire, UK). Densitometric analysis was performed using NIH ImageJ Software.

Body Weight, Plasma Volume, Hematocrit, and Fluid Content of Abdominal Fat Pads

Mice were kept in standard rodent cages, and the body weight was determined before and during 11 d of treatment with rosiglitazone. Subsequently, some of the mice were subjected to assessment of plasma volume using Evans Blue application. Briefly, Evans Blue (1.5 mg/ml in 0.85% NaCl; 1 μl/g body wt) was applied into the retroorbital vein plexus of the left eye under isoflurane anesthesia. After 7 min of distribution, the retroorbital vein plexus of the right eye was punctured to collect plasma for determination of Evans Blue concentrations (absorbance at 620 nm; the injection solution was diluted with naive mouse plasma to generate standards), and calculation of plasma volume was based on the volume of distribution of Evans Blue.¹⁶ In other mice, blood was withdrawn by puncturing of the retrobulbar plexus under brief isoflurane anesthesia for determination of hematocrit and plasma Na⁺, K⁺, and aldosterone (RIA, DSL-8600; Diagnostic Systems Laboratories, Webster, TX).⁴² Intra-abdominal fat pads were removed under terminal isoflurane anesthesia, and the fluid content determined after heating of the samples in an oven at 70°C over 7 d.

Lithium Experiments

Mice were kept in standard rodent cages and fluid intake was determined before and during lithium treatment for 8 d. Lithium was given in the drinking water (initial concentration of LiCl 20.8 mM). In Scnn1a^loxlox mice, the Li concentration of drinking water was adjusted daily to compensate for the lithium-induced increase in fluid intake, such that lithium intake in all mice was kept between 4 and 8 μmol lithium/24 h per g body wt). Urine was spontaneously collected before and after 8 d of lithium treatment to measure urine osmolality (Vapro; Wescor, Salt Lake City, UT).

Electrophysiology

Patch-clamp electrophysiology was used to assess ENaC activity in isolated, split-open CCDs and the activity of Na⁺ conducting channels in freshly isolated primary mouse IMCD cultures. Both preparations have been described previously.^43–47 In brief, pathogen-free C57BL/6 mice of either gender (approximately 8 wk of age) were maintained for 10 d on a control diet (Harlan TEKLAD; TD.7001) or with this diet supplemented with pioglitazone (220 mg/kg¹⁷) or rosiglitazone (320 mg/kg¹⁶). In control experiments, we tested the effects of dietary Na⁺ on ENaC activity. Mice were maintained with control diet (0.32% [Na⁺]; TD.9712) and diets nominally Na⁺ free (<0.01 [Na⁺]; TD.90228) and enriched in Na⁺ (2.0% [Na⁺]; TD.92034) for 7 d.

Mice were killed by cervical dislocation and the kidneys immediately removed. Kidneys were cut into thin slices (<1 mm), and slices placed in ice-cold physiologic saline solution (pH 7.4). CDs were mechanically isolated from these slices by microdissection using watchmaker forceps under a stereomicroscope. Isolated CCDs were allowed to settle onto 5 × 5-mm cover glass–coated with poly-l-lysine. Cover glass containing CDs were placed within a perfusion chamber mounted on an inverted Nikon TE2000 and superfused with a physiologic saline solution buffered with HEPES (pH 7.4). CDs were split open with a sharpened micropipette controlled with a micromanipulator to gain access to the apical membrane. CDs were used within 1 to 2 h of isolation.

Primary cultures of IMCD cells were generated using a modified protocol as described previously.⁴⁴ In brief, kidneys were dissected from adult mice. The inner medulla was dissected and minced into 1-mm fragments, which were digested in a bicarbonate-free Krebs solution ([in mM] 145 NaCl, 10 HEPES, 5 KCl, 1 NaH₂PO₄, 2.5 CaCl₂, 1.8 MgSO₄, and 5 glucose [pH 7.3]) with 1 mg/ml collagenase type I (Worthington, Lakewood, NJ) and 0.1 mg/ml DNase I (Sigma) at 37°C for 15 to 20 min until the tissue was dispersed into single tubule fragments. Single tubule fragments were collected by centrifugation at 400 × g for 5 min in 10% albumin in PBS. The pellet was resuspended in medium and cultured in 12-well polyester membrane (0.4 μm pore) transwells (Corning, Lowell, MA). The cells were cultured in DMEM/F-12 (1:1; Invitrogen) with 15 mM HEPES, 100 U/ml penicillin, 10 U/ml streptomycin, 2 mM glutamine, 10 μg/ml insulin, 5.5 μg/ml transferrin, 6.7 ng/ml selenium, 10⁻⁷ M dexamethasone, 5 nM T3, 10 ng/ml mouse EGF, and 10% FBS at 37°C until confluent.

Cell-attached patches were made under voltage-clamp conditions using pipettes having resistances of 10 to 15 megaohms on the apical plasma membranes of principal cells in isolated, split-open CDs and primary mouse IMCD cultures using standard procedures.^43,48,49 Bath and pipette solutions were as follows (in mM): 155 NaCl, 1 CaCl₂, 2 MgCl₂, 5 glucose, and 10 HEPES (pH 7.4) and 140 LiCl, 2 MgCl₂, and 10 HEPES (pH 7.4), respectively. Single-channel current data from gigaohm seals were acquired (and subsequently analyzed) with an Axopatch 200B (Axon Instruments, Union City, CA) patch-clamp amplifier interfaced via a Digidata 1322A (Axon Instruments) to a PC running the pClamp 9.2 suite of software (Axon Instruments). Currents were low-pass filtered at 100 Hz with an eight-pole Bessel filter (Warner Instruments, Hamden, CT). Unitary current (i) and the number of ENaC in a patch, N, were determined from all-point amplitude histograms. Channel activity defined as NP_o was calculated using the following equation: NP_o = Σ(t₁ + 2t₂ + … nt_n), where t_n is the fractional open time spent at each of the observed current levels. P_o was calculated by normalizing NP_o for the number of channels observed within a patch. Only patches containing five channels or fewer were used to estimate P_o.

Statistical Analysis

All summarized data were reported as means ± SEM. Data were compared with the two-tailed t test. P ≤ 0.05 was considered statistically significant. For presentation, current data from some cell-attached patches were subsequently software filtered at 50 Hz, and slow baseline drifts were corrected.

DISCLOSURES

Studies using pioglitazone were supported by Takeda Pharmaceuticals America, Inc.