• afterhyperpolarization;
  • l-type voltage-gated calcium channels;
  • phosphatidylinositol 3-kinase;
  • potassium channels;
  • prion protein;
  • PrP knock out mice


  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Previous neurophysiological studies on prion protein deficient (Prnp–/–) mice have revealed a significant reduction of slow afterhyperpolarization currents (sIAHP) in hippocampal CA1 pyramidal cells. Here we aim to determine whether loss of PrPC. directly affects the potassium channels underlying sIAHP or if sIAHP is indirectly disturbed by altered intracellular Ca2+ fluxes. Patch-clamp measurements and confocal Ca2+ imaging in acute hippocampal slice preparations of Prnp–/– mice compared to littermate control mice revealed a reduced Ca2+ rise in CA1 neurons lacking PrPC following a depolarization protocol known to induce sIAHP. Moreover, we observed a reduced Ca2+ influx via l-type voltage gated calcium channels (VGCCs). No differences were observed in the protein expression of the pore forming α1 subunit of VGCCs Prnp–/– mice. Surprisingly, the β2 subunit, critically involved in the transport of the α1 subunit to the plasma membrane, was found to be up-regulated in knock out hippocampal tissue. On mRNA level however, no differences could be detected for the α1C, D and β2–4 subunits. In conclusion our data support the notion that lack of PrPC. does not directly affect the potassium channels underlying sIAHP, but modulates these channels due to its effect on the intracellular free Ca2+ concentration via a reduced Ca2+ influx through l-type VGCCs.

Abbreviations used

artificial cerebrospinal fluid




afterhyperpolarization current


slow afterhyperpolarization current


chinese hamster ovary


charge transfer cytosolic shell


charge transfer membraneous shell




cellular prion protein


prion protein gene


region of interest

SK channel

small conductance channel


phosphatidylinositol 3-kinase


voltage gated Ca2 channels

The cellular prion protein (PrPC) is a copper-binding glycophosphatidylinositol (GPI)-linked glycoprotein. The misfolded isoform PrPSc causes neurodegenerative diseases like Creutzfeldt–Jakob disease, scrapie and Bovine Spongiform Encephalopathy. Although PrPC is expressed most abundantly in the CNS, several types of non-neuronal cells including blood lymphocytes, gastro-epithelial cells, heart, kidney and muscle also express PrPC (Horiuchi et al. 1995). In the brain, PrPC particularly localizes to synaptic membranes as shown by immunohistochemical, immunoelectron microscopic and synaptic plasma membrane fraction studies (Herms et al. 1999; Fournier et al. 2000).

The predominant synaptic localization of PrPC suggests a function related to synaptic transmission and neuronal excitability. Although PrPC-deficient mice develop normally, extensive studies revealed a high number of subtle alterations, including mossy fiber reorganization in the hippocampus (Colling et al. 1997), impaired long-term potentiation (Collinge et al. 1994; Manson et al. 1995), decreased field excitatory postsynaptic potentials (Carleton et al. 2001), reduced copper concentration in synaptosomes (Brown et al. 1997a; Herms et al. 1999), cognitive defects (Criado et al. 2005) and impaired neurite outgrowth (Santuccione et al. 2005). Other studies assign PrPC neuroprotective and antiapoptotic function (Li and Harris 2005; Roucou et al. 2005) or attribute PrPC to signal transduction pathways (Chiarini et al. 2002; Schneider et al. 2003; Vassallo et al. 2005; Krebs et al. 2006). A consistent electrophysiological phenotype observed in independently generated knock out mice lines is the impairment of slow afterhyperpolarization currents (Colling et al. 1996; Mallucci et al. 2002; Asante et al. 2004). These currents are of critical importance for neuronal excitability. Mouse hippocampal CA1 pyramidal cells display predominantly two afterhyperpolarization (AHP) currents, namely IAHP and slow IAHP(sIAHP) (Krause et al. 2002). Potassium channels underlying the IAHP belong to the small conductance (SK) channel family and are characterized by their sensitivity to the bee venom apamin (Stocker et al. 2004). In contrast, potassium channels underlying sIAHP are insensitive to apamin, but sensitive to a number of different neurotransmitters (Stocker et al. 2004). Moreover, IAHP and sIAHP can be clearly distinguished by their different decay times. While IAHP deactivates in the range of hundreds of milliseconds, sIAHP displays a much slower time course decaying in the range of seconds. Both currents are voltage insensitive and Ca2+-dependent. After membrane depolarization or action potential firing IAHP currents are activated by Ca2+ entering the cell via voltage-gated Ca2+ channels (VGCCs). The activation of sIAHP currents is delayed in comparison to the rise of intracellular free Ca2+ revealing a gap between intracellular free Ca2+ and onset of sIAHP. Possible reasons are slow diffusion of Ca2+ from the point of origin to the channels responsible for sIAHP (Lancaster and Nicoll 1987; Lancaster and Zucker 1994; Zhang and McBain 1995), Ca2+-induced Ca2+ release (Sah and McLachlan 1991; Zhang and McBain 1995; Lasser-Ross et al. 1997; Moore et al. 1998; Tanabe et al. 1998), involvement of an enzymatic step (Lasser-Ross et al. 1997; Moore et al. 1998), delayed facilitation of Ca2+ channels supplying the Ca2+ for the activation of sIAHP (Cloues et al. 1997; Bowden et al. 2001), intrinsically slow activation and deactivation kinetics of the sIAHP channels (Schwindt et al. 1992; Hocherman et al. 1992; Sah 1993; Sah and Clements 1999) or a very slow continued Ca2+ entry through l-type VGCCs (Marrion and Lima 2005).

Here we aim to answer the question whether loss of PrPC directly affects the potassium channel that underlies sIAHP or if this channel is secondarily affected due to a possible role of PrPC in modifying cellular Ca2+ homoeostasis. The latter was suggested from previous work of our group on cultured cerebellar granule cells (Herms et al. 2000), as well as from a recent study exploiting differentially localized aequorins to monitor intracellular Ca2+ concentration in different cell compartments (Brini et al. 2005). Both studies indicated that PrPC affects the cytosolic free Ca2+ concentration. In the present study, patch-clamp experiments were performed on mouse hippocampal Prnp–/– CA1 neurons to quantify sIAHP amplitudes, combined with confocal fluorimetric Ca2+ measurements to quantify the intracellular Ca2+ flux. Since IAHPs are mainly dependent on calcium influx via l-type voltage gated calcium channels we analyzed those channels as well.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References


Mice kept on a C57BL6 × 129Sv background were genotyped by PCR on genomic DNA from tail biopsies. Littermates were generated by crossing heterozygous C57BL6 × 129Sv Prnp+/– mice (Bueler et al. 1992). Mice were genotyped using the primer pair P2 (CTTCAGCCTAAATACTGGG) and P4 (ACCACGAGAATGCGAAGG), yielding a 840 bp fragment for Prnp+/+, a 1.4 kb fragment for Prnp–/– or both fragments for heterozygous Prn-p+/– mice. Mice used for each experiment were randomly grouped into Prnp+/+ and Prnp–/– by an independent person without any knowledge of the experimenter. All experiments were thus conducted in a double-blind manner, and the code was broken only after completion of the experimental work and data analysis.

Slice preparation

Acute slices were obtained from 18 to 21 day old C57BL6 × 129Sv Prnp+/+ and Prnp–/– littermate mice. Throughout the text, it is specified to which group we refer according to the experiment performed. Transversal hippocampal slices (300–400 µm) were cut with a vibratome (Leica VT1000E, Solms, Germany) in ice-cold oxygenated ACSF (artificial cerebrospinal fluid) and maintained in oxygenated ACSF at 37°C up to 6 h. ACSF contained in mM: 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, 10 Glucose, bubbled with 95% O2 and 5% CO2 (Carbogen), pH 7.3 with NaOH. Slices were allowed to recover for at least 45 min in oxygenated ACSF at 37°C before being transferred to the recording chamber.


Tight-seal whole-cell voltage-clamp recordings were obtained from somata of CA1 pyramidal cells visualized by differential interference contrast using an EPC-9 patch-clamp amplifier (HEKA Electronics, Lambrecht, Germany). Patch-clamp electrodes (3.5–5 M) were filled with an intracellular solution containing in mM: 135 potassium methylsulfate, 8 NaCl, 10 HEPES, 2 Mg2-ATP, and 0.3 Na3-GTP, 0.5 Calcium Green-5N (Calcium Green-5N was only included when performing Ca2+ imaging), pH 7.3 with KOH (osmolarity 300 mOsm). The intracellular solution used for the measurement of VGCCs contained (in mM): 120 CsCl2, 20 CsFl, 4 MgCl2, 1 EGTA, 4 Mg2-ATP, 10 HEPES, pH 7.3 with CsOH (320 mOsm). Recordings were performed at 23°C in a submerged recording chamber with a constant flow of ACSF (1 mL/min). When VGCCs were studied, bath solution contained (in mM): 148 TEA-Cl, 4 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, 0.001 TTX, and 0.01 nifedipine (where indicated), pH 7.3 with TEA-OH (320 mOsm). Bath and intracellular solution were devoid of sodium to avoid measurement of sodium currents. Additionally TTX was added to block sodium currents. Neurons were voltage-clamped at −60 mV, and 200 ms depolarizing pulses to +20 mV were delivered every 30 s. Depolarizing pulses were sufficient to activate AHP currents. Analysis was carried out using the HEKA software PulseFit (Heka Electronics, Lambrecht, Germany). The decays of IAHP and sIAHP currents were determined by fitting the trace with a single exponential equation. The amplitude of IAHP was determined at the peak of the current and the sIAHP amplitude was measured 700 ms post stimulus end. For whole-cell patch-clamp recordings of voltage-gated Ca2+ currents depolarizing pulse commands 200 ms in duration were applied from a holding potential of −70 mV. The pulse command was stepped in 10 mV steps from −100 to +30 mV. Between each sweep a 1 s interval was inserted. Current-voltage (I-V) relationships were determined by measuring the peak amplitude value at the different command potentials. The inactivating component of the current was determined for the maximum amplitude by calculating the percent difference between the peak amplitude value and the amplitude value at the end of the command potential step (Fig. 4A, ΔX).


Figure 4.  Nifedipine-sensitive voltage-gated l-type Ca2+-channel currents are diminished in hippocampal CA1 neurons of Prnp–/– mice. (a, b) Mean I-V relationships of Prnp+/+ (n = 10, 3 animals) and Prnp–/– (n = 10, 3 animals) hippocampal CA1 neurons treated with or without 10 µm nifedipine. Nifedipine blocked the l-type mediated part of the measured whole-cell current. Insets show representative example current traces evoked by a 200 ms depolarizing voltage step from a holding potential of −70 mV to − 20 mV that were partially blocked by the application of nifedipine. The symbol ΔX indicates the value of the inactivating component of the current. (c) The maximum current amplitude of Prnp–/– CA1 neurons (n = 9, 3 animals) is significantly reduced compared to Prnp+/+ (n = 9, 3 animals, *p < 0.05; Student's t-test). Application of nifedipine reduced current amplitudes of both Prnp+/+ (n = 9, 3 animals) and Prnp–/– (n = 9, 3 animals) CA1 neurons to a similar level (n.s. p > 0.05; Student's t-test). (d) Percent inhibition of the maximum current amplitude by nifedipine. Percent inhibition by nifedipine is significantly higher in Prnp+/+ (n = 9, 3 animals) CA1 neurons compared to Prnp–/– (n = 9, 3 animals, *p < 0.05; Student's t-test). (e) Inactivation of the maximum current amplitude of Prnp+/+ and Prnp–/– CA1 neurons treated with or without nifedipine (10 µm). The percent inactivation of Prnp+/+ (n = 9, 3 animals) CA1 neurons is significantly higher than in Prnp–/– (n = 9, 3 animals, *p < 0.05; Student's t-test) CA1 neurons. Application of nifedipine reduced the percent inactivation of Prnp–/– and Prnp+/+ to a comparable value (n.s. p > 0.05; Student's t-test). Box plot diagrams contain 10, 25, 75 and 90% percentiles; red/grey bar: mean; black bar: median.

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Ca2+ imaging

Fluorimetric Ca2+ measurements were performed using a confocal laser-scanning system (LSM510 Meta, Zeiss, Oberkochen, Germany) mounted to an upright microscope (Axioplan, Zeiss) equipped with a water-immersion objective (Achroplan, Zeiss; × 40 NA 0.75). The image-acquisition-rate was 10 Hz to ensure resolving of the time course of the Ca2+ transient. Calcium Green-5N (Molecular Probes, MaβiTec, Gőttingen, Germany) was excited at 488 nm with an Argon Laser (Zeiss). For fluorescence detection a band pass filter (500–550 nm) was used. Setting the pinhole to an airy unit of 1 yielded a confocal slice of 2–3 µm. All fluorescence data is expressed as ΔF/F0 = 100 × (F – Fr)/(Fr – B), where F is the measured fluorescence signal at any given time, Fr is the average fluorescence from the scans preceding the stimuli, and B is the average value of the background fluorescence of a region in the scan field that does not contain any part of the dye filled cell. Regions of interest (ROIs) were chosen off line after image acquisition. Two different ROIs containing the membrane shell of the cell and the cytosolic shell were applied to each cell. For image analysis the Zeiss LSM 510 Meta Software was used, data analysis was performed off line using Microsoft EXCEL (Microsoft, Seattle, WA, USA) and Sigma Plot (SPSS, Chicago, IL, USA).

If not otherwise stated, values represent Mean± standard error of the mean (SEM). To test significance, Student's t-test was performed and differences were considered statistically significant if p < 0.05.

Quantitative RT-PCR

Real-time quantitative reverse transcription-PCR (RT-PCR) was performed by using the Roche Light Cycler system (Roche, Basel, Switzerland). Briefly, 16-µg samples of total RNAs isolated from mouse hippocampi were prepared with a phenol-guanidine isothiocyanate reagent (Trizol; Invitrogen, Karlsruhe, Germany) and cleaned by the use of RNeasy columns (Qiagen, Hilden, Germany). From 16-µg samples of total RNA, first-strand cDNAs were synthesized with SuperScript II (Invitrogen) and the T7-(dT24) primer (MWG, Ebersberg, Germany), followed by the generation of double-stranded cDNAs. Two microliter of diluted cDNA (1 : 10) were subjected to further PCR cycles, which were done with Faststart DNA Master SYBR Green I (Roche). The PCR conditions were 95°C for 10 min for a hot start, followed by denaturing at 95°C for 10 s, annealing at 57°C for 15 s, and extension at 72°C for 10 s for 50 cycles. The beta-actin gene was used as a general housekeeping gene to normalize target gene mRNA expression levels. Primer sets for beta-actin and target genes were chosen from published studies. Care was taken to select primers that bound to exon-intron boundaries or spanned exon-exon splice sites to avoid amplification of contaminating traces of genomic DNA. The primer sets used for PCRs were as follows: for beta-actin 5′-AACCCTAAGGCCAACCGTGAAAAG-3′ and 5′-CTAGGAGCCAGAGCAGTAATCT-3′, for Cacna1C 5′-CGCAGCGTAAGGATGA-3′ and 5′-GCCCTTCGACCTAGAG-3′; for Cacna1d 5′-ACTGAGAAACCGCTGT-3′ and 5′-AGACCTAATGTAAGTCTCGT-3′, for Cacnb2 5′-TCTGCTTAGCCGGACT-3′ and 5′-GGTTATGCTCGCGGTG-3′, for Cacnb3 5′-CACAACTTGCCAAGACC-3′ and 5′-CCAGTAAACCTCTAAGTATTC AG-3′, and for Cacnb4 5′-AGCTTAGCGGAAGTACAA-3′ and 5′-GCTACTGCTCGTGTGG-3′. Relative transcriptional expression levels of the target genes were generated by a relative quantification method as recommended by Roche (technical note LC 13/01).

Western blot analysis

Western blotting was carried out using standard methods. Briefly, brain homogenates were separated on SDS-PAGE, blotted on a PVDF membrane (Millipore, Schwalbach, Germany) and probed with Cav1.2 and β2 specific antibodies (Moosmang et al. 2003). Equal loading of slots was ascertained by the use of a polyclonal Erk1/2 (Upstate, Dundee, UK) antibody. Antibodies were visualized by the ECL system (NEN, Montreal, Canada).


  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Electrophysiology of AHP currents

To further investigate the phenotype of impaired sIAHP in hippocampal Prnp–/– CA1 neurons obtained from intracellular recordings in current-clamp mode (Colling et al. 1996), AHP currents were analyzed exploiting patch-clamp technique in whole-cell voltage-clamp mode. Figure 1 shows typical AHP traces of a Prnp+/+ CA1 pyramidal neuron (Fig. 1a) compared to littermate Prnp–/– (Fig. 1b). Examining the decay of the IAHP current revealed a tau of 83 ± 8 ms for Prnp–/– (n = 19) and 85 ± 7 ms for Prnp+/+ (n = 17); the decay of IAHP was unchanged. Similarly, no statistical difference in sIAHP decay between Prnp–/–(1.2 ± 0.4 s; n = 16) and Prnp+/+ (1.5 ± 0.4 s; n = 17) was observed. The decay values are in the range of previously published data for mouse CA1 hippocampal neurons (Krause et al. 2002). The IAHP current amplitude was not found to be significantly altered in Prnp–/–(143 ± 12 pA; n = 31) compared to Prnp+/+ (162 ± 15 pA; n = 29) cells (Fig. 1c). However, hippocampal Prnp–/– CA1 neurons displayed significantly reduced sIAHP amplitude (Prnp–/–: 7.7 ± 1 pA, n = 31; Prnp+/+: 14.3 ± 2.5 pA, n = 29; Fig. 1d), consistent with previous results from intracellular recordings in current-clamp mode (Colling et al. 1996; Mallucci et al. 2002; Asante et al. 2004).


Figure 1.  Reduced slow afterhyperpolarization (sIAHP) in Prnp–/– mouse hippocampal CA1 pyramidal neurons. (a, b) Representative traces obtained from a Prnp+/+ neuron (+/+, a) and a neuron from a littermate Prnp–/– mouse (–/–, b) showing initial IAHP directly upon the depolarization artifact, followed by sIAHP current. AHP currents were elicited by a 200 ms depolarization step from − 60 mV to +20 mV, as indicated. (c, d) IAHP and sIAHP current amplitudes obtained from wild-type (+/+, n = 29) and Prnp–/– CA1 neurons (–/–, n = 31). No significant differences were observed in IAHP current amplitudes between wild-type and Prnp–/– CA1 neurons (c), however, sIAHP current amplitudes (d) were found to be significantly reduced in Prnp–/– neurons (n.s. p > 0.05, *: p < 0.05; Student's t-test). (c, d) Box plot diagrams contain 10, 25, 75 and 90% percentiles; red/grey bar: mean; black bar: median.

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Next, intracellular Ca2+ was measured exploiting the same stimulation protocol as for evoking AHP currents. In whole-cell mode dye diffusion into hippocampal CA1 neurons was allowed for 15 min. The intracellular solution additionally contained Calcium Green-5N (CG-5N). Figure 2a shows a typical CG-5N filled hippocampal CA1 neuron. Two representative Ca2+ traces of Prnp–/– (open circles) and Prnp+/+ (filled circles) neurons are depicted in Fig. 2(b). Comparison of littermate Prnp–/– and Prnp+/+ hippocampal CA1 neurons revealed significantly reduced Ca2+ amplitudes in Prnp–/– cells (p < 0.05; Fig. 2c). No difference was found analyzing the decay time constant of the Ca2+ trace between littermate wild-type and PrPC–deficient mice (Fig. 2d).


Figure 2.  Altered free Ca2+ concentration dynamic in the soma of PrPC-deficient CA1 neurons. (a) Example of a dye filled CA1 neuron in an acute hippocampal slice preparation of an 18 day old mouse. (b) Representative Ca2+ traces obtained from the soma of Prnp+/+ (filled circles) and Prnp–/– (open circles) hippocampal CA1 neurons. Cells were depolarized via patch pipette from a holding potential of − 60 mV to + 20 mV for 200 ms, at the time point indicated (arrow). Box plot diagrams show the maximal amplitudes (c) and the decay time constants (d) of Ca2+ traces obtained from 31 wild-type and 29 Prnp–/– neurons. The mean Ca2+ amplitudes were found significantly reduced in Prnp–/– hippocampal CA1 neurons compared to littermate control measurements (+/+: 13.6 ± 1.1%, n = 31; −/–: 9.8 ± 1%, n = 32). No differences were observed between the decay time constants (+/+: 3.2 ± 0.3 s, n = 31; −/–: 3.4 ± 0.3 s, n = 29). Box plot diagrams contain 10, 25, 75 and 90% percentiles; red bar: mean; black bar: median (*p < 0.05; n.s. p > 0.05 Student's t-test).

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In order to determine whether the Ca2+ influx through the plasma membrane is predominantly affected by loss of PrPC, we analyzed the confocal images in more detail. According to Eilers et al. (1995),Fig. 3(a) we compared the Ca2+ transient at the submembrane region to the cytosolic shell (Fig. 3a). Figures 3(b and c) depict representative submembrane and cytosolic shell Ca2+ traces of wild-type and PrPC–deficient hippocampal CA1 neurons, respectively, after a depolarizing voltage step of 200 ms to + 20 mV was applied. The fast component of the Ca2+ trace measured in the submembrane shell exhibited a decay of 1.9 ± 0.1 s in Prnp+/+ cells, whereas the cytosolic shell decay time constant was 3.4 ± 0.3 s. The fast component represents Ca2+ influx via VGCCs and the diffusion of Ca2+ to more central somatic sites, whereas the slow component is likely to be due to Ca2+ removal, via Ca2+ pumps and Na+– Ca2+ exchange together with the binding of Ca2+ to high affinity endogenous buffers (Eilers et al. 1995). The area under the curve, or charge transfer (CT), of submembrane and cytosolic shell Ca2+ transients was measured (Figs 3d and e). Cytosolic shell charge transfer value (CTCyt) of Prnp–/– (4.8 ± 0.7, n = 30; 4 animals) was significantly reduced in comparison to Prnp+/+ (7.9 ± 0.8; n = 31, 4 animals; p < 0.005). Similarly, submembrane shell charge transfer value (CTMem) was decreased from 12.3 ± 1.4 (Prnp+/+, n = 31, 4 animals) to 8.4 ± 1.2 (Prnp–/–, n = 30, 4 animals; p < 0.05).


Figure 3.  Both submembrane and cytosolic [Ca2+]i are affected by loss of PrPC. (a) Typical confocal images of a dye-loaded hippocampal CA1 neuronal soma, at different time points as indicated. Ca2+ concentration was analyzed in a submembrane (Mem) and a cytosolic (Cyt) shell. Following a 200 ms depolarization, a circular membrane-associated signal can be detected at 1.2 s. Images at later time points (2.2 and 7.8 s) show a predominantly cytosolic signal. (b, c) Representative traces of Prnp+/+ (+/+) vs. Prnp–/– (–/–) CA1 neurons comparing the submembrane and cytosolic region of interest. (d, e) Ca2+ charge transfer (CT) derived from the area under the curve of wild-type (+/+ n = 31) vs. PrP knock out (–/–; n = 30) neurons. Prnp–/– mouse CA1 hippocampal neurons display significantly reduced CTMem and CTCyt (*p < 0.05; **p < 0.005 Student's t-test). (d, e) Box plot diagrams contain 10, 25, 75 and 90% percentiles; red/grey bar: mean; black bar: median.

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The reduced CTMem value argues for an impaired Ca2+ flux via VGCCs in Prnp–/– mice, since VGCCs solely contribute to the initial Ca2+-influx measured after a depolarizing voltage step.

Electrophysiological characterization of VGCCs

To analyze the possible contribution of VGCCs to reduced sIAHP currents and Ca2+ charge transfer of Prnp–/– hippocampal CA1 neurons I-V relationships of Prnp–/– and Prnp+/+ neurons were compared. Figures 4(a and b) insets show representative currents of Prnp+/+ (Fig. 4a inset) and Prnp–/– (Fig. 4b inset) evoked by 200 ms depolarization from a holding potential of −70 to −20 mV. Application of nifedipine (10 µm), known to block l-type VGCCs, reliably reduced the current. The evoked currents always displayed an inactivating component (Fig. 4a, ΔX) that was reduced by nifedipine, indicating that it is partly mediated by l-type Ca2+ channels (Avery and Johnston 1996). Figure 4a displays the mean I-V relationship of Prnp+/+ CA1 neurons with or without application of nifedipine (n = 10 cells, 3 animals). Analogously Fig. 4b displays the mean I-V relationship of equally treated Prnp–/– CA1 neurons (n = 10 cells, 3 animals).

Comparing the mean maximum amplitude values of Prnp+/+ (1555 ± 157 pA; n = 9, 3 animals) and Prnp–/– (1141 ± 101 pA; n = 9, 3 animals) hippocampal CA1 neurons revealed a significant reduction of the maximum Ca2+ current amplitude of Prnp–/– neurons (Fig. 4c). Nifedipine partially reduced the maximum Ca2+ current amplitude of Prnp+/+ (865 ± 170 pA; n = 9, 3 animals) and Prnp–/– (946 ± 60 pA; n = 9, 3 animals) neurons to a comparable absolute value (Fig. 4c). It is important to note that in the presence of nifedipine the percent reduction of the maximum current of Prnp+/+ CA1 neurons (44 ± 10%) is significantly higher in comparison to Prnp–/– CA1 neurons (17 ± 2%) (Fig. 4d).

Investigating the nifedipine-sensitive inactivating component of the Ca2+ currents revealed a significantly smaller inactivating component in Prnp–/–(10.8 ± 0.7%) CA1 neurons compared to Prnp+/+ (21.2 ± 4.5%) (Fig. 4e). Administration of nifedipine slightly diminished the inactivating component of Prnp–/– CA1 neurons to 7.7 ± 1%, whereas the inactivating component of Prnp+/+ CA1 neurons was decreased by 50% to 10.6 ± 1.9% (Fig. 4e). These results argue for diminished Ca2+ currents through nifedipine-sensitive l-type VGCCs in Prnp-/– hippocampal CA1 neurons.

Expression of l-type VGCCs

To analyze whether the decreased Ca2+ current through l-type VGCCs of Prnp–/– CA1 neurons may be attributed to a reduced number of VGCCs, quantitative RT-PCR was performed to measure the expression levels of voltage gated l-type Ca2+ channels that are known to mediate Ca2+ influx in hippocampal neurons (Tanabe et al. 1998). As candidate genes we chose the l-type Ca2+ channels Cacna1c and Cacna1d. These channels are confidently expressed in somatodendritic compartments of hippocampal CA1 neurons (Westenbroek et al. 1990). Comparing the expression levels of Prnp+/+ with Prnp–/– hippocampi concerning Cacna1c (+/+1.8 ± 0.6, −/– 1.82 ± 0.4; p > 0.05; n = 3) and Cacna1d (+/+: 0.96 ± 0.16, −/–: 0.84 ± 0.24; p > 0.05; n = 3) no significant differences were detected (Fig. 5a). Furthermore the mRNA levels of regulatory β subunits that control and regulate the membrane incorporation of pore forming α-subunits were examined (Catterall 2000; Dalton et al. 2005). Cacnb2–4 were chosen as candidate genes. No significant differences could be detected comparing the expression levels of these genes: Cacnb2 (+/+1.09 ± 0.2, −/– 0.95 ± 0.2; p > 0.05; n = 3), Cacnb3 (+/+2.9 ± 0.5, −/– 3.3 ± 0.5; p > 0.05; n = 3), Cacnb4 (+/+0.23 ± 0.03, −/– 0.25 ± 0.02; p > 0.05; n = 3) (Fig. 5a).


Figure 5.  Protein and mRNA expression levels of pore forming and regulatory β subunits comparing Prnp–/– and Prnp+/+ hippocampal tissue. (a) Comparing relative mRNA expression levels of Cacna1c, Cacna1d and Cacnb2–4 genes for wild-type (+/+) and knock out (–/–) hippocampal tissue no significant differences were found (n = 3, +/+ and −/– hippocampal hemispheres). (b) Western blot analysis of the corresponding hippocampal hemispheres comparing protein expression levels of CaV1.2 (α1C) and β2 for Prnp+/+ and Prnp–/–. MAPK was used as loading control. While CaV1.2 levels were found to be unaltered, the protein expression of the β2 subunit was increased in all three Prnp–/– hippocampi.

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Western blot analysis of l-type VGCCs α1C pore forming unit showed no difference concerning the protein expression level of this protein (n = 3 Prnp+/+ and Prnp–/– hippocampal hemispheres). When comparing the protein expression level of β2 subunits of three wild-type and Prnp–/– hippocampi an increase in β2 subunit expression was detected in all three knock out mice (Fig. 5b).


  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The present study aimed to determine whether loss of PrPC directly affects sIAHP or if sIAHP alterations represent secondary effects due to alterations in [Ca2+]i. Summarizing, the previously described phenotype of impaired sIAHP in Prnp–/– mice (Colling et al. 1996; Mallucci et al. 2002; Asante et al. 2004) was confirmed using the whole-cell patch-clamp technique. Analyzing Ca2+ charge transfer following a depolarization protocol used to elicit AHP currents revealed reduced Ca2+ transients in hippocampal Prnp–/– CA1 neurons. Reduced Ca2+ charge transfer in hippocampal Prnp–/– CA1 neurons, as described here for the first time, indicates that the alteration in sIAHP is most likely not a direct consequence of loss of PrPC on K+-efflux. In fact, decreased cytosolic free Ca2+ concentration, on which sIAHP currents mainly depend, might be the reason for impaired sIAHP. These results obtained from CA1 hippocampal neurons stand in line with our previous findings derived from cerebellar Prnp–/– neurons, namely cultured cerebellar granule cells as well as purkinje cells in slice preparations. Microfluorometric measurements using Fura-2 showed a reduced Ca2+ rise following cell depolarization in both cell types (Herms et al. 2000; Herms et al. 2001). Also a very recent study using Ca2+-sensitive aequorin chimeras in Chinese hamster ovary (CHO) cells revealed convincing evidence that loss of PrPC modulates the intracellular free Ca2+ concentration (Brini et al. 2005). The fact that different types of neurons display similar phenotypes with regard to the intracellular free Ca2+ concentration indicates that loss of PrPC affects basal mechanisms of Ca2+ homeostasis. In general, three mechanisms have to be considered as to how PrPC deficiency might modulate intracellular free Ca2+ concentration following depolarization: Firstly, loss of PrPC may modulate Ca2+ influx from the extracellular space via voltage gated calcium channels. Secondly, PrPC deficiency may alter Ca2+-induced Ca2+ release from intracellular stores, or alter filling of these stores. Thirdly, lack of PrPC may result in enhanced removal of Ca2+ from the cytosol, for instance by increased Ca2+ buffering within the cytoplasm, extrusion of Ca2+ into organelles or through the plasma membrane.

Judging from our confocal microfluorimetric Ca2+ measurements, loss of PrPC primarily affects Ca2+ influx through the plasma membrane. In order to study this more directly we performed patch-clamp whole-cell recordings in hippocampal slice preparations. Indeed, we found a reduced Ca2+ influx following the activation of VGCCs in Prnp–/– CA1 neurons. Moreover, nifedipine (a blocker of l-type VGCCs) was found to reduce the Ca2+ influx about twice as strong in wild-type hippocampal neurons compared to Prnp–/– neurons, reaching similar levels. These observations strongly indicate that the Ca2+ influx through l-type VGCCs is altered in hippocampal neurons lacking PrPC. In line with this suggestion stands the observation that Ca2+-dependent inactivation of l-type VGCCs (Avery and Johnston 1996) is significantly more pronounced in wild-type neurons than in PrPC lacking cells (Fig. 4e). In order to analyze, whether the expression of l-type VGCCs is affected by loss of PrPC, we studied both mRNA and protein expression levels of l-type channels by quantitative RT-PCR and Western blot. Comparison of hippocampal mRNA expression levels from wild-type and Prnp–/– mice did not reveal any significant differences in VGCCs. Moreover, Western blot analysis did not reveal quantitative differences in the pore forming α1C subunit in hippocampal lysates. Unexpectedly, however, we observed an enhanced expression of the β2 subunit in hippocampal tissue of Prnp–/– mice. The β2 subunit is known to modulate the function of VGCCs via its critical role in transporting α1 subunits to the plasma membrane. This process is regulated by phosphatidylinositol 3-kinase (PI3K)/Akt-dependent phosphorylation of the β2 subunit (Viard et al. 2004). Interestingly, our previous studies showed strong evidence that PrPC activates a PI3K-mediated signaling cascade resulting in improved cell survival (Vassallo et al. 2005). In addition, a reduced PI3K activity in Prnp–/– mice brain lysates was observed (Vassallo et al. 2005). A reduced PI3K activity could indeed be a possible explanation for the enhanced expression of the β2 subunit in the brain of mice lacking PrPC. A reduced transport of α1 subunits to the plasma membrane and consequently a reduced Ca2+ influx through l-type VGCCs may induce an up-regulation of the β2 subunit as a compensatory phenomenon. Since the PI3K-mediated modulation of VGCCs via Akt is known to be critically involved in preventing apoptotic cell death (Blair et al. 1999), this hypothesis may further explain the enhanced vulnerability of cells lacking PrPC to oxidative stress (Brown et al. 1997b; Kim et al. 2004; Nishimura et al. 2004) or the increased size of infarcts induced in Prnp–/– mice (McLennan et al. 2004; Spudich et al. 2005; Weise et al. 2006). Both phenomena have been linked to reduced PI3K/Akt activation (Vassallo et al. 2005; Weise et al. 2006). We would like to underline that the above suggested mechanisms are part of a working hypothesis and require further investigation.

A role for PrPC in enhancing Ca2+ influx through VGCCs was initially suggested based on microfluorimetric measurements on synaptosomal preparations (Whatley et al. 1995). However, more recent patch-clamp studies on cerebellar granule cells revealed that the application of recombinant PrP depresses, rather than enhances, the Ca2+ influx through l-type VGCCs (Korte et al. 2003). Interestingly, a depression of VGCCs was also observed in cell lines infected with PrPSc (Sandberg et al. 2004). Whether this is due to a loss of function of PrPC in modulating VGCCs or caused by a different independent mechanism is a matter of speculation.

In conclusion, PrPC depletion in hippocampal CA1 neurons affects the Ca2+-activated potassium channels underlying sIAHP most likely via a reduction of Ca2+ influx through L-Type VGCCs. Since Ca2+ influx via l-type VGCCs is known to control several basal cellular functions in neurons and other cell types, the broad variety of Prnp–/– phenotypes may be at least in part related to an impaired function of L-Type VGCCs.


  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

This work was supported by the Bayerische Forschungsverbund ForPrion and the Deutsche Forschungsgemeinschaft (HE 2328/71). We kindly thank Neville Vassallo for helpful comments on the manuscript, Wei Xiang and Ramona Heptner to provide assistance with performing Quantitative RT-PCR analysis.


  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
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