HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 282, Issue 2, 1119-1127, January 12, 2007

This Article Abstract Full Text (PDF) All Versions of this Article: 282/2/1119    most recent M605066200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Priller, C. Articles by Herms, J. Search for Related Content PubMed PubMed Citation Articles by Priller, C. Articles by Herms, J. Social Bookmarking                      What's this?

Mutant Presenilin 1 Alters Synaptic Transmission in Cultured Hippocampal Neurons^*

Christina Priller, Ilse Dewachter, Neville Vassallo^¶, Sandra Paluch, Claudia Pace, Hans A. Kretzschmar, Fred Van Leuven, and Jochen Herms¹

From the Department of Neuropathology, Ludwig-Maximilians-Universität Munich, Feodor-Lynen-Strasse 23, 81377 München, Germany, the Experimental Genetics Group, KULeuven-Campus Gasthuisberg, B-3000 Leuven, Belgium, and the ^¶Department of Physiology and Biochemistry, University of Malta, Msida MSD06, Malta

Received for publication, May 26, 2006 , and in revised form, October 31, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mutations in presenilins are the major cause of familial Alzheimer disease, but the precise pathogenic mechanism by which presenilin (PS) mutations cause synaptic dysfunction leading to memory loss and neurodegeneration remains unclear. Using autaptic hippocampal cultures from transgenic mice expressing human PS1 with the A246E mutation, we demonstrate that mutant PS1 significantly depressed the amplitude of evoked -amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid and N-methyl-D-aspartate receptor-mediated synaptic currents. Analysis of the spontaneous miniature synaptic activity revealed a lower frequency of miniature currents but normal miniature amplitude. Both alterations could be rescued by the application of a -secretase blocker. On the other hand, the application of synthetic soluble A₄₂ in wild-type neurons induced the PS1 mutant phenotype on synaptic strength. Together, these findings strongly suggest that the expression of mutant PS1 in cultured neurons depresses synaptic transmission by causing a physical reduction in the number of synapses. This hypothesis is consistent with morphometic and semiquantitative immunohistochemical analysis, revealing a decrease in synaptophysin-positive puncta in PS1 mutant hippocampal neurons.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Alzheimer disease is an age-related neurodegenerative disorder characterized clinically by a progressive deterioration in memory and other cognitive abilities. Neuropathological hallmarks include a progressive loss of neurons, synaptic degeneration, and deposition of amyloid plaques and neurofibrillary tangles in characteristic brain regions. Dominantly inherited mutations in presenilin (PS)² 1 and 2 are the primary cause of familial Alzheimer disease (1). The exact pathogenic mechanism by which PS mutations cause memory loss and neurode-generation, however, remains unclear.

Presenilins are integral components of a multiprotein protease complex termed -secretase that is responsible for the intramembranous cleavage of the amyloid precursor protein (APP) and the Notch receptors (2, 3). Familial Alzheimer disease-linked PS mutations selectively enhance the production of the amyloidogenic 42-residue -amyloid peptide (A₄₂), often at the expense of the less amyloidogenic A₄₀ generation (4, 5), suggesting a toxic gain-of-function pathogenic mechanism. In mammalian cells mutant forms of PS1 produce reduced levels of the intracellular domains of Notch and APP, which can function as transcriptional activators (4-6).

Neuropathological studies have indicated that synaptic abnormalities may represent the proximate cause of dementia in Alzheimer disease. Synaptic loss correlates better with the pattern and severity of cognitive impairment than formation of amyloid plaques or neurofibrillary tangles (7). In addition, synaptic and dendritic loss precedes neuronal loss and is most prominent in limbic regions corresponding to the clinical deficits (7, 8).

Abnormalities in learning and synaptic function have been observed in transgenic mice expressing mutant APP or mutant PS1. However, the results obtained by different groups using different animal models are very heterogeneous. For instance, initial studies examining long-term potentiation, a widely used model for synaptic plasticity, reported a reduction of long-term potentiation in transgenic mice with A accumulation (9-11), indicating that an alteration in synaptic plasticity may underlie memory deficits in transgenic Alzheimer disease mice. However, later studies were not able to reproduce these findings fully; rather, they pointed to an impairment of basal synaptic transmission in these mice (12-18). In transgenic mice expressing mutant PS1 alone, long-term potentiation was even found to be facilitated (19-21). The heterogeneity of the findings in different experimental settings may be due to the possibility that mutant PS1 exerts pathogenic effects not only by increasing A₄₂ production but also by influencing synapse formation or synaptic plasticity (22, 23).

Here we aimed to analyze pre- and postsynaptic mechanisms of synaptic vesicle release in mutant PS1 transgenic mice in detail by studying autapses in single-grown hippocampal neurons. Autapses are synapses between the axon and dendrites of the same neuron. These structures are built when neurons are cultured separately, a condition that is achieved by a special coating of the cell culture wells. The main advantage compared with conventional preparations is that these monosynaptic cultures ensure that the release from the same set of synapses is monitored regardless of the stimulation method and in absence of complex polysynaptic circuitry. Our results essentially show that mutant PS1 strongly affects both the amplitude of evoked excitatory currents as well as the frequency of spontaneous excitatory synaptic currents by decreasing the number of functional synapses.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mice—The parent PS1 transgenic mice (wild-type human PS1 and human mutant PS1[A246E]) were generated by microinjection of mini-gene constructs containing PS1 cDNA in a mouse Thy1 gene cassette into prenuclear embryos as described (10, 24). Transgenic strains were maintained on the FVB/N background and back-crossed to obtain homozygous mice for either wild-type (PS1wt) or mutant human PS1 (PS1mut). FVB/N mice were used as non-transgenic (Non-tg) controls.

Cell Culture—Microisland cultures of mouse hippocampal neurons were prepared and maintained as described (25, 26). Briefly, astrocyte feeder wells enriched in type 1 astrocytes (>95%) were first prepared from postnatal day 1 mouse hippocampal tissue. After dissection and removal of the choroid plexus and meninges, the material was dissociated and trypsinized. Astrocytes were grown in culture media consisting of Dulbecco's modified Eagle's medium with 4.5 g/liter glucose (Pan, Aidenbach, Germany), 5% fetal bovine serum (Pan), 100 IU/ml penicillin/streptomycin (Invitrogen), N2 supplement (Invitrogen), Glutamax (Invitrogen), and Mito⁺ serum extender (BD Bioscience) at their indicated concentrations. Astrocytes were grown at 37 °C in a 10% CO₂-enriched atmosphere. Once a monolayer was achieved in the flasks, cells were trypsinized and replated onto microdot-coated glass coverslips. Hippocampi were dissected from newborn mice pups and digested in Dulbecco's modified Eagle's medium solution containing 0.2 mg/ml cysteine, 100 mM CaCl₂, 50 mM EDTA, 10 units/ml papain (Roche Applied Science) at 37 °C. Digestion was stopped after 60 min, and tissue was gently triturated using fire-polished Pasteur pipettes. Cell suspensions diluted to 5–8 x 10⁴ cells/ml were then added to the prepared astrocyte feeder wells (see above). To prevent excessive proliferation of astrocytes, the anti-mitotic agent 5-fluoro-2'-deoxyuridine (8.1 mM) was applied to the cultures on the day after neuronal plating. For the -secretase inhibitor experiment, autaptic cultures were incubated with 1 µM N-[N-(3,5-difluorophenacetyl-L-alanyl)]-S-phenylglycine t-butyl Ester (DAPT, Calbiochem) 48 h before the measurements. For the A incubation studies, freshly diluted monomeric A₄₂ (100 nM, Bachem, Switzerland) was added to the autaptic cultures,and electrophysiological measurements were carried out after 48 h.

Electrophysiology—For each experiment approximately equal numbers of cells from the respective genotypes were measured in parallel and blindly on the same day in vitro (18–20 days in vitro (DIV). Whole-cell recordings were performed using an EPC 9 amplifier (HEKA Elektronic, Lambrecht, Germany). The standard extracellular medium contained 140 mM NaCl, 2.4 mM KCl, 10 mM HEPES, 10 mM glucose, 4 mM CaCl₂, 4 mM MgCl₂, and 15 µM bicuculline, 300 mosM, pH 7.3. NMDA-excitatory post-synaptic currents (EPSCs) were measured at 4 mM external Ca²⁺ in the presence of 5 µM 6-cyano-7-nitroquinoxaline-2,3-dione, 10 µM glycine and in the absence of external Mg²⁺. Internal pipette recording solution consisted of 10 mM NaCl, 125 mM potassium gluco-nate, 1 mM EGTA, 4.6 mM MgCl₂, 4 mM Na₂ATP, 15 mM creatine phosphate, and 20 mM units/ml phosphocreatine kinase, 300 mosM, pH 7.3. Data were filtered at 2 kHz and acquired at 10 kHz using Pulse 8.5 software (HEKA Elektronic). Patch electrodes had resistances of 3–4 megohms. Series resistance and capacity of the electrodes were compensated; only recordings with access resistance below 15 megohms and with an initial membrane potential <-50 mV (measured in the current-clamp modus shortly after cell access was obtained) were included in the analysis. Access resistance and cell capacities (5–25 picofarads) were compensated. Cells were voltage-clamped at -70 mV, except for 1-ms depolarizations to 0 mV to elicit an action potential-like stimulus. The rise time constant (10–90%) of the synaptic currents was determined by fitting a linear function to the activating current, and the decay time constant was determined by fitting a single exponential function to the current decay. Detection of miniature EPSCs was usually performed for at least 60 s in the presence of 200 nM tetrodotoxin (Biotrend, Köln, Germany). Data were analyzed with a template detection program (Axograph 4.1, Axon Instruments, Sunnyvale, CA). Threshold for detection was set to 3.5 times the base-line S.D. Captured miniature EPSCs (mEPSCs) of individual cells were averaged to determine mean amplitude and charge per cell. Application of a hypertonic solution for pool size determination was made by pulsed application of 500 mM sucrose to the extracellular solution for 4 s using a fast perfusion system (Warner Instruments, Hamden, CT). Evoked responses and responses to hypertonic sucrose solutions were always recorded successively from the same cell. The readily releasable vesicle pool (RRP) per cell was calculated by dividing the charge of the transient sucrose response with the mean mEPSC charge of the same cell, if available. To further determine the mean vesicular release probability, the number of vesicles released by a single action potential-like stimulus was divided by the number of vesicles in the RRP. Recordings and data analysis were performed blindly on each sample.

Immunocytochemistry—Hippocampal cultures of 18–20 DIV were used for immunocytochemistry. For fluorescent microscopic determinations of synaptic density in neurons, cultures were fixed for 10 min with 4% paraformaldehyde, blocked, and permeabilized with phosphate-buffered saline containing 2% bovine serum albumin, 10% goat serum (Dako Hamburg, Germany), 0.3% Triton X-100 (Calbiochem) for 30 min. Cells were incubated with rabbit polyclonal anti microtubule-associated protein 2 (1:100, Chemicon, Hampshire, UK) for 60 min, washed with phosphate-buffered saline, and incubated with fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (1:100) for another 60 min. After another washing step, fluorescent-labeled mouse monoclonal anti-synaptophysin (1:1000, Synaptic Systems, Göttingen, Germany) was added, incubated for 60 min, and finally fixed in fluorescent mounting medium (Dako). Images were acquired on an Axiovert 200M microscope (Zeiss, Oberkochen, Germany) with a 63x LD Achroplan objective and Axiovision 4.5 software (Zeiss). For quantification of the total dendritic length and total amount of synapses, analySIS software (Soft Imaging System, Münster, Germany) was used. The summary graph represents four independent experiments using different culture preparations from PS1mut animals and PS1wt controls. All chemicals were obtained from Sigma-Aldrich unless otherwise noted.

Immunohistochemistry—Six hemispheres (3 of each geno-type) of 30-day- and 3-month-old wild-type human PS1, human mutant PS1[A246E], and controls were removed and fixed in 4% paraformaldehyde overnight. Coronal 50-µm free floating sections were prepared with a VT 1000S vibratome (Leica, Wetzlar, Germany) and immunostained with anti-synaptophysin and Cy^TM3-conjugated streptavidin. Briefly, sections were placed in phosphate-buffered saline, blocked with antibody diluent (Ventana Medical Systems, Inc., Illkirch, France), and then incubated overnight at 4 °C with the polyclonal antibody against synaptophysin (1:300, Chemicon). This step was followed by incubation in biotinylated swine anti-rabbit IgG (1:150, Dako), and finally, the sections were fluorescence labeled with Cy^TM3-conjugated streptavidin (1:25, Dianova, Hamburg, Germany). The sections were mounted on precoated glass slides (Super-frost-plus, Menzel, Braun-schweig, Germany) and covered (Coverquick, Labonord, Templemars, France). All sections were processed simultaneously under the same conditions. The immunofluorescent labeling protocol was repeated three times to assess the reproducibility of results. Immunolabeled brain sections were viewed with a Zeiss 40x/0.8 IR Achroplan objective on a Zeiss Axioskop 2FS mot LSM510 confocal microscope.

Cy3 was excited with a HeNe Laser at 543 nm, and the emitted fluorescence was detected with a LP560 (longpass 560 nm) emission filter. Frames of 230 x 230-µm size were scanned with 0.22 µm/pixel, and the pinhole diameter was set to an area unit of 1. The z-distance of each z-stack ranged from 10 to 15 µm and consisted of 20–30 0.5-µm-spaced images.

Synaptophysin levels were assessed as described previously (27)in the stratum radiatum and stratum moleculare of the hippocampus in three coronal sections per animal. Two serial optical z-stacks (one for each marker) of the synaptophysin-labeled structures were collected from each section. All series of z-stacks from each section were maximum intensity projected and acquired under standard conditions maintaining the same gain, aperture, and black level settings. This method of analysis was previously evaluated by Masliah et al. (28, 29). The digitized images were then analyzed regarding the signal intensity and density (covered area of immunohistochemistry-positive objects) of synaptophysin-labeled structures (PictureAnalyzer Software, Bjarne Krebs, Zentrum für Neuropathologie und Prion Forschung, Munich, Germany). A threshold was set so that all objects with an intensity value below the threshold were set equal to the background value. For each mouse, the values obtained for the stratum radiatum and stratum moleculare of the hippocampus were averaged.

Western Blot—For measurement of the expression of synaptophysin in hippocampus, coronal brain sections were collected on glass slides and Nissl stained. The hippocampal region was then separately lysed and subjected to SDS-PAGE as previously reported (30). After electrotransfer of proteins to the polyvinylidene difluoride membrane, immunoblotting was followed by the incubation with primary antibody against synaptophysin (1:5000, Chemicon) and -actin (1:5000, actin I-19, Santa Cruz Biotechnology, Heidelberg, Germany) overnight. The visualization was performed with a secondary antibody coupled to alkaline phosphatase (polyclonal goat anti-mouse and anti-rabbit immunoglobulin/alkaline phosphatase, Dako) and chromo-gene nitro blue tetrazolium salt with 5-bromo-4-chloro-3-indolylphosphate. Western blots were scanned with a high resolution flatbed scanner, and the density of bands was determined using Total Lab Version 2.01 software (Nonlinear Dynamics, Newcastle-upon-Tyne, UK).

View larger version (25K): [in this window] [in a new window]   FIGURE 1. Evoked neurotransmitter release in Non-tg neurons and in neurons expressing PS1wt or PS1mut. A, representative AMPA-EPSC traces from Non-tg, PS1wt, and PS1mut neurons. B, AMPA-EPSC amplitude in neurons from Non-tg (n = 19), PS1wt (n = 30), and PS1mut (n = 41) mice. C, representative NMDA-EPSC traces from Non-tg, PS1wt, and PS1mut neurons. D, NMDA-EPSC amplitude in neurons from Non-tg (n = 15), PS1wt (n = 32), and PS1mut (n = 27) mice (*, p < 0.05; **, p < 0.005).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Evoked Neurotransmitter Release in Non-tg, PS1wt, and PS1mut Neurons—Patch clamp recordings from single isolated neurons were used to assess possible defects in neurotransmitter release induced by mutant PS1. We concentrated our analysis on glutamatergic neurons, because they are much more abundant in autaptic cultures than in inhibitory cells producing -aminobutyric acid. We first tested the action potential-evoked neurotransmitter release. AMPA and NMDA synaptic responses were evoked by brief somatic depolarization (1-ms depolarization from -70mV holding potential to 0 mV) and measured as peak inward currents a few ms after action potential induction (Fig. 1, A and C). The amplitude of the evoked autaptic amplitudes of AMPA-EPSCs was reduced in PS1mut cells compared with neurons expressing human PS1wt and Non-tg mice (3.66 ± 0.46 nA for Non-tg, 3.24 ± 0.42 nA for PS1wt, and 2.16 ± 0.22 nA for PS1mut; p < 0.05; Fig. 1B). Similarly, NMDA receptor-mediated EPSCs, analyzed by blocking AMPA receptors by 6-cyano-7-nitroquinoxaline-2,3-dione (5 µM), revealed a significantly reduced amplitude in PS1 mutant cells when compared with cells from mice expressing human wild-type PS1 or from non-transgenic mice (2.14 ± 0.28 nA for Non-tg, 1.68 ± 0.19 nA for PS1wt, and 0.77 ± 0.08 nA for PS1mut; p < 0.05; Fig. 1D). No significant differences were observed in the kinetics (rise time and decay time constant ) of both AMPA and NMDA receptor-mediated EPSCs between PS1 mutant, PS1 wild-type, and non-transgenic neurons (Table 1).

Spontaneous Neurotransmitter Release in Non-tg, PS1wt, and PS1mut Neurons—To define whether the pre- or the postsynaptic side is affected by mutant PS1, we analyzed the amplitude and frequency of spontaneous mEPSC activity (mEPSCs; Fig. 2A). We found that the frequency of mEPSCs was significantly lower in PS1mut neurons than in hPS1 and non-transgenic cells (8.03 ± 1.52 s^-1 for Non-tg, 7.95 ± 1.53 s^-1 for PS1wt, and 3.02 ± 0.59 s^-1 for PS1mut neurons; p < 0.05; Fig. 2B). The mean mEPSC amplitude, however, was not affected (24.34 ± 0.87 pA for Non-tg, 23.36 ± 0.92 pA for PS1wt, and 25.06 ± 1.12 pA for PS1mut cells; p > 0.05; Fig. 2C), indicating that the responsiveness and number of postsynaptic receptors is not altered due to mutation of PS1. In addition, we calculated the number of vesicles released by an action potential-like stimulus by dividing the charge of evoked responses by that of a single mEPSC. For those neurons in which both EPSCs and mEPSCs were measured, the quantity of vesicles released during an EPSC was determined and found to be significantly lower in PS1mut neurons (526 ± 148 for Non-tg, 481 ± 111 for PS1wt, and 169 ± 68 for PS1mut cells; p < 0.05; Fig. 2D).

View larger version (34K): [in this window] [in a new window]   FIGURE 2. Spontaneous neurotransmitter release in Non-tg neurons and in neurons expressing PS1wt or PS1mut. A, representative mEPSC traces from Non-tg, PS1wt, and PS1mut neurons. B, mEPSC frequency in neurons from Non-tg (n = 22), PS1wt (n = 24), and PS1mut (n = 30) mice. C, mEPSC amplitude in neurons from Non-tg (n = 22), PS1wt (n = 24), and PS1mut mice (n = 30) mice. D, estimated EPSC vesicles for Non-tg (n = 9), PS1wt (n = 11), and PS1mut mice (n = 8)(*, p < 0.05).

To estimate whether the efficiency of the action potential-induced vesicle release is influenced by PS1 mutation, we analyzed the vesicular release probability (P_vr). We determined P_vr by dividing the number of vesicles released during an action potential by the number of vesicles in the RRP. We found that P_vr did not vary significantly between cells from non-transgenic (4.24 ± 0.89%), PS1wt (4.25% ± 0.67%), and PS1mut neurons (4.41 ± 1.37%; Fig. 3D). The P_vr values obtained are similar to the values obtained in other studies on autaptic hippocampal neurons (33). In summary, these results indicate that the content of presynaptic vesicles seems to be released efficiently and that the decrease in synaptic responses in PS1mut autapses can be attributed to the decrease in RRP.

View larger version (21K): [in this window] [in a new window]   FIGURE 3. Estimated RRP, RRP refilling, and release probability in Non-tg neurons and in neurons expressing PS1wt or PS1mut. A, representative raw traces of responses to hypertonic solution from Non-tg, PS1wt, and PS1mut neurons. Hypertonic solution was applied for 4 s as indicated by the black bar. B, estimated RRP vesicles for Non-tg (n = 26), PS1wt (n = 18), and PS1mut (n = 29) mice. C, average ratio of the second hypertonic sucrose response to the initial sucrose response measured 4 s before (the inset shows sample recording). D, bar diagram showing the average vesicular release probability (Pvr) of Non-tg (n = 9), PS1wt (n = 11), and PS1mut (n = 8) mice (*, p < 0.05).

View larger version (19K): [in this window] [in a new window]   FIGURE 4. Paired pulse ratio of Non-tg neurons and neurons expressing PS1wt or PS1mut. A, average EPSC amplitudes of Non-tg (n = 18), PS1wt (n = 24), and PS1mut (n = 40) neurons to different stimulus intervals at 1 mM [Ca2+]e normalized to the size of the first response. B, average EPSC amplitudes of Non-tg (n = 12), PS1wt (n = 16), and PS1mut (n = 18) neurons to different stimulus intervals at 4 mM [Ca2+]e normalized to the size of the first response.

Prolonged Repetitive and High Frequency Stimulation in Non-tg, PS1wt, and PS1mut Neurons—Repetitive stimulation gives insights into the nature of the release probability, P_r, and the size of the RRP in the central synapse (34). For prolonged repetitive stimulation we used a 10-Hz protocol, and for high frequency stimulation we used a 50-Hz protocol at 4 mM [Ca²⁺]_out (Fig. 5, A and C). There was no significant difference in the gradual reduction of the relative EPSC amplitude during repetitive stimulation at 10 Hz between Non-tg, PS1wt, and in PS1mut neurons (Fig. 5B). Similar properties were observed in evoked responses elicited by high frequency stimulation at 50 Hz (Fig. 5D).

-Secretase Inhibitor Enhances Evoked and Spontaneous Neurotransmitter Release in PS1mut Neurons—To determine whether the reduction of EPSC amplitude is caused by -secretase cleavage products, we blocked -secretase activity in PS1mut neurons by the addition of 1 µM DAPT. DAPT-treated PS1mut neurons showed significantly higher AMPA-EPSC responses compared with untreated PS1mut neurons (1.83 ± 0.26 nA for control PS1mut and 2.65 ± 0.28 nA for DAPT-treated PS1mut; p < 0.05; Fig. 6A). The frequency of spontaneous neurotransmitter release was also found to be increased in DAPT-treated PS1mut neurons (1.72 ± 0.49 s^-1 for control PS1mut and 4.28 ± 0.98 s^-1 for DAPT-treated PS1mut neurons; p < 0.05; Fig. 6B). Both results indicate that the PS1mutant phenotype can be rescued by inhibiting -secretase activity.

The addition of A₄₂ Decreases Evoked and Spontaneous Neurotransmitter Release in PS1mut Neurons—If the DAPT effect of rescuing the PS1mut phenotype is mediated by a decreased production and extracellular release of A peptides, then monomeric A₄₂ should negatively affect synaptic function in non-transgenic cells. Indeed, non-transgenic neurons treated with A₄₂ (100 nM) showed a significant reduction in AMPA-EPSC responses compared with untreated non-transgenic neurons (2.69 ± 0.34 nA for control Non-tg and 1.61 ± 0.15 nA for A-treated Non-tg neurons; p < 0.05; Fig. 6C). The frequency of spontaneous neurotransmitter release was also significantly lower in nontransgenic neurons treated with A₄₂ (4.98 ± 1.4 s^-1 for control Non-tg and 1.60 ± 0.41 s^-1 for A-treated Non-tg cells; p < 0.05; Fig. 6D). In summary, A₄₂ applied extracellularly reduces both the amplitude and the frequency of excitatory synaptic currents in autaptic hippocampal cultures of non-transgenic mice, thus mimicking the effect of mutant PS1.

View larger version (41K): [in this window] [in a new window]   FIGURE 5. Prolonged repetitive and high frequency stimulation of Non-tg neurons and neurons expressing PS1wt or PS1mut. A, sample recordings of the first 10 responses in a 10 Hz train. B, average EPSC amplitudes of Non-tg (n = 16), PS1wt (n = 25), and PS1mut (n = 32) during 10-Hz train normalized to the size of first response. C, sample recordings of 5 responses in a 50-Hz train. D, average EPSC amplitudes of Non-tg (n = 21), PS1wt (n = 28), and PS1mut (n = 35) during 50-Hz train normalized to the size of first response.

View larger version (26K): [in this window] [in a new window]   FIGURE 6. Evoked and spontaneous neurotransmitter release in neurons treated either with DAPT or A42. A, AMPA-EPSC amplitude in neurons from control PS1mut (n = 27) and PS1mut neurons treated with DAPT (n = 25). B, mEPSC frequency in control PS1mut neurons (n = 22) and PS1mut neurons treated with DAPT (n = 22). C, AMPA-EPSC amplitude in control Non-tg neurons (n = 9) and Non-tg neurons treated with A42 (n = 11). D, mEPSC frequency in control Non-tg neurons (n = 9) and Non-tg neurons treated with A42 (n = 10) (*, p < 0.05).

View larger version (49K): [in this window] [in a new window]   FIGURE 7. Structural analysis of synapses in microisland cultures from PS1wt and PS1mut neurons. A, immunostains of autaptic neurons. Aa and Ab, staining for the dendritic marker microtubule-associated protein 2 to assess the size of PS1wt and PS1mut neurons. Ac and Ad, staining of the same culture for synaptophysin to assess the number of synapses. Ae and Af, merged picture of microtubule-associated protein 2 and synaptophysin staining. Scale bar,25 µm. B, density of synapses on dendrites measured as number of synaptophysin-positive spots per 10 µm dendrite from PS1wt (n = 56) and PS1mut (n = 63) neurons from three independent preparations (*, p < 0.05).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The ensuing question is: how can mutant PS1 affect the formation of excitatory synapses in cultured neurons? A recent electrophysiological study on cultured neurons of PS1 knockout mice favors a gain-of-function hypothesis for explaining our results (37). The enhanced mEPSC frequency as well as enhanced synapse formation observed (37) corroborate our observation that the mutant PS1 defect in synaptic strength can be rescued by inhibition of -secretase activity. Therefore, a pathological gain-of-function has to be considered, pertaining to four possible mechanisms. First, mutant PS1 affects synaptic formation due to enhanced generation of A₄₂ at the expense of A₄₀ (38). Second, altered generation of other APP cleavage products, for example reduced generation of the C-terminal cleavage product AICD (APP intracellular domain), may be critical (4). Third, mechanisms secondary to alterations in the intracellular calcium storage, as described in cells expressing mutant PS1, may be involved (21, 39-42). Fourth, cleavage of other substrates of the -secretase complex could be involved, e.g. Notch1 and homologs, Notch ligands Jagged2 and Delta, ErbB4, CD44, low density lipoprotein receptor-related protein, N- and E-cadherins, nectin-1, p75 neurotrophin receptor, syndecan3, telencephalin, and deleted in colorectal cancer (DCC) (43). A loss-of-function mechanism in PS1-mediated cleavage of any of these proteins cannot account for the reduced synaptic responses in mutant PS1 neurons, since we show that the reduced synaptic response in mutant PS1 can be rescued by the application of -secretase blocker. The involvement of alterations in calcium storage by mutant PS1 (21, 41, 42), however, cannot be excluded.

View larger version (116K): [in this window] [in a new window]   FIGURE 8. Synaptophysin staining of sections derived from PS1wt and PS1mut mice. A, synaptophysin-positive presynaptic terminals in mouse hippocampus. Aa and Ab, low magnification view of PS1wt and PS1mut hippocampus from 1-month-old mice: 1, stratum granulosum; 2, stratum moleculare; 3, stratum lacunosum; 4, stratum radiatum; 5, stratum pyramidale; 6, stratum oriens. Scale bar, 100 µm. Ac and Ad, synaptophysin staining of the stratum radiatum from 1-month-old PS1wt and PS1mut mice. Ac and Ad, synaptophysin-positive presynaptic terminals in the stratum radiatum and stratum moleculare (Ae and Af) of 1-month-old PS1wt and PS1mut mice. Scale bar,25 µm. B, average staining intensity in the stratum radiatum of PS1wt (n = 18 areas, 9 sections) and PS1mut (n = 18 areas; 9 sections) mice at the age of 1 and 3 months, respectively. C, average staining intensity in the stratum moleculare of PS1wt (n = 18 areas, 9 sections) and PS1mut (n = 18 areas, 9 sections) mice at the age of 1 and 3 months, respectively. D, Western blot analysis of synaptophysin expression relative to -actin in hippocampal brain sections of three PS1wt and PS1mut mice of 1 and 3 months of age. Relative band intensity of synaptophysin expression normalized to-actin levels revealed a lower expression of synaptophysin in PS1mut mice compared with PS1wt both at 1 and in 3 months of age mice (*, p < 0.05).

This study analyzing synaptic transmission in mutant PS1 transgenic mice further supports the idea that soluble A₄₀ may play a normal physiological role in synapse formation and maintenance. This role can become compromised by decreased production of A₄₀ and/or by increased generation of A₄₂, particularly oligomer formation, thus altering A function from a beneficial to a pathological one (49). This toxic gain-of-function of A in synapse formation and remodeling underscores the need for therapeutic attempts aimed at reducing the level of A₄₂ in this cellular compartment, thereby preventing synaptic degeneration and ameliorating cognitive deficits in Alzheimer disease. In light of the accumulating evidence for a physiological role of A₄₀ in modulating synaptic strength (44-48), methods for preferentially decreasing A₄₂ without affecting A₄₀ production are needed. This may include the generation of specific antibodies targeted against oligomeric A₄₂ or the use of selective -secretase inhibitors.

	FOOTNOTES

 
^* This work was supported by Deutsche Forschungsgemeinschaft Grant SFB 596 and the Bayerische Forschungsverbunt ForPrion. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ To whom correspondence should be addressed. Tel.: 89-2180-78010; Fax: 89-2180-78037; E-mail: jochen.herms{at}med.uni-muenchen.de.

² The abbreviations used are: PS, presenilin; APP, amyloid precursor protein; DIV, days in vitro; RRP, releasable vesicle pool; DAPT, [N-(3,5-difluorophenacetyl-L-alanyl)]-S-phenylglycine t-butyl ester; NMDA, N-methyl-D-aspar-tate; AMPA, -amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; EPSC, excitatory post-synaptic current; mEPSC, miniature EPSC; non-tg, non-transgenic; PS1wt, wild-type human PS1; PS1mut, mutant human PS1.

	ACKNOWLEDGMENTS

 
We thank Dr. Michael Mansour for helpful suggestions throughout the project, Dr. Gerda Mitteregger for excellent support in animal breeding, and the late Doris Schechinger for excellent technical assistance.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

1. Hardy, J., and Gwinn-Hardy, K. (1998) Science 282, 1075-1079[Abstract/Free Full Text]
2. De Strooper, B., Saftig, P., Craessaerts, K., Vanderstichele, H., Guhde, G., Annaert, W., von Figura, K., and Van Leuven, F. (1998) Nature 391, 387-390[CrossRef][Medline] [Order article via Infotrieve]
3. De Strooper, B., Annaert, W., Cupers, P., Saftig, P., Craessaerts, K., Mumm, J. S., Schroeter, E. H., Schrijvers, V., Wolfe, M. S., Ray, W. J., Goate, A., and Kopan, R. (1999) Nature 398, 518-522[CrossRef][Medline] [Order article via Infotrieve]
4. Moehlmann, T., Winkler, E., Xia, X., Edbauer, D., Murrell, J., Capell, A., Kaether, C., Zheng, H., Ghetti, B., Haass, C., and Steiner, H. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 8025-8030[Abstract/Free Full Text]
5. Schroeter, E. H., Ilagan, M. X., Brunkan, A. L., Hecimovic, S., Li, Y. M., Xu, M., Lewis, H. D., Saxena, M. T., De Strooper, B., Coonrod, A., Tomita, T., Iwatsubo, T., Moore, C. L., Goate, A., Wolfe, M. S., Shearman, M., and Kopan, R. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 13075-13080[Abstract/Free Full Text]
6. Cao, X., and Sudhof, T. C. (2001) Science 293, 115-120[Abstract/Free Full Text]
7. Terry, R. D., Masliah, E., Salmon, D. P., Butters, N., DeTeresa, R., Hill, R., Hansen, L. A., and Katzman, R. (1991) Ann. Neurol. 30, 572-580[CrossRef][Medline] [Order article via Infotrieve]
8. Braak, E., and Braak, H. (1997) Acta Neuropathol. (Berl.) 93, 323-325[CrossRef][Medline] [Order article via Infotrieve]
9. Chapman, P. F., White, G. L., Jones, M. W., Cooper-Blacketer, D., Marshall, V. J., Irizarry, M., Younkin, L., Good, M. A., Bliss, T. V., Hyman, B. T., Younkin, S. G., and Hsiao, K. K. (1999) Nat. Neurosci. 2, 271-276[CrossRef][Medline] [Order article via Infotrieve]
10. Moechars, D., Dewachter, I., Lorent, K., Reverse, D., Baekelandt, V., Naidu, A., Tesseur, I., Spittaels, K., Haute, C. V., Checler, F., Godaux, E., Cordell, B., and Van Leuven, F. (1999) J. Biol. Chem. 274, 6483-6492[Abstract/Free Full Text]
11. Nalbantoglu, J., Tirado-Santiago, G., Lahsaini, A., Poirier, J., Goncalves, O., Verge, G., Momoli, F., Welner, S. A., Massicotte, G., Julien, J. P., and Shapiro, M. L. (1997) Nature 387, 500-505[CrossRef][Medline] [Order article via Infotrieve]
12. Fitzjohn, S. M., Morton, R. A., Kuenzi, F., Rosahl, T. W., Shearman, M., Lewis, H., Smith, D., Reynolds, D. S., Davies, C. H., Collingridge, G. L., and Seabrook, G. R. (2001) J. Neurosci. 21, 4691-4698[Abstract/Free Full Text]
13. Hsia, A. Y., Masliah, E., McConlogue, L., Yu, G. Q., Tatsuno, G., Hu, K., Kholodenko, D., Malenka, R. C., Nicoll, R. A., and Mucke, L. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 3228-3233[Abstract/Free Full Text]
14. Larson, J., Lynch, G., Games, D., and Seubert, P. (1999) Brain Res. 840, 23-35[CrossRef][Medline] [Order article via Infotrieve]
15. Roder, S., Danober, L., Pozza, M. F., Lingenhoehl, K., Wiederhold, K. H., and Olpe, H. R. (2003) Neuroscience 120, 705-720[CrossRef][Medline] [Order article via Infotrieve]
16. Gong, B., Vitolo, O. V., Trinchese, F., Liu, S., Shelanski, M., and Arancio, O. (2004) J. Clin. Investig. 114, 1624-1634[CrossRef][Medline] [Order article via Infotrieve]
17. Trinchese, F., Liu, S., Battaglia, F., Walter, S., Mathews, P. M., and Arancio, O. (2004) Ann. Neurol. 55, 801-814[CrossRef][Medline] [Order article via Infotrieve]
18. Gureviciene, I., Ikonen, S., Gurevicius, K., Sarkaki, A., van Groen, T., Pussinen, R., Ylinen, A., and Tanila, H. (2004) Neurobiol. Dis. 15, 188-195[CrossRef][Medline] [Order article via Infotrieve]
19. Parent, A., Linden, D. J., Sisodia, S. S., and Borchelt, D. R. (1999) Neurobiol. Dis. 6, 56-62[CrossRef][Medline] [Order article via Infotrieve]
20. Barrow, P. A., Empson, R. M., Gladwell, S. J., Anderson, C. M., Killick, R., Yu, X., Jefferys, J. G., and Duff, K. (2000) Neurobiol. Dis. 7, 119-126[CrossRef][Medline] [Order article via Infotrieve]
21. Schneider, I., Reverse, D., Dewachter, I., Ris, L., Caluwaerts, N., Kuiperi, C., Gilis, M., Geerts, H., Kretzschmar, H., Godaux, E., Moechars, D., Van Leuven, F., and Herms, J. (2001) J. Biol. Chem. 276, 11539-11544[Abstract/Free Full Text]
22. Saura, C. A., Choi, S. Y., Beglopoulos, V., Malkani, S., Zhang, D., Shankaranarayana Rao, B. S., Chattarji, S., Kelleher, R. J., III, Kandel, E. R., Duff, K., Kirkwood, A., and Shen, J. (2004) Neuron 42, 23-36[CrossRef][Medline] [Order article via Infotrieve]
23. Marjaux, E., Hartmann, D., and De Strooper, B. (2004) Neuron 42, 189-192[CrossRef][Medline] [Order article via Infotrieve]
24. Dewachter, I., Van Dorpe, J., Smeijers, L., Gilis, M., Kuiperi, C., Laenen, I., Caluwaerts, N., Moechars, D., Checler, F., Vanderstichele, H., and Van Leuven, F. (2000) J. Neurosci. 20, 6452-6458[Abstract/Free Full Text]
25. Rosenmund, C., Clements, J. D., and Westbrook, G. L. (1993) Science 262, 754-757[Abstract/Free Full Text]
26. Rosenmund, C., Feltz, A., and Westbrook, G. L. (1995) J. Neurophysiol. 73, 427-430[Abstract/Free Full Text]
27. Priller, C., Bauer, T., Mitteregger, G., Krebs, B., Kretzschmar, H. A., and Herms, J. (2006) J. Neurosci. 26, 7212-7221[Abstract/Free Full Text]
28. Masliah, E., Terry, R. D., Mallory, M., Alford, M., and Hansen, L. A. (1990) Am. J. Pathol. 137, 1293-1297[Abstract]
29. Masliah, E., Terry, R. D., Alford, M., and DeTeresa, R. (1990) J. Histochem. Cytochem. 38, 837-844[Abstract]
30. Krebs, B., Kohlmannsperger, V., Nolting, S., Schmalzbauer, R., and Kretzschmar, H. A. (2006) J. Histochem. Cytochem. 54, 559-565[Abstract/Free Full Text]
31. Bekkers, J. M., and Stevens, C. F. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 7834-7838[Abstract/Free Full Text]
32. Rosenmund, C., and Stevens, C. F. (1996) Neuron 16, 1197-1207[CrossRef][Medline] [Order article via Infotrieve]
33. Reim, K., Mansour, M., Varoqueaux, F., McMahon, H. T., Sudhof, T. C., Brose, N., and Rosenmund, C. (2001) Cell 104, 71-81[CrossRef][Medline] [Order article via Infotrieve]
34. Schneggenburger, R., Meyer, A. C., and Neher, E. (1999) Neuron 23, 399-409[CrossRef][Medline] [Order article via Infotrieve]
35. Cohen-Cory, S. (2002) Science 298, 770-776[Abstract/Free Full Text]
36. Zaman, S. H., Parent, A., Laskey, A., Lee, M. K., Borchelt, D. R., Sisodia, S. S., and Malinow, R. (2000) Neurobiol. Dis. 7, 54-63[CrossRef][Medline] [Order article via Infotrieve]
37. Parent, A. T., Barnes, N. Y., Taniguchi, Y., Thinakaran, G., and Sisodia, S. S. (2005) J. Neurosci. 25, 1540-1549[Abstract/Free Full Text]
38. Murayama, O., Tomita, T., Nihonmatsu, N., Murayama, M., Sun, X., Honda, T., Iwatsubo, T., and Takashima, A. (1999) Neurosci. Lett. 265, 61-63[CrossRef][Medline] [Order article via Infotrieve]
39. Herms, J., Schneider, I., Dewachter, I., Caluwaerts, N., Kretzschmar, H., and Van Leuven, F. (2003) J. Biol. Chem. 278, 2484-2489[Abstract/Free Full Text]
40. Leissring, M. A., Murphy, M. P., Mead, T. R., Akbari, Y., Sugarman, M. C., Jannatipour, M., Anliker, B., Muller, U., Saftig, P., De Strooper, B., Wolfe, M. S., Golde, T. E., and LaFerla, F. M. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 4697-4702[Abstract/Free Full Text]
41. Leissring, M. A., Paul, B. A., Parker, I., Cotman, C. W., and LaFerla, F. M. (1999) J. Neurochem. 72, 1061-1068[CrossRef][Medline] [Order article via Infotrieve]
42. LaFerla, F. M. (2002) Nat. Rev. Neurosci. 3, 862-872[Medline] [Order article via Infotrieve]
43. Vetrivel, K. S., Cheng, H., Kim, S. H., Chen, Y., Barnes, N. Y., Parent, A. T., Sisodia, S. S., and Thinakaran, G. (2005) J. Biol. Chem. 280, 25892-25900[Abstract/Free Full Text]
44. Snyder, E. M., Nong, Y., Almeida, C. G., Paul, S., Moran, T., Choi, E. Y., Nairn, A. C., Salter, M. W., Lombroso, P. J., Gouras, G. K., and Greengard, P. (2005) Nat. Neurosci. 8, 1051-1058[CrossRef][Medline] [Order article via Infotrieve]
45. Chang, E. H., Savage, M. J., Flood, D. G., Thomas, J. M., Levy, R. B., Mahadomrongkul, V., Shirao, T., Aoki, C., and Huerta, P. T. (2006) Proc. Natl. Acad. Sci. U. S. A. 103, 3410-3415[Abstract/Free Full Text]
46. Kamenetz, F., Tomita, T., Hsieh, H., Seabrook, G., Borchelt, D., Iwatsubo, T., Sisodia, S., and Malinow, R. (2003) Neuron 37, 925-937[CrossRef][Medline] [Order article via Infotrieve]
47. Almeida, C. G., Tampellini, D., Takahashi, R. H., Greengard, P., Lin, M. T., Snyder, E. M., and Gouras, G. K. (2005) Neurobiol. Dis. 20, 187-198[CrossRef][Medline] [Order article via Infotrieve]
48. Roselli, F., Tirard, M., Lu, J., Hutzler, P., Lamberti, P., Livrea, P., Morabito, M., and Almeida, O. F. (2005) J. Neurosci. 25, 11061-11070[Abstract/Free Full Text]
49. Huang, S. M., Mouri, A., Kokubo, H., Nakajima, R., Suemoto, T., Higuchi, M., Staufenbiel, M., Noda, Y., Yamaguchi, H., Nabeshima, T., Saido, T. C., and Iwata, N. (2006) J. Biol. Chem. 281, 17941-17951[Abstract/Free Full Text]

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				S. Knafo, L. Alonso-Nanclares, J. Gonzalez-Soriano, P. Merino-Serrais, I. Fernaud-Espinosa, I. Ferrer, and J. DeFelipe Widespread Changes in Dendritic Spines in a Model of Alzheimer's Disease Cereb Cortex, March 1, 2009; 19(3): 586 - 592. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 282/2/1119    most recent M605066200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Priller, C. Articles by Herms, J. Search for Related Content PubMed PubMed Citation Articles by Priller, C. Articles by Herms, J. Social Bookmarking                      What's this?