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Widely expressed in the adult central nervous system, the cellular prion protein (PrPC) is implicated in a variety of processes, including neuronal excitability. Dipeptidyl aminopeptidase-like protein 6 (DPP6) was first identified as a PrPC interactor using in vivo formaldehyde cross-linking of wild type (WT) mouse brain. This finding was confirmed in three cell lines and, because DPP6 directs the functional assembly of K+ channels, we assessed the impact of WT and mutant PrPC upon Kv4.2-based cell surface macromolecular complexes. Whereas a Gerstmann-Sträussler-Scheinker disease version of PrP with eight extra octarepeats was a loss of function both for complex formation and for modulation of Kv4.2 channels, WT PrPC, in a DPP6-dependent manner, modulated Kv4.2 channel properties, causing an increase in peak amplitude, a rightward shift of the voltage-dependent steady-state inactivation curve, a slower inactivation, and a faster recovery from steady-state inactivation. Thus, the net impact of wt PrPC was one of enhancement, which plays a critical role in the down-regulation of neuronal membrane excitability and is associated with a decreased susceptibility to seizures. Insofar as previous work has established a requirement for WT PrPC in the Aβ-dependent modulation of excitability in cholinergic basal forebrain neurons, our findings implicate PrPC regulation of Kv4.2 channels as a mechanism contributing to the effects of oligomeric Aβ upon neuronal excitability and viability.
Prion diseases involve the structural conversion of the primarily α-helical, cellular prion protein (PrPC)5 to an infectious, β-sheet-enriched form, PrPSc. The high degree of primary to tertiary structural conservation of mammalian PrPC (1, 2) leads to an expectation of an explicit phenotype in Prnp0/0 mice, but, aside from a total resistance to prion disease, this is not the case (3, 4). That said, within the inventory of subtle or disputed phenotypic changes in these mice are reports of impairment in GABAA receptor-mediated inhibition and long term potentiation (5–8). The diversity of altered end point measures is, to a certain extent, paralleled in the large number of reported PrPC-interacting proteins: the laminin receptor (9–11), the neural cell adhesion molecule (12, 13), stress-inducible protein-1 (14, 15), and NMDA receptors (16, 17) to name but a few (reviewed in Ref. 18).
In addition to previously described interacting proteins, dipeptidyl aminopeptidase-like protein 6 (DPP6; also known as DPPX) was identified using time-controlled transcardiac perfusion cross-linking while probing the PrPC interactome (19). DPP6 is an auxiliary subunit of pore-forming Kv4.2 channels (20, 21) and, together with most K+ channel-interacting protein (KChIP) isoforms (22), DPP6 increases Kv4.2 trafficking to the cell surface and is required for the reconstitution of the properties of the native channel complex in heterologous cells (23). A type II transmembrane protein, DPP6 has a number of splice variants differing in the length of the cytoplasmic, N-terminal portion (DPP6-S; short) (24). KChIPs are a family of intracellular Ca2+-binding proteins with four major isoforms (1–4) and at least 16 splice variants (25). Interactions between Kv4.2, KChIPs and DPP6 have been confirmed by proteomic analyses demonstrating the pull-down of KChIP1 to -3 in comparable ratios (26). Assembled Kv4 channel complexes mediate sub-threshold operation somatodendritic transient outward K+ currents (A-type K+ currents), which play important roles in the regulation of neuronal membrane excitability, somatodendritic signal integration, and long term potentiation (27, 28). Given the interplay between Kv4.2 channels and DPP6 in neuronal function (29) and the ability to cross-link DPP6 and PrPC in vivo (19), we sought to delineate the nature and repercussions of a DPP6-PrPC interaction. To this end, we investigated the impact of PrPC upon the properties of A-type K+ currents mediated by Kv4.2 channel complexes derived from co-expression of Kv4.2, KChIP2, and DPP6-S (30, 31).
cDNAs encoding Kv4.2 (MMM1012-9498428 clone 30356567, pYX-Asc) and KChIP2 (MMM1013-7511937 clone 4503251, pCR4-TOPO) (Open Biosystems) were inserted behind the CMV and EF1α promoters, respectively, of pBud.CE4 with Quick Ligase (New England Biolabs) to create pBud.CE4.Kv4.2.KChip2. To construct pBud.DPP6-S.RFP (co-expression driven from the CMV and EF1α promoters, respectively), total RNA was isolated from half mouse brains using acid guanidinium thiocyanate/phenol/chloroform extraction (32) and mRNA-purified using the Oligotex mRNA minikit (Qiagen). cDNA synthesis was performed in the presence of SUPERase-In RNase inhibitor (Ambion) using 0.5 μg of mRNA, Omniscript reverse transcriptase (Qiagen), and an oligo(dT) primer. PCR amplification of DPP6-S and DPP6-E was conducted using Pfu Turbo DNA polymerase (Invitrogen) with a nested PCR strategy and inserted between the BamHI and XbaI sites of the pcDNA3 mammalian expression vector (Invitrogen). A PstI site was added to the pcDNA3.DPP6-S for insertion into pBud.empty.GFP with Quick Ligase (New England Biolabs) after gel purification (QIAquick gel extraction kit (Qiagen)). GFP was replaced with RFP by digestion of pBud.DPP6-S.GFP and pBud.empty.RFP with NdeI and NheI, isolation of the desired fragments, and ligation. HA-tagged DPP6-S, DPP6Δcyto, and DPP6 deletion mutants were generated by standard PCR-based techniques. The secreted DPP6 ectodomain construct was generated by fusing the DPP6 ectodomain to the PrP N-terminal signal sequence using an introduced BsrGI site. The Thy-1 plasmid (Thy-1.2 isoform) was generated by amplification of the Thy-1 open reading frame from the MGC:62652 cDNA clone by PCR and then insertion into pcDNA3. PrP A116V and M128V HD dup PrP were created using the QuikChange (Stratagene) site-directed mutagenesis procedure with Pfu Turbo DNA polymerase. Other PrP mutants and Doppel plasmids were generated as described previously (33). All plasmids used for transfection were enriched using the EndoFree Plasmid Maxi kit (Qiagen).
HEK293T cells were maintained in DMEM with 10% FBS (Invitrogen). Transfections were performed with Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions. Cells used for electrophysiological recording were transfected with the following plasmids (fluorescent proteins co-expressed): pBud.DPP6-S.RFP, pBud.Kv4.2/KChIP2, pBud.wtPrP.GFP, pBud.octa13PrP.GFP, and pBud.empty.GFP. These transfection mixtures included (as noted) siRNA directed against the 3′-non-coding region of human PRNP mRNA (OriGene Trilencer-27 siRNA duplex; rGrGrCrUrUrArCrArArUrGrUrGrCrArCrUrGrArArUrCrGTT) or a scrambled control siRNA. 24 h after transfection, cells were trypsinized and plated on coverslips for electrophysiological recording the following day. Cells used for complex formation assays were transfected as above using constructs based on a pcDNA plasmid vector backbone (with identities indicated in the figure legends) except for Fig. 3C, which used a pBud vector.
Peptides were synthesized (containing an N-terminal cysteine for KLH conjugation), conjugated to maleimide-activated KLH (Pierce), and then injected into New Zealand White rabbits. 03J2 was raised against residues 507–522 (DKRRMFDLEANEEVQK), and 03K1 was raised against residues 788–803 (QDKLPTATAKEEEEED). Polyclonal antibodies were precipitated from serum using ammonium sulfate and affinity-purified using the corresponding immunogenic DPP6 peptide conjugated to a SulfoLink column (Pierce).
The assay was performed according to the manufacturer's instructions (Pierce).
24 h after transfection, cells (HEK293T, RK13, or N2a) were washed twice with PBS and incubated for 15 min at room temperature with 2 or 0.4% formaldehyde in PBS. The cross-linking reaction was quenched with 0.125 m glycine for 10 min. Lysis was performed with radioimmune precipitation assay buffer (50 mm Tris base, 150 mm NaCl, 0.1% SDS, 0.5% sodium deoxycholate, 1% Triton X-100) containing protease inhibitors (Roche Applied Science) at 4 °C.
120 μg of cell lysate was incubated overnight at 4 °C with 0.7 μg of α-HA antibody (Sigma) in a total volume of 50 μl. 100 μl of protein A/G-agarose beads were washed three times with radioimmune precipitation assay buffer containing protease inhibitor (Roche Applied Science) before adding the overnight incubation and were then incubated at room temperature for 2 h rotating end over end. After three washes with radioimmune precipitation assay buffer, the beads were washed a final time with water, and the complexes were eluted from the beads (and cross-links were reversed) by incubation in sample buffer at 95 °C for 30 min.
All animal protocols were in accordance with the Canadian Council on Animal Care and were approved by the Institutional Animal Care and Use Committees at the University of Alberta and the University of Toronto.
Prnp0/0 mice (ZrchI strain) were maintained on a C57/B6 background. DPP6df5J/Rw mice were a generous gift from John Schimenti and were maintained on a C3H background. Mice were perfused with saline, and half-brains were extracted and then either homogenized directly in nine volumes of 0.32 m sucrose with protease inhibitors (Roche Applied Science) or snap-frozen and stored at −80 °C.
Mice (either C57/B6 or FVB strains) were subjected to time-controlled transcardiac perfusion cross-linking, as described previously (19). Brains were homogenized in 0.32 m sucrose containing protease inhibitor (Roche Applied Science).
Cells were lysed with radioimmuneprecipitation assay lysis buffer. Brains were homogenized in 0.32 m sucrose. A BCA assay (Pierce) was performed to determine protein concentration. Samples were boiled in loading buffer (with the exception of samples to be used for Kv4.2 detection due to protein aggregation), and were electrophoresed with either Tris-glycine gels (8 or 12%) or 4–12% NuPAGE gradient gels (Invitrogen) and transferred to PVDF membranes (Millipore). Membranes were blocked with either 3% (Kv4.2) or 5% milk (all others except for SHA31 blots, which are not blocked) in TBS with 0.1% Tween (Fisher). Membranes were incubated overnight in the indicated primary antibody (DPP6 clones 03K1 and 03J2 (created in house), actin (Sigma), Kv4.2 clone K57/1 and KChIP2 clone K60/73 (NeuroMab), SHA31 (Spi-bio), 8B4 and 7A12 (generous gifts from Man-Sun Sy), HA (Sigma), and Thy-1 (graciously provided by Roger Morris)) and after washing were incubated in the appropriate HRP-conjugated secondary antibody for 2 h at room temperature (goat α-mouse or goat α-rabbit (Bio-Rad)), washed again, exposed to ECL (Pierce), and visualized with light-sensitive film (Fujifilm). Quantification was performed using ImageJ. The intensity of the ~191-kDa species detected by Western blotting was normalized to intensity of “monomeric input” PrP in the same gel lane and plotted using an arbitrary scale. p values were determined using Student's t test.
HEK293T cells were transfected with mouse PrP and HA-tagged DPP6-S (both on a pcDNA backbone) and an siRNA against endogenous human PrP. 24 h after transfection, cells were replated on glass coverslips and given 24 h to recover. After rinsing twice with PBS and fixation with 4% paraformaldehyde, cells were washed three times with PBS and incubated in Sha31 (1:5000) overnight at 4 °C with rocking. Following three PBS washes, cells were blocked with 2% goat serum (Invitrogen) and incubated with goat α-mouse Alexa Fluor 594 (1:300; Invitrogen) for 2 h at room temperature. After washing and permeabilization with 0.2% Triton X-100, cells were blocked and incubated overnight in 1:500 α-HA (Sigma) as above, washed three times, blocked, and incubated for 2 h with 1:300 goat α-mouse Alexa Fluor 488 (Invitrogen) at room temperature. Nuclei were stained with 1 μg/ml Hoechst and visualized using a Nikon Eclipse 90I motorized upright microscope (Nikon) and a CFI PL ×40/numerical aperture 0.75 lens (Nikon) using the following excitation/emission filter properties: 325–375/500–575 nm with a 495-nm long pass filter (blue channel), 440–510/475–575 nm with a 495-nm long pass filter (green channel), and 505–615/570–720 nm with a 595-nm long pass filter (red channel). Images were acquired with a Retiga 2000R monocooled camera, fast 1394, using NIS-Elements AR advanced research software at room temperature.
Whole-cell recordings were always applied to two groups of HEK293T cells on the same day to minimize variation resulting from cell manipulation prior to recordings. Fluorescence-positive individual HEK293T cells were visualized (by way of the GFP and RFP encoded in the bigenic plasmids) and selected for recording under an Axioscope 2 Fs microscope (Zeiss) at ×60 magnification. Oxygenated external solution was bath-perfused at a rate of 1.0 ml/min. The external solution contained 140 mm NaCl, 2.5 mm KCl, 1.2 mm MgCl2, 1.5 mm CaCl2, 10 mm HEPES, and 10 mm glucose, pH 7.4. Recording pipette solution was composed of 140 mm potassium methylsulfate, 5 mm MgCl2, 10 mm HEPES, 2 mm Na2ATP, 0.2 mm NaGTP, and 0.5 mm EGTA, pH 7.3. Borosilicate glass capillaries (thin wall with filament, 1.5 mm; World Precision Instruments) were pulled with a Narishige (PP-83) puller to yield recording pipettes with resistances of 2–4 megaohms. Series resistance of 4–7 megaohms in whole-cell configuration was compensated by 60%. Three different stimulation protocols in our experiments were employed to evoke Kv4.2-mediated A-type K+ currents, which were used for building relationship of voltage-dependent activation (activation curve), steady-state inactivation (inactivation curve), and rate of recovery time from steady-state inactivation (curve of recovery time). A-type K+ currents for making the activation curves were evoked for 300 ms, depolarizing membrane potential ranging from −60 to 50 mV following a conditioning pulse of −110 mV for 400 ms. The A-type K+ currents used to build the inactivation curves were evoked during a fixed depolarizing potential of 30 mV following 400-ms conditioning pulses of various membrane potentials, ranging from −110 to −20 mV. A-type K+ currents for plotting the curve of the recovery rate were elicited during a fixed 30-mV depolarizing membrane potential for 300 ms following a series of increasing time intervals of conditioning hyperpolarizing potential at −110 mV. All A-type currents presented were subject to subtraction of the outward currents (endogenous currents) evoked by a conditioning pulse of −20 mV from that evoked by any protocols mentioned above. The current signals were acquired at a bandwidth of 10 kHz using pClamp version 9.2 software and filtered with a 5-kHz low pass Bessel filter using an Axopatch 200B amplifier (Molecular Devices).
Time to half-inactivation (t½) is the time at which 50% of peak amplitude is inactivated at the indicated voltage. The activation curve was obtained by plotting normalized conductance of peak amplitude of A-type K+ currents against its corresponding depolarizing membrane potential. The inactivation curve was built by plotting normalized peak amplitude of A-type currents against its corresponding conditioning membrane potential. The curve of recovery rate from steady-state inactivation was obtained by plotting the peak amplitude of A-type currents against its corresponding time intervals at a conditioning hyperpolarizing membrane potential of −110 mV. The time constant (trec) of recovery rate was measured by fitting the curve to a single exponential function. Average values were expressed as mean ± S.E., and statistical significance was evaluated by means of the two-tailed Student's unpaired t test. The significance level for the t tests was set at p < 0.05.
To facilitate our analyses of DPP6 and DPP6-containing protein complexes, two polyclonal peptide antisera, 03K1 and 03J2, were raised in rabbits (Fig. 1A). Western blot analysis of brain homogenate from WT and DPP6-deficient mice (DPP6df5J/Rw) (34) assessed specificity (not shown), revealing immunoreactive species of ~110 kDa in accord with glycosylation of full-length DPP6 (~91 kDa) (35). Immunohistochemistry demonstrated the widespread expression of DPP6 in the mouse brain (not shown), confirming previous reports (36). The crystal structure of recombinant DPP6 is dimeric (35), and protein species with an electrophoretic mobility consistent with dimers were observed in transfected cells and mouse brain homogenate after cross-linking and analysis by Western blot (Fig. 1, B and C). When N2a cells expressing endogenous PrPC were transfected with plasmids encoding DPP6 (DPP6-S and DPP6-E) (23), cross-linked, and immunoprecipitated with 7A12 (α-PrP) and membranes were probed with 03K1, a novel species was observed with a mobility of ~190 kDa (Fig. 1D). These data indicate that PrPC is located in membrane subdomains that harbor dimeric DPP6 or that PrP and DPP6 exist in direct physical contact in a protein complex with a stoichiometry that totals to an Mr of ~190,000. Reversal of cross-links yielded a species that co-migrated with full-length glycosylated DPP6-S. In a reciprocal analysis, following cross-linking, cells expressing HA-tagged DPP6-S yielded high Mr species and, after reversal, a signal compatible with glycosylated PrPC (Fig. 1E). Co-immunoprecipitation was also achieved in WT mouse brain without cross-linking using either 8B4 (α-PrP) or 03K1 for pull-down (Fig. 1F).
Subsequently, mutant alleles were used to map regions within PrPC and DPP6 required for association (Figs. 2A, ,33A, and and44A). Deleting the intracellular, N-terminal portion of DPP6 (“DPP6Δcyto”) had no effect upon complex formation with PrPC (Fig. 2, A, B, and E). “secDPP6,” where DPP6 residues 56–803 are prefaced by the PrP N-terminal signal peptide, causing secretion from the cell, was not associated with complex formation with PrPC (Fig. 2, A, C, E, and F). Because this secDPP6 allele was readily detected in the culture medium, this implies a requirement for membrane association for complex formation rather than presence in the extracellular milieu. Also, anchoring to the membrane by a glycosylphosphatidylinositol moiety was not sufficient for complex formation because the assay failed to detect DPP6·Thy-1 complexes (37) (Fig. 2D).
To delineate the role of the DPP6 ectodomain (35) in complex formation with PrPC, we used a series of DPP6 deletions with an N-terminal HA tag (Fig. 3A). Control experiments confirmed that these DPP6 constructs were expressed at the cell surface (Fig. 3B), and following cross-linking and immunoprecipitation with an α-HA antibody, cross-links were reversed before blot analysis. PrPC was recovered in conjunction with all DPP6 deletion mutants tested (Fig. 3C), leading to the inference that DPP6 residues 56–80 contribute to complex formation. This was investigated further by performing the cross-linking assay with an internally deleted form of DPP6 Δ56–81 (this deletion covers the same interval but extends one residue further to isoleucine 81 to maintain the length of the hydrophobic TM region). For DPP6 Δ56–81, the ability to form complexes was lower than for WT DPP6 (Fig. 3D), underscoring a role for the juxtamembrane region.
With regard to PrP, N-terminal deletions Δ23–88 and Δ32–121, which encompass most or all of the octarepeats, had subtle effects on complex formation (Fig. 4, A, B, and E). The contribution of an interval that encompasses the first β-strand of the PrP structure (38, 39) to complex formation with DPP6 is illustrated by the performance of the Δ32–134 PrP allele (Fig. 4, A and B). We also created PrP alleles with more C-terminal deletion intervals; aside from theoretical caveats (see “Discussion”), these were also limited by lower expression levels than WT PrP and were not examined further. A survey of PrP alleles from genetic prion diseases defined a striking result associated with the octarepeat region (33, 41) but not with mutations C-terminal to this position (42–44). octa13 PrP, an expansion of the octarepeats to a total of 13, was less efficient than WT PrP at complex formation with DPP6 (Fig. 4, A, C, and D). Densitometry revealed that octa13 PrP formed ~191-kDa complexes at a level of 23.3 ± 16.4% that of WT PrP (100 ± 12.4%; n = 3, p < 0.002). As a preface to the functional studies described below, this result was confirmed in cross-linking studies of HEK293T cells (Fig. 4D). Furthermore, we confirmed that full-length HA-tagged DPP6-S and PrPC co-localize at the cell surface of the HEK293T cells used for electrophysiology (Fig. 4F). Last, interactions between PrPC and DPP6 were found to take place in cis in a cell biological sense, as demonstrated by the lack of complex formation when cells expressing PrPC or DPP6 are co-cultured before cross-linking and lysis (Fig. 4G). Interestingly, Doppel, with a three-dimensional fold similar to that of the PrPC C terminus (40), can also form complexes with DPP6 isoforms (Fig. 5).
Following co-expression of the components of the Kv4.2 channel complex (Kv4.2, KChIP2, and DPP6-S; Fig. 6A), whole-cell recordings were performed on transiently transfected HEK293T cells. Two series of A-type K+ currents mediated by the Kv4.2 channel complex were investigated in the presence and absence of exogenous PrPC (the isolate of HEK293T cells used here expressed endogenous human PrPC; Figs. 6A, ,88A, ,99A, and and1010A). As seen in Fig. 6B, an A-type outward K+ current was generated in response to depolarizing potentials. Following a rapid rise to peak amplitude, the current rapidly decayed despite a continued depolarizing step command. In the presence of exogenous PrPC, the peak amplitude of the A-type K+ currents at 20 mV was larger (14.5 ± 0.9 nA; average ± S.E., n = 17) than that mediated by the Kv4.2 channel complex in its absence (10.2 ± 1.0 nA; n = 21, p < 0.05) (Fig. 6C). The curve of voltage-dependent activation of A-type K+ currents mediated by Kv4.2 channel complexes in the presence (triangles) and absence (squares) of exogenous PrPC was created by plotting averaged normalized conductance (G/Gmax) against corresponding depolarizing potential (Fig. 6D). There was no significant difference in the voltage-dependent activation between the two groups.
To establish voltage-dependent steady-state inactivation, A-type K+ currents were evoked by another voltage stimulus protocol (see “Experimental Procedures”). To highlight differences, we have shown two traces of A-type K+ currents mediated by the Kv4.2 channel complex alone (left) and with exogenous PrPC (right) at conditioning potentials of −110 and −60 mV, respectively (Fig. 6E). The ratio of normalized peak amplitudes of the currents evoked at conditioning potentials of −60 and −110 mV was 0.29 in the absence of PrPC versus 0.56 in its presence. The plot for voltage-dependent steady-state inactivation was obtained by plotting corresponding average normalized currents (I/Imax) against conditioning potential in the presence (triangles) and absence of exogenous PrPC (squares) (Fig. 6F). Significant differences between these two groups were noted at conditioning potentials of −50, −60, and −70 mV (n = 10, p < 0.05).
To measure decay of A-type K+ current from peak amplitude to base line, we used the time at which 50% of peak amplitude was inactivated at a given depolarizing potential (half-inactivation time) to quantitatively describe the time course for inactivation (20). Two representative traces of A-type K+ currents evoked by a depolarizing potential of 50 mV in Fig. 6B are shown with an expanded time scale and correspond to the Kv4.2 channel complex with or without exogenous PrPC (Fig. 7A). The half-inactivation time was increased from 17.6 to 31.2 ms by the presence of exogenous PrPC. On average, the half-inactivation time of A-type K+ currents mediated by Kv4.2 channel complexes in the presence of exogenous PrPC (n = 8) was significantly longer than that mediated by the Kv4.2 channel complex alone (n = 10) at all but two depolarizing potentials (Fig. 7B; p < 0.05).
We noted above (Fig. 6F) for HEK293T cells expressing Kv4.2 complexes that when the conditioning potential applied is more positive than −40 mV, it is not possible to evoke an A-type K+ current by stepping to a depolarizing potential of 30 mV. This status can be referred to as complete voltage-dependent steady-state inactivation of Kv4.2 channels and can be removed by first stepping to a conditioning hyperpolarizing potential (e.g. −110 mV) for a short duration. Consequently, to measure how fast A-type currents can recover from complete voltage-dependent steady-state inactivation, we recorded A-type K+ currents evoked by a different stimulus protocol (Fig. 7C). Representative traces of the resulting A-type K+ currents are shown for the presence and absence of exogenous PrPC (Fig. 7C) with conditioning potentials of −110 mV for 5 and 170 ms, respectively. The ratios of the amplitudes of the currents measured with a pulse duration of 5-ms versus 170 ms at conditioning potentials of −110 mV were 0.24 (without) and 0.58 (with exogenous PrPC). The average recovery rate from steady-state inactivation was obtained by plotting the time of conditioning hyperpolarizing potential at −110 mV against corresponding normalized currents (I/Imax) (Fig. 7D). The recovery rate was significantly faster in the presence of exogenous PrPC (n = 8) than in its absence (n = 10) at the first two time intervals of 5 and 25 ms (p < 0.05). In both groups, recovery from steady state was virtually complete after 100 ms following a conditioning hyperpolarizing potential of −110 mV.
To ascertain whether or not the effects of PrPC in this system are mediated through DPP6-S, we characterized the modulatory properties of PrPC upon Kv4.2-mediated A-type K+ currents in the absence of co-expressed DPP6-S (Fig. 8A). These A-type K+ currents showed smaller peak amplitudes, a right shift of the activation curve, a right shift of the steady-state inactivation curve, a longer half-inactivation time, and a slower recovery rate from steady-state inactivation. These changes in the properties of A-type K+ currents without DPP6-S are consistent with the previous findings observed by another group (24). However, exogenous PrPC did not have a significant effect upon any A-type K+ current properties in the absence of DPP6-S (Fig. 8, B–F).
Next, we studied the impact of co-expression of octa13 PrP (Fig. 4A) with Kv4.2 complexes that include DPP6-S. Although the stimulus protocols for these studies remained the same as in previous recordings (with the exception of those used to investigate recovery rate from steady-state inactivation; Fig. 7, C and D), we adjusted two aspects of the experimental design. First, the time interval for conditioning hyperpolarizing potential was reduced from the previous 20 ms to 5 ms to acquire more data points. Second, we used siRNA against human PrP mRNA to reduce the levels of endogenous PrPC (Fig. 9A); by this means, we reduced cross-talk from WT PrPC that could complicate the interpretation of data associated with transgene-encoded PrP. In this experimental context, WT PrPC was found to modulate the Kv4.2 channel complex in the same manner as our previous data (Fig. 9, B–F). However, the properties of A-type K+ currents in the presence of octa13 PrP showed no significant difference from those mediated by the Kv4.2 channel complex alone (Fig. 9, B–F, and Table 1). These data indicate that octa13 PrP is a loss of function with regard to modulation of Kv4.2, a finding that parallels its >75% loss in efficiency at forming cross-linked complexes with DPP6-S (Fig. 4, C and D).
In a final set of analyses, we further investigated the inference that residues 56–80 of DPP6 are required for complex formation with PrPC. A-type K+ currents mediated by the Kv4.2 complex composed of Kv4.2, KChIP2, and DPP6 Δ56–81 showed smaller peak amplitudes, a right shift of the activation curve, a right shift of the steady-state inactivation curve, a longer half-inactivation time, and a slower recovery rate from steady-state inactivation (Fig. 10). These changes in the properties of A-type K+ currents are similar to those observed in the absence of DPP6-S (Fig. 8), indicating that DPP6 Δ56–81 is not fully functional. However, exogenous PrPC did not modulate the properties of these A-type K+ currents, further supporting the idea that this juxtamembrane region of DPP6 is required for complex formation with PrPC (Figs. 3 and and1010).
PrPC is highly expressed in the central nervous system (45–47), and consequently, its role in regulating neuronal excitability is of great interest. Prior studies have centered upon perturbations in electrophysiological recordings made from brain slices of Prnp0/0 mice (5–8, 16, 17, 48–51). Here we have investigated the influence of PrPC upon Kv4.2 channel complexes by their reconstitution in HEK293T cells to study the currents produced by these complexes in isolation. These cells have been reported to have small endogenous delayed rectifier K+ currents (52). We found identical currents in our HEK 293T cell isolate (generally smaller than 200 pA) but did not detect A-type K+ currents, as previously reported by another group (53). Therefore, the A-type K+ currents produced here (and measuring in the nanoampere range) by reconstitution of the Kv4.2 channel complex are not contaminated by the presence of endogenous currents. More recently, expression of PrP Δ105–125 in transfected cells has been reported to induce spontaneous non-selective, cation-permeable ion currents (54, 55). In this instance, because of the striking nature of the effect observed in HEK293T cells, we undertook parallel studies with the same allele inserted in our expression vectors, but none displayed a spontaneous ionic current (n = 10). To observe the currents, the cells were held at either −80 or +80 mV for several min. Under these conditions, it would be unlikely to miss any spontaneous current activity (54, 55). These experiments were performed both at 23 and 34 °C and with pipette solutions containing either 0.5 mm EGTA or 10 mm EGTA (54, 55). Although these data (not shown) did not support the concept of a solitary action of PrP for this particular allele under our designated experimental conditions, WT PrPC impacted the performance of co-expressed DPP6·Kv4.2 complexes, as elaborated below.
In prior analyses, PrPC has been implicated in the modulation of a variety of ion channels, including GABAA receptor/channels (5, 6, 8), calcium-dependent K+ channels, and NMDA receptors/channels (16, 17, 48, 56, 57); stress-inducible protein-1-dependent intracellular Ca2+ fluxes mediated by the α7 nicotinic acetylcholine receptor (14, 15, 58); and an AMPA-dependent Zn2+ reuptake phenomenon (59). Remarkably, these studies convey a diversity of mechanisms whereby PrPC modulates ion channels and neuronal excitability. For instance, enhanced and drastically prolonged NMDA-evoked currents in PrPC knock-out mouse neurons were the result of a functional up-regulation of NMDA receptors containing NR2D subunits (16). On the other hand, impaired and depressed Ca2+-dependent after-hyperpolarization potential in PrPC knock-out mouse neurons arises from an increased intracellular Ca2+ buffering capability (48, 57). In this case, the free Ca2+ through influx of activated voltage-gated Ca2+ channels is decreased and in turn depresses the after-hyperpolarization potential. Here, the modulation of Kv4.2 properties by PrPC requires interaction with DPP6-S and indicates that yet another, presently unknown, mechanism is employed. One possibility is increased trafficking of DPP6-S in the presence of PrPC. A larger current amplitude and a faster recovery time from steady-state inactivation can be attributed to an increase of DPP6-S at the cell surface (20). However, the modulation of the voltage dependence of inactivation and half-inactivation time by PrPC in this study counters the effect of DPP6-S. This suggests that the modulation of these channels by PrPC occurs, at least partially, in a manner distinct from trafficking. Before addressing the puzzle presented by the pleiotropic actions of PrPC, we will first consider molecular and mechanistic aspects of the PrPC/A-type current paradigm.
In genetic mapping of determinants necessary for PrPC-DPP6 interactions (Figs. 11–5), WT N-terminal sequences up to residue 121, which are considered natively unstructured in the unmetallated form of PrP (39, 60), were not required. Deletions that begin to encroach on the C-terminal globular domain diminished the interaction (Fig. 4B). Although there are certain caveats concerning expression levels, chaperone interactions, and global folding that apply to the use of C-terminal deletions (61, 62), as yet we have been unable to find a crucial, common segment of PrP that is required for complex formation. We infer that a natively structured PrPC globular domain (rather than a linear “epitope”) is essential for complex formation. This is supported by the finding that Doppel also forms high molecular weight complexes with DPP6 (Fig. 5). For DPP6-S, the intracellular portion had no effect on complex formation with PrPC, but there was a requirement for anchoring to the cell membrane (Fig. 2). With progressive ectodomain deletions, we determined that residues 1–81 retained the ability to immunoprecipitate PrPC following cross-linking (Fig. 3, A–C). Because the 55 N-terminal residues of DPP6-S are found either on the cytoplasmic side of or spanning the membrane (and thus inaccessible to PrPC), these data lead to an inference that the DPP6-S juxtamembrane region (residues 56–80) either complexes directly with the globular domain of PrP or is retained in PrP-enriched membrane domains by the action of an intermediary protein. This inference was bolstered by analyses of cells expressing an internal deletion (residues 56–81) incorporated into full-length DPP6-S (Fig. 3, A and D). These findings have a potential parallel in analyses and complement data indicating that the extracellular portion of DPP6 is not necessary for modulation of Kv4.2 channel properties (63).
To explain the curiously diverse actions of PrPC in different experimental paradigms, Linden et al. (64) hypothesized its action as a dynamic scaffold at the cell surface, one that not only can assemble membrane proteins in a cellular signaling microenvironment but also can impact or adjust function. Findings presented here seem compatible with this proposal as can the loss-of-function phenotype of the octa13 allele in electrophysiological assays (Fig. 9). Aside from the inefficiency of octa13 PrP at forming DPP6-S-containing complexes (Fig. 4, C and D), our studies revealed that it is incapable of forming another type of complex. This effect is apparent in both RK13 and HEK293T cells (Fig. 4, C and D), where a complex of ~110 kDa containing WT PrP (indicated by open arrows) is absent for octa13 PrP. The ~110-kDa complex could represent a PrPC·DPP6-S monomer complex or interaction with a different protein, as suggested by the failure to detect endogenous DPP6 in HEK293T cells (Figs. 6A, ,99A, and and1010A). In terms of potentially analogous effects for protein-protein interactions, it is notable that PrP with 14 octarepeats was loss of function for inhibiting β-cleavage of APP, whereas PrPC without the octarepeat region retained activity (65). Although a direct physical interaction between PrPC and the β-cleaving enzyme has been questioned (66), the concept of plasma membrane-signaling microdomains orchestrated by PrPC may have considerable merit.
The >75% reduction in formation of ~191- and ~110-kDa complexes by octa13 PrP is notable (Fig. 4, C and D) and serves as a useful control to measure against the performance of WT PrPC. However, two other GSS alleles tested here did not behave in the same way, and GSS pathogenesis is normally considered as “gain of function” due to misfolding of PrP. Whether the loss-of-function effect is due to an approximately one-third reduction in protein at the cell surface (50.1 ± 7.5% versus 78.1 ± 3.9% for octa13 versus WT, as measured by a biotinylation assay; not shown) is not clear. The electrophysiological observations made here apply to non-neuronal HEK293T cells, and it remains possible that excitable cells may behave differently, but given prominent expression of DPP6 and PrPC in the CNS, where they are located in close proximity (19), our findings do broach the question as to how modulation of A-type K+ currents by WT PrPC via DPP6-S might feature within a broader spectrum of neurological diseases. Although the verdict may still be out on the DPP6 locus as a significant risk factor for autism spectrum disorders (67, 68) and ALS (69–71), other possibilities remain. WT PrPC modifies the A-type K+ currents from reconstituted Kv4.2 channel complexes by increasing peak amplitude, shifting the voltage-dependent steady-state inactivation curve to the right (more positive membrane potential), slowing inactivation, and decreasing recovery time from steady-state inactivation. This overall impact of enhancement prompts two speculations. First, enhancement plays a critical role in the down-regulation of neuronal membrane excitability and is associated with a decreased susceptibility to seizures (27, 28, 72); interestingly, a reported phenotype of Prnp0/0 mice is an increased vulnerability to drug-induced seizures (73, 74). Second, our previous work has established that PrPC is essential for the modulation of neuronal excitability by Aβ oligomers in cholinergic basal forebrain neurons (75). Thus, the present findings implicate PrPC regulation of Kv4.2 channels as a mechanism that could contribute to the observed effects of oligomeric Aβ (and perhaps other types of protein aggregate assemblies (76)) on neuronal excitability and viability.
DPP6df5J/Rw mice were a generous gift from Dr. John Schimenti, and we thank Dr. Nam-Chaing Wang (Hospital for Sick Children, Toronto, Canada) for peptide syntheses. α-Thy-1 was a gift from Dr. Roger Morris (King's College, London).
*This work was supported by grants from the PrioNet Network Centres of Excellence, Alberta Prion Research Institute Grant APRI 200600070, Alberta Innovates-Health Solutions Grant AHFMR 201000628, Canadian Institutes of Health Research Grants MOP36377 and 93601, and the Canadian Foundation for Innovation.
5The abbreviations used are: