PKD2 Functions as an Epidermal Growth Factor-Activated Plasma Membrane Channel

Abstract

PKD2, or polycystin 2, the product of the gene mutated in type 2 autosomal dominant polycystic kidney disease, belongs to the transient receptor potential channel superfamily and has been shown to function as a nonselective cation channel in the plasma membrane. However, the mechanism of PKD2 activation remains elusive. We show that PKD2 overexpression increases epidermal growth factor (EGF)-induced inward currents in LLC-PK₁ kidney epithelial cells, while the knockdown of endogenous PKD2 by RNA interference or the expression of a pathogenic missense variant, PKD2-D511V, blunts the EGF-induced response. Pharmacological experiments indicate that the EGF-induced activation of PKD2 occurs independently of store depletion but requires the activity of phospholipase C (PLC) and phosphoinositide 3-kinase (PI3K). Pipette infusion of purified phosphatidylinositol-4,5-bisphosphate (PIP₂) suppresses the PKD2-mediated effect on EGF-induced conductance, while pipette infusion of phosphatidylinositol-3,4,5-trisphosphate (PIP₃) does not have any effect on this conductance. Overexpression of type Iα phosphatidylinositol-4-phosphate 5-kinase [PIP(5)Kα], which catalyzes the formation of PIP₂, suppresses EGF-induced currents. Biochemical experiments show that PKD2 physically interacts with PLC-γ2 and EGF receptor (EGFR) in transfected HEK293T cells and colocalizes with EGFR and PIP₂ in the primary cilium of LLC-PK₁ cells. We propose that plasma membrane PKD2 is under negative regulation by PIP₂. EGF may reduce the threshold of PKD2 activation by mechanical and other stimuli by releasing it from PIP₂-mediated inhibition.

ADPKD (autosomal dominant polycystic kidney disease) is a common systemic disease affecting multiple organs and cell types (12, 17). ADPKD affects 1 in 1,000 individuals, primarily by the development of large, fluid-filled renal cysts that ultimately may lead to kidney failure. ADPKD is caused by mutations in at least two separate genes, pkd1 and pkd2 (1, 5, 23, 49). Their protein products, called PKD1 and PKD2 (or polycystin 1 and 2, respectively), are large, membrane-associated proteins with putative roles in signal transduction and Ca²⁺ regulation (1, 5, 23, 49).

A large amount of PKD2 is present in the endoplasmic reticulum, where it functions as a Ca²⁺-activated intracellular Ca²⁺ release channel (25). A significantly smaller amount of PKD2 is present in the plasma membrane, where it functions as a nonselective cation channel (15, 18, 28). In addition to these two subcellular sites, PKD2 expression has been documented in the primary cilium of kidney epithelial cells, where it is believed to have an essential role in mediating Ca²⁺ entry in response to flow rate changes (34), suggesting that it may be part of a mechanosensing machinery residing in the primary cilium. While all these functions may very well represent physiological functions of PKD2, its biological role in the cilium may be more closely related to the pathophysiology of ADPKD, as several independent studies have shown that loss of function of other ciliary proteins often results in kidney cysts (7, 48). However, the exact function and mechanism of PKD2 activation in the cilium and/or other subcellular sites remain largely unknown.

Several of the mammalian transient receptor potential (TRP) channels, particularly members of the canonical (TRPC) and the vanilloid (TRPV) groups, function downstream of phospholipase C (PLC) activation (9, 32). Because PKD2 belongs to the TRP superfamily and is likely to be activated by similar mechanisms, we searched for mutations in components of known PLC-activating signal transduction pathways that have been reported to cause kidney cysts in mice. Targeted deletion of the epidermal growth factor receptor (EGFR) gene by homologous recombination in CD-1 mice resulted in cystic dilatation of collecting ducts (43), an area that is also affected by mutations in the PKD2 gene (50). Therefore, these previous results implied that there might be functional interaction between EGFR and PKD2 and prompted us to test whether PKD2 activity can be directly modulated by epidermal growth factor (EGF) in kidney epithelial cells.

EGF mediates its effects via the activation of the EGFR, a prototypical receptor tyrosine kinase (RTK) (24). Activated EGFR forms a binding site for PLC-γ isoforms (γ1 and γ2) which are recruited to the activated receptor to catalyze the formation of inositol-1,4,5-trisphosphate (IP₃) and diacylglycerol (DAG) from phosphatidylinositol-4,5-bisphosphate (PIP₂). Activated EGFR also activates phosphoinositide 3-kinase (PI3K) to phosphorylate PIP₂ to phosphatidylinositol-3,4,5-triphosphate (PIP₃). IP₃ is a short-lived second messenger that binds to the IP₃ receptor to induce a Ca²⁺ release transient mainly from the endoplasmic reticulum. IP₃-induced depletion of the internal Ca²⁺ stores activates the so-called store-operated Ca²⁺ channels (SOCs) to admit Ca²⁺ from the extracellular space to replenish the stores (3). While the biophysical properties of SOCs operating mainly in hematopoietic cells have been studied in detail (22, 52), their molecular identity remains unknown. Therefore, it is not surprising that the channels mediating the EGF conductance are also unknown. However, there is growing evidence that EGF-induced currents can be carried by channels that do not necessarily require store depletion for activation in certain cell types. Specifically, inhibition of EGFR's tyrosine kinase activity by tyrphostin A23 and A25 in guinea pig ventricular myocytes implicated the L-type voltage-gated Ca²⁺ channels as the channels mediating EGF-induced currents (36). In contrast, EGF-induced Ca²⁺ entry in A431 cells was not affected by nifedipine, a potent antagonist of voltage-gated Ca²⁺ channels, indicating that EGF-induced currents are not likely to be carried by voltage-activated Ca²⁺ channels in A431 cells (33). A recent study by Zhang et al. suggested that EGF-stimulated Ca²⁺ influx occurs via a mechanism distinct from store-operated Ca²⁺ entry in a human salivary cell line (51). However, our previous data demonstrated that EGF-induced channels possessed biophysical and pharmacological properties similar to those of SOCs but did not require depletion of internal stores for activation in cultured human mesangial cells (27, 29). Therefore, while it is expected that in most cell types EGF would activate SOCs through the PLC-γ pathway (3), EGF-induced conductance can be mediated by channels activated by modes other than store depletion. Examples of such regulation related to TRP channels include the EGF-mediated activation of TRPC5 (4) and of TRPC6 (21). Particularly relevant to our study is the first example, which shows that EGF induces the vesicular translocation of TRPC5 to the plasma membrane through a Rac1/type Iα phosphatidylinositol-4-phosphate 5-kinase [PIP(5)Kα]-dependent mechanism (4).

In the present study, we show that EGF activates PKD2 by releasing it from inhibition by PIP₂ through PLC-γ2-induced PIP₂ hydrolysis and by PI3K-induced conversion of PIP₂ to PIP₃. PKD2 activation by EGF is independent of store depletion. This regulation may be mediated by interactions between PKD2, PLC-γ2, and EGFR, as these proteins interact in transfected cells and colocalize with PIP₂ at the primary cilium of kidney epithelial cells, where PKD2 is expected to be activated by mechanical stimulation. Our data provide evidence for a novel mechanism of PKD2 regulation by EGF.

MATERIALS AND METHODS

Plasmids.

PKD2-D511V was constructed by standard site-directed mutagenesis using QuikChange (Stratagene). The cDNA for human EGFR was moved from pOT12-hEGFR (obtained from T. Carter) to pCDNA3. Expression plasmids of PLC-γ1 and -γ2 in pCMV-SPORT6 were obtained from Open Biosystems. All other constructs were described earlier (45).

Reagents and antibodies.

Purified brain PIP₂ and mouse monoclonal α-EGFR (GR-15 or GR-15L) were obtained from Calbiochem, while synthetic PIP₃ and monoclonal α-PIP2 (immunoglobulin M) were obtained from Echelon. Rabbit polyclonal antibodies against PLC-γ1, -γ2, and EGFR were obtained from Santa Cruz Biotechnology.

Cell culture and stable transfections.

LLC-PK₁ cells were cultured in M199 medium supplemented with 3% fetal bovine serum. Expression construct containing the m5 muscarinic acetylcholine receptor subtype cDNA (LLC-PK₁^Control), wild-type human PKD2 (LLC-PK₁^PKD2), human PKD2-D511V (LLC-PK₁^PKD2-D511V), or human hemagglutinin (HA)-PKD2 (LLC-PK₁^HA-PKD2) was transfected into LLC-PK₁ cells using Lipofectamine. Transfected cells were selected for stable integration using G418 (200 μg/ml). Individual clones were picked using cloning cylinders, expanded, and analyzed for PKD2 expression by immunoprecipitation with rabbit α-PKD2 followed by immunoblotting using chicken α-PKD2 antibody. LLC-PK₁^Control, LLC-PK₁^PKD2, LLC-PK₁^PKD2-D511V, and LLC-PK₁^HA-PKD2 represent single clones.

RNA interference.

To inactivate pig PKD2 in LLC-PK₁ cells, we used our previously described constructs (40). PKD2^KD2-2 targeted specifically porcine PKD2, while PKD2^KD2-3 targeted porcine and human PKD2. These constructs were made in our previously described RNA interference (RNAi) vector, pUB/H1/RNAi vector (30). As a control off-target sequence, we used pUB/RNAi/Ubc-H6 targeting the human Ubc-H6 enzyme. Inactivating constructs were transiently cotransfected with a CD8α expression plasmid into LLC-PK₁^PKD2 cells using Lipofectamine Plus reagent. CD8α⁺ cells were identified by a 10-min incubation with magnetic beads coupled to α-CD8α (DYNAL).

Electrophysiology.

The conventional whole-cell voltage-clamp configuration was used to measure transmembrane currents in single cells as described previously (30). Briefly, patch-clamp recordings were obtained from single cells at room temperature using a Warner PC-505B amplifier (Warner Instrument Corp., Hamden, CT) and pClamp 8 software (Axon Instrument, Foster City, CA). Glass pipettes (plain; Fisher Scientific, Pittsburgh, PA) with resistances of 5 to 8 MΩ were prepared with a pipette puller and polisher (PP-830 and MF-830, respectively; Narishige, Tokyo, Japan). After the whole-cell configuration was achieved, cell capacitance and series resistance were compensated before each recording period. From a holding potential of 0 mV, voltage steps were applied from −100 to 80 mV in 20-mV increments with 100-ms durations at 5-s intervals. Current traces were filtered at 1 kHz and analyzed offline with pClamp 8. Statistical analysis was employed with SigmaStat (Chicago, IL) software. Data were reported as means ± standard errors of the means. The Student t test was used for comparisons between groups. Differences were considered significant at P values of <0.05. The pipette solution contained (in mM): 135 cesium methane sulfonate, 8 NaCl, 1 MgCl₂, 0.3 Mg-ATP, 0.03 GTP, 10 HEPES, and 10 EGTA (pH 7.2). The standard external (bath) solution contained (in mM): 110 NaCl, 5 CsCl, 1 MgCl₂, 20 CaCl₂, 10 HEPES, and 10 glucose (pH 7.4).

Whole-cell current measurements in low (low K⁺) or high (high K⁺) extracellular K⁺ concentrations were obtained and analyzed as described above, with the following exceptions: from a holding potential of −60 mV, voltage steps were applied from −100 to 100 mV in 20-mV increments with 200-ms durations at 3-s intervals. The pipette solution contained (in mM): 100 K-aspartate, 30 KCl, 0.3 Mg-ATP, 10 HEPES, 10 EGTA, and 0.03 GTP (pH 7.2). The extracellular solution (ECS) in low K⁺ contained (in mM) 110 NaCl, 5 KCl, 1 MgCl₂, 20 CaCl₂, 10 HEPES, and 10 glucose (pH 7.4), and the ECS in high K⁺ contained (in mM) 130 KCl, 1 MgCl₂, 10 HEPES, 0.1 CaCl₂, and 5 glucose (pH 7.4).

Ratiometric Ca²⁺ measurements.

To determine intracellular Ca²⁺ concentration ([Ca²⁺]_i) in single cells (see Fig. 4B), LLC-PK₁^PKD2 cells were plated onto glass coverslips and loaded with 5 μM Fura-2/AM in ECS containing 140 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 1.8 mM CaCl₂, 10 mM glucose, and 15 mM HEPES, pH 7.4 {extracellular Ca2+ concentration ([Ca²⁺]_o), 1.8 mM } in the presence of 0.05% Pluronic F-127 for 1 h at room temperature. Cells were washed twice in ECS and incubated for 15 min in 37°C before intracellular imaging. To determine EGF-induced release transient, cells were incubated in a nominally Ca²⁺-free solution (same as ECS but 1.8 mM CaCl₂ was replaced by 0.5 mM EGTA [ECS-EGTA]; [Ca²⁺]_o, <1 nM) and stimulated with 100 nM EGF at the indicated time. Ca²⁺ influx in response to ECS-EGTA alone or to ECS-EGTA and EGF (100 nM) was determined by the readdition of ECS containing a 10 mM [Ca²⁺]_o. Individual cells were imaged with a charge-coupled-device camera (CoolSnap HQ; Photometrics) driven by Metafluor software (Universal Imaging). Intracellular Ca²⁺ concentration was expressed as a 340/380 ratio.

FIG. 4.

Functional contribution of native PKD2 to EGF-induced currents in LLC-PK₁ cells. (A) LLC-PK₁ cells were transiently cotransfected with wild-type myc-tagged human PKD2 (M-PKD2), pUB/RNAi/Ubc-H6, and green fluorescent protein (GFP) (lane 1); M-PKD2, RNAi construct PKD2^KD2-2, and GFP (lane 2); or M-PKD2, RNAi construct PKD2^KD2-3, and GFP (lane 3). Cell lysates were immunoblotted with rabbit α-myc (1:1,000; Santa Cruz Biotechnology) (upper panel) or α-GFP (1:1,000; Santa Cruz Biotechnology) (lower panel). (B) LLC-PK₁ cells were transiently transfected with pUB/RNAi/Ubc-H6 (lane 1), PKD2^KD2-2 (lane 2), or PKD2^KD2-3 (lane 3). Cells were lysed and endogenous PKD2 was immunoprecipitated with rabbit α-PKD2. Immunocomplexes were immunoblotted with chicken α-PKD2 (1:3,000) (upper panel). Lysates were immunoblotted with rabbit α-TRPC1 (1:500). Two isoforms of endogenous TRPC1, of 80 kDa and 65 kDa, are shown (lower panel). (C) Current-voltage (I-V) curves before and 3 min after EGF of LLC-PK₁^PKD2 cells transiently transfected with pUB/RNAi/Ubc-H6 (n = 8) *, P < 0.05. (D) curves before and 3 min after EGF of LLC-PK₁^PKD2 cells transiently transfected with PKD2^KD2-3 (n = 8). (E) I-V curves before and 3 min after EGF of LLC-PK₁^Control cells transiently transfected with PKD2^KD2-2 (n = 7). (F) I-V curves before and after EGF in LLC-PK1^PKD2-D511V cells; n = 6.

To determine intracellular Ca²⁺ concentration in cell populations (∼2 × 10⁶ cells) (see Fig. 5C), cells were harvested in phosphate-buffered saline containing 0.5 mM EDTA, washed with normal ECS, and loaded with 2 μM Indo-1/AM in the presence of 0.05% Pluronic F-127 (Molecular Probes) in ECS for 40 min at room temperature. After the 40-minute incubation, cells were washed three times with ECS-EGTA and resuspended in 2 ml of ECS-EGTA. Cells in ECS-EGTA were first incubated with 100 nM EGF at the indicated time, and then Ca²⁺ entry was determined by CaCl₂ readdition (10 mM CaCl₂) (see Fig. 5C). Ratiometric measurements representing free intracellular Ca²⁺ concentrations were obtained by a PTI QuantaMaster spectrofluorometer equipped with an excitation monochromator set at 350 nm and two emission monochromators set at 405 and 485 nm.

FIG. 5.

EGF-induced Ca²⁺ entry in LLC-PK₁^PKD2 cells. (A) Current-voltage (I-V) curves showing EGF-induced currents with and without 10 mM EGTA in the pipette solution. (B) Fura-2/AM fluorescent measurement of [Ca²⁺]_i (shown as 340/380 fluorescence ratio) in response to EGF using a Ca²⁺ readdition protocol in single LLC-PK1^PKD2 cells. Three (red) or four (black) LLC-PK₁^PKD2 cells were incubated in ECS-EGTA in the presence (red) or absence (black), respectively, of 100 nM EGF. A high concentration of extracellular Ca²⁺ (10 mM) was added to EGF-treated cells (red) 1 min later than it was added to control cells. Top and bottom bar graphs are diagrammatic representations of the Ca²⁺ readdition protocol followed in control (black) and EGF-treated (red) cells, respectively. (Inset) Summary data of Ca²⁺ entry in EGF-stimulated (+) (n = 20) or unstimulated (−) (n = 19) LLC-PK₁^PKD2 cells. *, P < 0.05. (C) Indo-1/AM fluorescence measurement of [Ca²⁺]_i (shown as 405/485 fluorescence ratio) in response to EGF using a Ca²⁺ readdition protocol in a population (∼2 × 10⁶) of LLC-PK₁^PKD2 cells.

Immunoblotting-immunoprecipitations in total cell lysates were done as described earlier (30, 45). To determine the cell surface expression of PKD2, we generated a stable line expressing the human PKD2 tagged with HA at its first extracellular loop (HA-PKD2). Live HA-PKD2-expressing cells (LLC-PK₁^HA-PKD2; 8 × 10⁶/150-mm dish) were incubated with 20 μg/ml of rabbit α-HA (Sigma) or our own rabbit α-PKD2 raised against the intracellular segment (679 to 742 amino acid residues) of human PKD2 in normal ECS containing 0.5% bovine serum albumin for 1 h at 4°C. Unbound antibodies were washed five times with ECS, and cells were lysed in standard immunoprecipitation solution. Bound antibodies were captured with protein A, and HA-PKD2 was detected by mouse α-HA (at 5 μg/ml; Boehringer) by Western analysis. To determine the total amount of HA-PKD2 in 8 × 10⁶ cells, cells were lysed and HA-PKD2 was directly immunoprecipitated using 4 μg of each antibody in 1 ml of total cell lysates. Immunocomplexes were probed with mouse α-HA.

Immunofluorescence.

LLC-PK₁ cells were grown on glass coverslips for 7 days to allow the formation of cilia. Cells were fixed with 3.7% formaldehyde in cytoskeletal stabilizing buffer {PHEM buffer; 60 mM PIPES [piperazine-N,N′-bis(2-ethanesulfonic acid)], 25 mM HEPES, 10 mM EGTA, 2 mM MgCl₂, adjusted to pH 6.9 with KOH} at room temperature for 15 min, which was followed by three rinses in PHEM buffer. The cells were extracted with 0.1% Triton X-100 in PHEM buffer for 90 seconds. Blocking was done at 4°C overnight or at room temperature for 1 h in PHEM buffer containing 0.1% Triton X-100 and 10% normal goat serum. Primary antibodies were diluted in PHEM buffer containing 0.1% Triton X-100 and 10% normal goat serum for overnight incubation. Rabbit affinity-purified antibody to PKD2 was used at 1:500 (or 0.5 μg/ml), mouse antibody to PIP₂ was used at 1:500 (or 2 μg/ml), and mouse antibody to EGFR was used at 1:500 (or 0.2 μg/ml). Coverslips were mounted with ProLong (Molecular Probes). Antibody staining was visualized with laser excitation from Ar-488 and Kr-568 lasers and viewed on a Leica TCS NT microscope. Images were processed with Adobe Photoshop software.

RESULTS

EGF-induced currents are enhanced in LLC-PK₁^PKD2 cells.

We generated several stable lines of wild-type nontagged PKD2 (PKD2; LLC-PK₁^PKD2 cells), D511V PKD2 mutant (PKD2-D511V; LLC-PK₁^PKD2-D511V cells), and wild-type HA-tagged PKD2 (HA-PKD2, LLC-PK₁^HA-PKD2 cells) in LLC-PK₁ cells. Figure 1A shows expression levels of endogenous PKD2 (lanes 1 and 4), transfected PKD2 (lanes 2 and 5), and PKD2-D511V (lanes 3 and 6). To test whether transfected PKD2 can reach the plasma membrane, we engineered the HA tag in the first extracellular loop of HA-PKD2 and tested whether it can be recognized by α-HA added in the extracellular space. As a control antibody, we used our own α-PKD2 raised against the intracellular C-terminal tail of human PKD2, which should not detect PKD2 in nonpermeabilized cells. Incubation of live cells with α-HA or α-PKD2 followed by washing out unbound antibodies, immunocapture by protein A, and detection by Western blotting revealed that less than 5% of total transfected HA-PKD2 reached the plasma membrane (Fig. 1B). By analogy, transfected wild-type nontagged PKD2 should also be present in the plasma membrane at a similar ratio. Thus, we proceeded with functional studies using the LLC-PK₁^PKD2 cells.

FIG. 1.

Expression levels of endogenous and transfected PKD2 in LLC-PK₁ cells. (A) Expression levels of wild-type PKD2 in LLC-PK₁^Control (lanes 1 and 4) and LLC-PK₁^PKD2 (lanes 2 and 5) cells and PKD2-D511V in LLC-PK₁^PKD2-D511V cells (lanes 3 and 6) determined by immunoprecipitation using rabbit α-PKD2 followed by immunoblotting using chicken α-PKD2 (1:3,000). Ten μl (lanes 1 to 3) or 30 μl (lanes 4 to 6) of immunocomplexes was used to allow accurate comparisons between endogenous and transfected PKD2. (B) Cell surface expression of HA-PKD2 in LLC-PK₁^HA-PKD2 cells. Plasma membrane HA-PKD2 (Ext, external) was immunoprecipitated with α-HA (lane 4) or α-PKD2 (lane 5) from live cells. Fifty percent (lane 1), 20% (lane 2), and 5% (lane 3) of total α-HA immunocomplexes are also shown.

Previous work had demonstrated that LLC-PK₁ cells express endogenous PKD2 and functional EGFR (16, 19, 25, 28, 42). Initial single channel recordings in cell-attached patches of LLC-PK₁^PKD2 cells indicated the existence of an EGF-activated channel with a conductance in K⁺ higher than that in Na⁺ or in Ca²⁺ (see the supplemental material). These data prompted us to measure basal and EGF-induced whole-cell K⁺ currents in LLC-PK₁^PKD2 cells. Furthermore, we asked whether cell dialysis with our PKD2 antibody could modulate basal and EGF-induced K⁺ currents. A rabbit antibody raised against the α subunit of human CD8 (α-CD8α) was used as a control antibody. In these experiments, whole-cell recordings were initiated 5 min following break-in to allow sufficient time for antibody diffusion. Switching the extracellular solution from low K⁺ (5 mM KCl) to high K⁺ (130 mM KCl) resulted in an increase in inward holding current at a membrane potential (V_m) of −60 mV as would be expected for a background K⁺ conductance. Voltage steps from −100 to 100 mV from a holding potential (V_h) of −60 mV resulted in large inward and outward currents, indicating the activation of a K⁺ conductance in cells dialyzed with α-CD8α (Fig. 2A, B, D, E, G, and H). Interestingly, cell dialysis with α-PKD2 reduced the holding current at −60 mV in high K⁺ as well as at potentials between −100 and −40 mV (Fig. 2H), suggesting the contribution of PKD2 to basal K⁺ conductance. At all tested potentials, bath application of EGF augmented K⁺ currents (Fig. 2C, F, and I), which were completely blocked by cell dialysis with α-PKD2 (Fig. 2I). These data clearly showed that PKD2 was directly involved in the formation of EGF-induced K⁺ currents and were consistent with the property of purified PKD2 of mediating K⁺ currents (14). They also revealed that EGF activated PKD2 to mediate both inward and outward currents that were best resolved when K⁺ was used as the charge carrier.

FIG. 2.

PKD2-mediated EGF-induced K⁺ currents. (A to F) Whole-cell current measurements obtained by a voltage step protocol in LLC-PK₁^PKD2 cells dialyzed with 200 ng/ml of α-CD8α (A to C) or α-PKD2 (D to F). Step currents were obtained at low K⁺ (5 mM KCl) (A and D), 1 min after switching to high K⁺ (130 mM KCl) (B and E), or 3 min after the addition of 100 nM EGF to the extracellular solution in high K⁺ (C and F). Pulses were applied from a holding potential of −60 mV and covered a voltage range from −100 mV to 100 mV with 20-mV increments. Each pulse had a duration of 200 ms, and currents were measured at the end of the pulse. (G to I) Current-voltage (I-V) curves of LLC-PK₁^PKD2 cells dialyzed with 200 ng/ml of α-CD8α (n = 7) or α-PKD2 (n = 5) at low K⁺ (G), high K⁺ (H), or high K⁺ plus EGF (I). *, P < 0.05.

Next, we wished to determine whether EGF could activate PKD2-mediated currents using Na⁺ and Ca²⁺ as the charge carriers, which would carry EGF-induced inward currents under physiological conditions. To block K⁺ currents, we included Cs⁺ in the bath and pipette solution. Continuous recordings at a V_h of −100 mV showed that currents were induced shortly after the administration of EGF (100 nM), reached maximum levels within 3 min (Fig. 3A and B), and were inhibited by 20 μM La³⁺ (Fig. 3A and B) in both cell types. EGF-induced currents in LLC-PK₁^PKD2 cells were slightly greater in amplitude and noise level.

FIG. 3.

EGF-induced whole-cell current in LLC-PK₁^Control and LLC-PK₁^PKD2 cells. (A and B) Time course experiments with EGF-induced current in LLC-PK₁^Control and LLC-PK₁^PKD2 cells. La³⁺ (20 μM) was added about 4.5 min following the addition of EGF (100 nM). (C and D) Net EGF-induced currents in LLC-PK₁^Control and LLC-PK₁^PKD2 cells obtained by a voltage step protocol. Background currents were subtracted from EGF-induced currents offline. (E and F) Current-voltage (I-V) relation curves generated from a voltage step protocol showing basal (Pre-EGF) and EGF-induced (Post-EGF) currents 3 min after the addition of 100 nM EGF to the extracellular solution in LLC-PK1^Control (n = 15; membrane capacitance [Cm] = 16.6 ± 1.37 pF) (E) or LLC-PK₁^PKD2 (n = 9; Cm = 16.5 ± 1.0 pF) (F) cells. *, P < 0.05. Current measurements were obtained at 50 ms of each pulse. Membrane currents were expressed as current densities (pA/pF).

Using a voltage step protocol before and 3 min after the administration of 100 nM EGF, we found that EGF significantly increased the inward currents in both cell types. Specifically, EGF increased basal inward currents from −3.7 ± 0.32 pA/pF to −5.7 ± 0.48 pA/pF (n = 15 and P < 0.05 for LLC-PK₁^Control cells) (Fig. 3C and E) and from −3.4 ± 0.4 pA/pF to −7.3 ± 0.9 pA/pF (n = 9 and P < 0.05 for LLC-PK₁^PKD2 cells) (Fig. 3D and F) at −100 mV. Therefore, EGF-induced inward currents were almost doubled in LLC-PK₁^PKD2 (−3.8 ± 0.51 pA/pF; n = 9) (Fig. 3E) compared to those in LLC-PK₁^Control (−1.9 ± 0.18 pA/pF; n = 15) (Fig. 3F). To confirm that this effect was not due to clonal variability, we transiently transfected PKD2 into parental LLC-PK₁ cells. EGF induced a similar increase in inward currents (from −3.7 ± 0.47 pA/pF to −6.9 ± 0.94 pA/pF; n = 3). Inward currents were most likely carried by Na⁺ and Ca²⁺ and not by Cl⁻, as the Cl⁻ concentration was much higher in the extracellular solution (157 mM) than in the pipette solution (10 mM). We noticed that while outward currents were increased in response to EGF, they did not reach significant levels in either cell type (Fig. 3E and F). This can be explained by the fact that outward currents would normally carry K⁺, which has been replaced by Cs⁺ in our pipette solution. In sum, these experiments showed that EGF induced inward currents carried by Na⁺ and Ca²⁺ in LLC-PK₁ cells and PKD2 enhanced these currents. The amplitude of EGF-induced whole-cell Na⁺/Ca²⁺ currents at −100 mV in LLC-PK₁^PKD2 cells (Fig. 3F) was ∼10-fold lower than the amplitude of K⁺ currents in the same cells (Fig. 2I), further supporting the idea that EGF-activated currents were mediated through PKD2, which has been shown to display conductance in K⁺ higher than that in Na⁺ or Ca²⁺ (14, 15, 28, 47).

Endogenous PKD2 is required for EGF-induced currents in LLC-PK₁ cells.

To determine whether native PKD2 contributed to EGF-activated currents in control and PKD2-overexpressing cells, we knocked down endogenous porcine or both porcine and transfected human PKD2 in LLC-PK₁^Control or LLC-PK₁^PKD2 cells, respectively. Construct PKD2^KD2-3 targeted a common region in porcine and human PKD2, while PKD2^KD2-2 was designed to target specifically porcine PKD2. Both of these constructs were described earlier and were shown by immunofluorescence staining to silence porcine PKD2 in LLC-PK₁ cells (40). Western blot analysis of LLC-PK₁ lysates transiently cotransfected with an expression plasmid of human myc-tagged PKD2 (M-PKD2) and each of the inactivating constructs showed that PKD2^KD2-3 effectively knocked down human PKD2 (Fig. 4A, lane 3), while PKD2^KD2-2 did not affect M-PKD2 expression (Fig. 4A, lane 2). This experiment verified that RNAi worked in LLC-PK₁ cells. The silencing effect of these constructs on endogenous porcine PKD2 in transiently transfected LLC-PK₁ cells was also determined by Western analysis (Fig. 4B, upper panel). However, no silencing effect was seen in either of the two isoforms (80 and 65 kDa) of endogenous pig TRPC1, demonstrating the specificity of our constructs for pig PKD2 (Fig. 4B, lower panel). However, it should be noted that silencing of endogenous PKD2 by transient transfection was evident only at transfection efficiencies higher than 50%.

Transient transfection of LLC-PK₁^PKD2 cells with an off-target RNAi construct, pUB/RNAi/Ubc-H6, did not significantly affect basal or EGF-induced currents (Fig. 4C). Silencing endogenous (porcine) and transfected (human) PKD2 by PKD2^KD2-3 (porcine and human) completely eliminated the EGF-induced currents in LLC-PK₁^PKD2 cells (Fig. 4D). Similar to the effect of PKD2^KD2-3 construct on LLC-PK₁^PKD2 cells, PKD2^KD2-2 (porcine only) transfection into LLC-PK₁^Control cells resulted in the elimination of EGF-induced currents (Fig. 4E), demonstrating that either endogenous or transfected PKD2 was required for EGF-induced conductance.

A pathogenic missense variant, PKD2-D511V, functions as a dominant negative allele.

To provide additional evidence that PKD2 mediated EGF-induced currents and to also test whether PKD2 activity, rather than a scaffolding role for PKD2, was essential for EGF-induced conductance, LLC-PK₁ cells were stably transfected with PKD2-D511V (LLC-PK₁^PKD2-D511V) (Fig. 1A, lanes 3 and 6) and were tested for EGF-induced conductance. Because PKD2-D511V was a pathogenic mutant (39) and maintained the homodimerization domain of PKD2 (46), we reasoned that it should function as a dominant negative allele if endogenous PKD2 mediated EGF-induced currents. Consistent with our prediction, Fig. 4F shows that EGF failed to induce a significant response in these cells. In complete agreement with RNAi experiments (Fig. 4D and E), PKD2-D511V suppressed EGF-induced inward currents (Fig. 4F). Therefore, these data suggested that PKD2 channel activity was required for EGF-induced ionic responses in LLC-PK₁ cells. It should also be noted that knockdown of endogenous PKD2 or overexpression of D511V resulted in a decrease in outward currents (Fig. 3E and 4C to F), indicating that PKD2 activity may be required for basal outward currents.

EGF activates PKD2 independently of store depletion.

It has been reported that PKD2 was positively regulated by intracellular Ca²⁺ (15, 25, 47). We found that EGF-induced currents were unaffected by the inclusion of 10 mM EGTA in the pipette solution (Fig. 5A). To further investigate whether PKD2 was insensitive to Ca²⁺ or whether EGF simply did not stimulate significant Ca²⁺ release from the internal stores, we assessed [Ca²⁺]_i in response to EGF by using ratiometric intracellular Ca²⁺ measurements for single cells (Fig. 5B) and for a population of cells (Fig. 5C). Both assays revealed that EGF failed to elicit a detectable Ca²⁺ release transient when Ca²⁺ was omitted from the extracellular solution in LLC-PK₁^PKD2 cells (Fig. 5B and C), but it did result in higher [Ca²⁺]_i following Ca²⁺ readdition (Fig. 5B), which may reflect increased influx, Ca²⁺-induced Ca²⁺ release, and/or reduced Ca²⁺ sequestration. These results suggested that EGF-induced enhancement of whole-cell currents in LLC-PK₁^PKD2 cells was unlikely to involve the traditional Ca²⁺-release-activated Ca²⁺ pathway. This statement was also supported by the failure of xestospongin C (10 μM), an inhibitor of IP₃ receptors, to suppress EGF-induced currents in LLC-PK₁^PKD2 cells (data not shown). The lack of significant EGF-stimulated Ca²⁺ release from the internal stores has been shown for LLC-PK₁ and other cell types (13, 20, 27, 31, 37).

EGF induces PKD2 activity by reducing PIP₂.

To understand the mechanism by which EGF induced PKD2 activity, we first tested whether EGFR tyrosine kinase activity was required for PKD2 activation. As shown in Fig. 6A, in the presence of 10 μM tyrphostin AG1478, a specific inhibitor of EGFR tyrosine kinase activity, EGF was unable to stimulate PKD2 channel activity, while tyrphostin AG1478 itself did not affect basal currents. A similar suppression was seen even at 100 nM AG1478 (from −4.5 ± 0.45 pA/pF to −5.3 ± 0.54 pA/pF at −100 mV; n = 8; P > 0.05). In the presence of 100 μM tyrphostin A1, an agent that is structurally similar to tyrphostin AG1478 but is an inactive inhibitor of RTK (used as a negative control for tyrphostin AG1478), EGF significantly induced PKD2 channel activity. These results suggest that RTK activity is required for EGF-induced channel activity but not for the basal activity of PKD2.

FIG. 6.

Pharmacologic characterization of EGF-induced current. (A) Summary data showing the effect of 100 μM tyrphostin A1 (n = 5) and 10 μM AG1478 (n = 8) on whole-cell current densities at a −100-mV holding potential in the presence or absence of EGF in LLC-PK₁^PKD2 cells; *, P < 0.05. (B) Effect of U73122 (1 μM) (n = 9) and U73433 (10 μM) (n = 7) on basal and EGF-induced currents at a −100-mV holding potential in LLC-PK₁^PKD2 cells; *, P < 0.05. (C) Effect of 100 μM OAG on basal and EGF-induced currents in LLC-PK₁^Control (n = 4) and LLC-PK₁^PKD2 (n = 4) cells at a −100-mV holding potential; *, P < 0.05. (D) Effect of 10 nM wortmannin (WMN) on basal and EGF-induced currents in LLC-PK₁^PKD2 cells (n = 8). (E) Effect of pipette infusion of 8 μM purified brain PIP₂ on basal and EGF-induced whole-cell current densities in LLC-PK₁^PKD2 cells (n = 6). (F) Effect of pipette infusion of 10 μM purified PIP₃ on basal and EGF-induced whole-cell current densities in LLC-PK₁^PKD2 cells (n = 6); *, P < 0.05. (G) AVP-induced currents in LLC-PK₁^Control (n = 8), U73122 (1 μM)-treated LLC-PK₁^Control (n = 4), and LLC-PK₁^PKD2 (n = 7) cells at a −100-mV holding potential; *, P < 0.05 between basal and AVP-treated cells. (H) Effect of AVP on whole-cell current densities in LLC-PK₁^PKD2 cells in the absence (n = 7) or presence (n = 4) of 8 μM purified brain PIP₂ in the recording electrode; *, P < 0.05. (I) Effect of transiently transfected PIP(5)Kα in LLC-PK₁^PKD2 cells. Capacitance-normalized net EGF-induced current at −100 mV in untransfected (−) or PIP(5)Kα-transfected (+) cells is expressed as change in current density (ΔIpA/pF); n = 9 or n = 11, respectively. *, P < 0.05.

Next, we determined whether PLC activity was required for EGF-induced activation of PKD2. LLC-PK₁^PKD2 cells were incubated with U73122, a widely used inhibitor of PLC isoforms, or U73433, an agent structurally similar to U73122 but lacking an inhibitory effect on PLCs. EGF-induced currents were blocked by U73122 but not by U73433 (Fig. 6B), suggesting that PLC activity was essential for EGF-induced activation of PKD2. PLC catalyzes the formation of IP₃ and DAG from PIP₂. To test whether DAG could account for the activation of PKD2 by EGF, LLC-PK₁^Control and LLC-PK₁^PKD2 cells were stimulated with 100 μM OAG (1-oleoyl-2-acetyl-sn-glycerol), a cell permeant analog of DAG, in the absence of EGF. OAG induced similar responses in both cell types (Fig. 6C), indicating that the OAG-induced current was unaffected by the overexpression of PKD2. Next, we determined whether PI3K had an effect on EGF-induced activation of PKD2 by preincubating LLC-PK₁^PKD2 cells with 10 nM wortmannin for 15 min before the addition of EGF. Wortmannin suppressed EGF-induced currents by almost 80% (from −3.8 pA/pF in untreated cells [Fig. 3F] to −0.67 pA/pF in wortmannin-treated cells [Fig. 6D]), suggesting that PI3K contributed to the EGF-induced activation of PKD2. We also noted that wortmannin reduced basal currents from −3.4 ± 0.6 pA/pF to −1.9 ± 0.45 pA/pF (Fig. 6D), suggesting that it might have inhibited the trafficking of PKD2 and possibly other channels to the plasma membrane. We reasoned that if PLC and PI3K activities were crucial for the activation of PKD2 by EGF, either the reduction of PIP₂ or the production of PIP₃ may be responsible for the activation mechanism. To test the first possibility, 8 μM purified brain PIP₂ was included in the recording electrode, and EGF-induced currents were determined. In the presence of PIP₂, EGF did not induce a significant increase in the amplitude of inward current (from −2.7 ± 0.18 pA/pF to −3.7 ± 0.29 pA/pF; P > 0.05 [Fig. 6E]). In contrast, in cells dialyzed with 10 μM purified PIP₃, EGF did induce a significant increase in current amplitude (from −2.6 ± 0.11 pA/pF to −5.0 ± 0.3 pA/pF; P < 0.05 [Fig. 6F]). Pipette infusion of PIP₃ did not augment basal or EGF-induced response in LLC-PK₁^PKD2 cells (Fig. 6F). These results suggested that reduction of PIP₂ by either hydrolysis or conversion to PIP₃, rather than increased production of PIP₃, might have been predominantly responsible for the activation of PKD2 by EGF. To further confirm the specificity of the inhibitory effect of PIP₂ on EGF-induced current, we showed that PIP₂ did not have an effect on arginine-vasopressin (AVP)-induced current, which was not mediated by the PLC pathway in these cells (38) (Fig. 6G and H). To provide additional evidence that PIP₂ functions as an inhibitor of PKD2, we transiently transfected PIP(5)Kα in LLC-PK₁^PKD2 cells and measured EGF-induced currents. PIP(5)Kα has been shown to increase PIP₂ production in permeabilized platelets, and its activity can be further augmented by activated Rac1 (44). Moreover, the PIP(5)Kα-induced production of PIP₂ resulted in increased cell surface expression of TRPC5 in response to EGFR activation (4). Our results showed that, in contrast to the effect of PIP(5)Kα on TRPC5, PIP(5)Kα reduced net EGF-induced currents by almost 85% (from −3.8 ± 0.5 pA/pF to −0.518 ± 0.09 pA/pF) (Fig. 6I), supporting previous results showing that PIP₂ had an inhibitory effect on PKD2 (Fig. 6E). In sum, these molecular biological and pharmacological data indicated that EGF-induced reduction of PIP₂ through the activation of PLC-γ isoforms and PI3K may account for the activation of PKD2 by EGF.

PKD2 associates with PLC-γ2 and EGFR in vitro.

We reasoned that if EGFR signals the activation of PLC-γ to release PKD2 from PIP₂-mediated inhibition, PKD2, EGFR, and PLC-γ isoforms may form a multimeric complex. Similar associations have been demonstrated between TRPC3 and TrkB (26) and between TRPV1 and TrkA (8). Coimmunoprecipitation experiments revealed that PLC-γ2, but not -γ1, was able to associate with the shortest truncation mutant of PKD2 containing the N-terminal cytosolic region (Fig. 7A), suggesting that a physical interaction between PLC-γ2 and the N-terminal cytosolic region of PKD2 could be possible in vivo. We also tested whether PKD2 could interact with EGFR. Wild-type PKD2 and the shortest [HA-PKD2(1-379)] or longest [HA-PKD2(1-871)] of the pathogenic PKD2 mutants were cotransfected with human EGFR in the presence or absence of PLC-γ2 in HEK293T cells, and the association between PKD2 and EGFR was determined by coimmunoprecipitation experiments. Interestingly, while wild-type PKD2 associated with EGFR, HA-PKD2(1-379) failed to interact, and HA-PKD2(1-871) displayed very weak interaction, if any (Fig. 7C). These data suggested that PKD2-EGFR association may be directly mediated by residues within the 97 most C-terminal residues of PKD2 or that these residues may be required for the stabilization of the interaction mediated by a domain located between residues 379 and 871. Consistent with a possible role for PLC-γ2 in EGF-mediated regulation of PKD2, LLC-PK₁ cells expressed endogenous PLC-γ2 (Fig. 7D), while PLC-γ1 was undetectable (data not shown).

FIG. 7.

Association of PKD2 with PLC-γ2 and EGFR. (A) HEK293T cells were transfected with PLC-γ2 alone (lane 1), PLC-γ2 and HA-PKD2(1-379) (lane 2), PLC-γ2 and HA-PKD2(1-702) (lane 3), PLC-γ2 and HA-PKD2(1-745) (lane 4), PLC-γ2 and HA-PKD2(1-871) (lane 5), or PLC-γ2 and wild-type HA-PKD2 (lane 6). HA-tagged proteins were immunoprecipitated with mouse monoclonal α-HA, and immunocomplexes were immunoblotted with rabbit α-PLC-γ2 at 0.2 μg/ml. The second panel from the top indicates the PLC-γ2 input. The third panel from the top shows the lack of association between PKD2 and PLC-γ1, in cells transfected with HA-tagged forms of PKD2 and PLC-γ1 as described for PLC-γ2. The bottom panel indicates the PLC-γ1 input. (B) Diagrammatic representation of wild-type and mutant PKD2 constructs showing association with PLC-γ2. (C) HEK293T cells were transfected with EGFR alone (lane 1); EGFR and wild-type HA-PKD2 (lane 2); EGFR, wild-type HA-PKD2, and PLC-γ2 (lane 3); EGFR and HA-PKD2(1-379) (lane 4); EGFR, HA-PKD2(1-379), and PLC-γ2 (lane 5); EGFR and HA-PKD2(1-871) (lane 6); or EGFR, HA-PKD2(1-871), and PLC-γ2 (lane 7). HA-tagged proteins were immunoprecipitated with mouse monoclonal α-HA, and immunocomplexes were immunoblotted with rabbit α-EGFR at a ratio of 1:1,000. Lower panel indicates EGFR input. (D) Expression of endogenous PLC-γ2 in LLC-PK₁ cells. Lysates of HEK293T cells transfected with PLC-γ2 (positive control, lane 1), crude lysates of LLC-PK₁ cells (lane 2), or immunoprecipitated PLC-γ2 from LLC-PK₁ lysates by using rabbit α-PLC-γ2 (lane 3) were blotted with rabbit α-PLC-γ2.

PKD2, PIP₂, and EGFR colocalize in the primary cilium of LLC-PK₁ cells.

To determine whether the EGF-induced regulation of PKD2 could be operative in the cilium, we determined the expression of native PKD2, EGFR, and PIP₂ in the primary cilium of LLC-PK₁ cells. Figure 8 shows that PKD2 colocalizes with PIP₂ (Fig. 8D to F) and EGFR (Fig. 8G to I) in this organelle, supporting the idea that cilium-expressed PKD2 might be under negative regulation by PIP₂, and inhibition of PKD2 channel activity could be reversed in the presence of EGF.

FIG. 8.

Expression of EGFR, PKD2, and PIP₂ in the primary cilium of LLC-PK₁ cells. Seven-day confluent cultures of LLC-PK₁ cells were stained with a monoclonal antibody (6-11B-1 clone; Sigma) against acetylated α-tubulin (1:2,000) (A), mouse monoclonal antibody (GR-15; Calbiochem) to EGFR (1:500 or 0.2 μg/ml) (B), or mouse monoclonal antibody to PIP₂ (Echelon) (1:500 or 2 μg/ml) (C). Each inset shows antibody staining of a single cilium. Colocalization of PKD2 (0.5 μg/ml) and PIP₂ (D to F) or EGFR (G to I) in cilia of LLC-PK₁ cells. LLC-PK₁ cells were stained with mouse monoclonal α-PIP₂ (2 μg/ml) and rabbit polyclonal α-PKD2 (0.5 μg/ml) or mouse monoclonal α-EGFR (0.2 μg/ml). Control cells stained with 0.5 μg/ml of rabbit immunoglobulin G (IgG) (K) and counterstained with ToPro3 (J). Primary rabbit or mouse antibodies were detected with goat α-rabbit Alexa 488- or α-mouse Alexa 566-conjugated secondary antibodies, respectively. Images were collected with a Leica TCS NT confocal microscope, and images were analyzed with Adobe Photoshop.

DISCUSSION

Our study shows that PKD2 enhances EGF-induced conductance and that native PKD2 activity is required for this conductance. Purified PIP₂, inhibition of PLC or PI3K, or transfection with PIP(5)Kα suppresses EGF-activated currents. In addition, PKD2 associates with PLC-γ2 and EGFR in transfected cells and colocalizes with PIP₂ and EGFR in the primary cilium of LLC-PK₁ cells. Our study suggests that PKD2 is under negative regulation by PIP₂, which can be reversed by EGFR activation. The implication is that EGF may sensitize PKD2 to mechanical or other stimuli by releasing it from PIP₂-mediated inhibition.

Single-channel measurements in purified PKD2 reconstituted in lipid bilayers (14, 15, 25) or in cell-attached patches of mIMCD3 cells (28) revealed that PKD2 showed a conductance higher in K⁺ than in Na⁺ or Ca²⁺. Our single-channel data in cell-attached patches of PKD2-transfected LLC-PK₁ cells showed the existence of a highly conductive K⁺ channel which was activated by EGF (see the supplemental material). Finally, whole-cell current measurements in high K⁺ showed that EGF-induced currents were increased by about 10-fold when K⁺ was used as the charge carrier over currents carried by Na⁺ and Ca²⁺. These large EGF-induced K⁺ currents were completely suppressed by a PKD2 antibody applied to the cytoplasm through the recording pipette. Taking into account previous studies of purified PKD2 reconstituted in lipid bilayers showing high K⁺ permeability (14, 15) and the lack of specific PKD2 blockers, we are prompted to conclude that EGF-activated currents are mediated through PKD2 in transfected cells. In support of this idea, it should be noted that our antibody targeted a cytoplasmic region of 63 residues immediately following the sixth transmembrane segment, which has been shown to be directly involved in the permeation properties of other TRP channels (35).

Under physiological conditions, Na⁺ and Ca²⁺ would be normally carried by inward currents. However, when we used Na⁺ and Ca²⁺ as the charge carriers, we found that EGF induced an approximately twofold (from 1.9 to 3.8 pA/pF)-higher response in cells expressing PKD2 than in mock-transfected cells. This increase did not directly correlate with the increase in expression levels of PKD2 following stable transfection (>10-fold), raising the issue of whether it represented a typical response obtained from an overexpression system where increases are usually expected to be >10-fold. This modest increase seen in our experiments could be explained by the limited expression of PKD2 in the plasma membrane following stable transfection, as biochemical experiments showed that less than 5% of total transfected PKD2 reached the plasma membrane. The reasons for the limited expression of PKD2 in the plasma membrane are not known. It has been shown that PKD2 requires PKD1 for targeting to the plasma membrane (18). Therefore, it is very possible that endogenous expression of PKD1 may be the rate-limiting factor for the plasma membrane expression of PKD2 in LLC-PK₁ cells.

Using a combination of complementary loss- and gain-of-function approaches, we show that PKD2 activity is regulated by EGF. This was most directly supported by the finding that PKD2 overexpression increased the amplitude of inward currents in response to EGF compared to control-transfected cells. In contrast, a pathogenic point mutant harboring a D511V substitution in the third transmembrane segment not only was unable to increase EGF-induced currents but also suppressed endogenous EGF-induced currents to levels seen when cells were transfected with a PKD2-specific RNAi construct. These results prompted us to conclude that PKD2 not only is sufficient to enhance but also is required for EGF-induced conductance in LLC-PK₁ cells.

To understand the mechanism by which EGF activated PKD2, we employed a series of molecular biological and pharmacological experiments. We first examined whether EGF activated through the activation of Ca²⁺ release from the internal stores. The lack of a measurable Ca²⁺ release transient supported by the lack of xestospongin C and intracellular EGTA effects on EGF-induced whole-cell currents simply indicates that EGF might not be able to induce production of IP₃ to levels adequate to drive a significant Ca²⁺ release transient. Alternatively, spatial constraints may prevent EGF-induced IP₃ from efficiently activating Ca²⁺ release transients. Regardless of the mechanism, while PKD2 can function very well as an intracellular channel in response to agents that provoke a Ca²⁺ release transient, PKD2 appeared to function in our system as an EGF-activated channel independently of store depletion.

Pharmacological studies showed that the inhibitions of PLCs and PI3K by U-73122 and wortmannin, respectively, suppressed EGF-induced currents in PKD2-overexpressing cells. In addition, cell dialysis with purified PIP₂ suppressed EGF-induced currents, while cell dialysis with PIP₃ did not augment basal or EGF-induced activity. These data prompted us to favor the idea that the reduction of PIP₂ secondary to PLCs and/or PI3K activation may be responsible for PKD2 activation. It has been reported that PI3K induces the phosphorylation, translocation, and activation of PLC-γ1, but not those of -γ2, in response to antigen stimulation in RBL-2H3 cells (2). Therefore, the wortmannin sensitivity might have been due to a block in PLC-γ1 activation rather than to the failure to convert PIP₂ to PIP₃. Because (i) pipette infusion of PIP₃ did not enhance PKD2 activity and (ii) PLC-γ1 was undetectable in LLC-PK₁ cells and even if it were to be present in small amounts it would be unlikely to associate with PKD2, we favor the idea that wortmannin blocked EGF-induced currents by reducing the size of the PIP₂ pool. However, we cannot formally exclude the possibility that small amounts of phosphatidylinositol-3,4-bisphosphate produced by PI3K could activate PKD2 currents.

It was recently shown that EGF induced the translocation of TRPC5 from vesicles residing in the cytoplasm to the plasma membrane (4). This translocation was dependent on Rac1/PIP(5)Kα activity. It was proposed that increased PIP₂ production in response to EGF-induced activation of PIP(5)Kα was responsible for the vesicular translocation of TRPC5. To test whether a similar mechanism may underlie PKD2 activation, we tested whether PIP(5)Kα could increase EGF-induced currents. In sharp contrast to TRPC5, PKD2 activity was suppressed by almost 85% by PIP(5)Kα. These results supported the idea that reduction of PIP₂ is required for EGF-induced conductance in LLC-PK₁ cells. However, they cannot exclude the possibility that both mechanisms may be operative for PKD2. For instance, EGF-induced activation of the Rac1/PIP(5)Kα/PIP₂ pathway may be initially required to translocate PKD2 to the plasma membrane, but subsequently PIP₂ in the microenvironment of the channel would need to be reduced to allow channel activation. The difference between TRPC5 and PKD2 could be that PKD2 is inhibited by PIP₂ while TRPC5 is simply insensitive to PIP₂. At present, our data favor the concept that the reduction of PIP₂ may be a necessary step in the activation mechanism of PKD2 by EGF. This idea is also supported by the physical interaction of PKD2, PLC-γ2, and EGFR.

Reduced PIP₂ in the microenvironment of channel proteins has been shown to have negative effects on TRPM7 (41) and positive effects on TRPL (10) and TRPV1 (8), respectively. Therefore, we would like to propose a similar mechanism of PKD2 activation by the EGF-induced reduction of the cellular PIP₂ pool size. However, we cannot exclude the possibility that reduced PIP₂ may have indirectly resulted in the activation of PKD2. It is interesting to note the similarity in the activation mechanisms of PKD2 and TRPV1 in response to growth factor stimulation (Fig. 8). Both channels are activated by growth factors, PKD2 by EGF and TRPV1 by NGF, through the reduction of PIP₂. In addition, TRPV1 can form a complex with TrkA and PLC-γ1 in transfected cells (8). We show here that PKD2 is also capable of forming a complex with EGFR and PLC-γ2. Interestingly, TRPV1 was unable to interact with EGFR, suggesting that specificity is based on the association of the channel and the RTK (8). Because PKD2 manifested a specific interaction with PLC-γ2 and not with PLC-γ1, we propose an additional level of specificity of the interaction between PLC isoforms and the channel. In agreement with in vitro coimmunoprecipitation experiments and functional data, LLC-PK₁ cells expressed predominantly PLC-γ2, while PLC-γ1 was undetectable.

The physiological significance of the NGF-induced activation of TRPV1 is to increase the sensitivity of TRPV1 to noxious stimuli which are unable to promote the breakdown of PIP₂ by themselves (8). Our data lead us to suggest an analogous mechanism of PKD2 activation, whereby EGF may sensitize PKD2 to mechanical stimulation (Fig. 9). This concept is supported by the finding that fluid flow-induced activation of PKD2 was insensitive to PLC inhibitors (34), suggesting that fluid shear stress alone may not be able to release PKD2 from PIP₂-mediated inhibition. Therefore, endogenous EGF could reduce the threshold of PKD2 activation to fluid shear stress through the reduction of PIP₂ in the primary cilium. EGF is present at very high levels in human urine (6); however, its function there has been a mystery because of the failure to detect EGFR in the apical surface of cells lining kidney tubules (11). Our finding that EGFR localized in the primary cilium which protrudes into the tubular lumen suggests that EGF may have a role in ciliary sensory function.

FIG. 9.

Diagrammatic representation of NGF-induced activation of TRPV1 and EGF-induced activation of PKD2.

Our study describes a novel mechanism on the regulation of PKD2 channel activity by EGF. Several points prompt us to conclude that this regulation is physiologically relevant. First, the deletion of EGFR produced kidney cysts in mice (43). Second, endogenous EGFR colocalized with PKD2 and PIP₂ at the primary cilium, an organelle very relevant to the pathophysiology of polycystic kidney disease. Third, PKD2 was essential for EGF-induced conductance. The mechanism by which the EGF-mediated regulation of PKD2 may contribute to normal tubulogenesis is unknown. However, based on previous studies of similar regulations of related channels by growth factors (8), we propose that EGF may increase the sensitization of PKD2 to mechanical stimulation at the primarily cilium. Certainly, other PLC-coupled signal transduction pathways may also result in PKD2 activation through a similar mechanism.