FMRP Involvement in Formation of Synapses among Cultured Hippocampal Neurons

Katharina Braun and Menahem Segal1

1 Leibniz Institute for Neurobiology, POB 1860, D-39008 Magdeburg, Germany and Department of Neurobiology, The Weizmann Institute, Rehovot 76100, Israel


    Abstract
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
Fragile-X, the main cause of inherited human mental retardation is associated with the absence of a recently identified fragile-X mental retardation protein (FMRP). Mice in which this protein is lacking due to a knockout (KO) mutation are reported to express altered dendritic spines on their cortical neurons compared with wild type (WT) controls. We have used tissue-cultured neurons to examine differences in morphology and synaptic connectivity between WT and FMRP-deficient mice. Hippocampal neurons taken from KO mice and grown in culture for 3 weeks have shorter dendrites and fewer dendritic spines than their WT counterparts. Also, KO cells tend to express fewer functional synaptic connections, which develop more slowly and produce smaller excitatory synaptic currents than WT controls. These observations may have important implications for the understanding of mental retardation associated with the absence of FMRP.


    Introduction
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
In mental retardation there has been an apparent lack of correlation between the structure of the brain and its cognitive functions. However, exciting new information focuses on the dendritic spine as the putative locus of synaptic interaction, neuronal plasticity and long-term memory (Bliss and Collingridge, 1993Go; Harris and Kater, 1994Go; Segal et al., 2000Go). The fragile-X mental retardation protein (FMRP) is absent in fragile-X mentally retarded children. The inability to synthesize FMRP, due to an abnormally long CGG repeat in the promoter region leading to hypermethylation and silencing of the Fmr-1 gene, is the leading cause of genetically inherited mental retardation (Hagerman, 1997Go). The pathological changes in the brain, observed in magnetic resonance imaging studies, include abnormalities in the cerebellum, most prominent in the posterior vermis (Reiss et al., 1991aGo,bGo), decreased volume of the superior temporal gyrus (Reiss et al., 1994Go) and enlarged volume of the caudate nucleus (Reiss et al., 1995Go). In young but not older patients increased hippocampal volume was also detected (Kooy et al., 1996Go).

Recent morphological studies have located FMRP to the dendrite near dendritic spines. These protrusions from the dendrite represent morphologically and functionally specialized post-synaptic structures, which undergo dramatic proliferative and regressive changes during brain development, learning and memory formation (Harris and Kater, 1994Go; Segal et al., 2000Go). Qualitative observations in several cortical regions, including the anterior cingulate area of human autopsy material from patients suffering from fragile-X mental retardation syndrome detected abnormal spine morphology and reduced synaptic size, indicative of immature spines (Rudelli et al., 1985Go; Hinton et al., 1991Go). These observations were confirmed in a quantitative analysis of spine morphology in fragile-X brains, where an abundance of long, thin immature spines was found compared with control brains (Irwin et al., 2000Go). Morphological analysis on cortical neurons of an FMRP-deficient mouse (Pieretti et al., 1991Go; Consortium, 1994Go; Oostra and Hoogeveen, 1997Go), however, revealed no differences in spine densities in the Fmr1-knock out mutant mice, but they display a similar abundance of immature thin and long spines (Comery et al., 1997Go; Irwin et al., 2000Go), as has been described in human tissue (Irwin et al., 2000Go).

These data promote the hypothesis that fragile-X mental retardation may be manifested by abnormal synaptic connections, associated with altered densities and morphology of dendritic spines and their functions. The abundance of spines and their immature morphology in fragile-X brains may be the result of impaired synaptic maturation and reduced pruning of super-numerary spines, a developmental process which normally occurs during postnatal brain maturation (Huttenlocher et al., 1982Go) and in response to environmental changes, experience and learning (Bock and Braun, 1998Go, 1999Go).

Recent studies suggest that FMRP may be a critical factor for both the maturation of dendritic spines and synaptic elimination (Weiler and Greenough, 1999Go). FMRP is synthesized at the synapse, in a synaptoneurosome preparation (Weiler et al., 1997Go), in response to glutamate, KCl and class I metabotropic glutamate receptor agonists. The protein kinase C inhibitor calphostin prevents the synthesis of FMRP (Weiler et al., 1997Go; Weiler and Greenough, 1999Go). FMRP appears to be associated with somatic and dendritic polyribosomes (Feng et al., 1997Go), and it may serve as a putative transporter (chaperone) of mRNA from the nucleus to the cytoplasm (Khandjian, 1999Go). However, a role for FMRP in targeting mRNAs to dendrites has yet to be established, as no differences in the patterns of mRNA localization for MAP2, CAM kinase II and dendrin in dendrites and in the dendritic transport of ARC mRNA were found between normal and Fmr1 knockout (KO) mice (Steward et al., 1998Go). An involvement of FMRP in the regulation of cAMP synthesis has been shown in platelets (Berry-Kravis and Huttenlocher, 1992Go), lymphoblastoid cells (Berry-Kravis et al., 1995Go) and neurotumor hybrid cell lines (Berry-Kravis and Ciurlionis, 1998Go).

To study spine morphology and function in controlled conditions, an in vitro system of cultured hippocampal neurons from the KO mouse mutant and its respective wild type (WT) controls was developed. In human fetal brain as well as in mice the hippocampus is one of the major sites of Fmr1 gene expression (Abitbol et al., 1993Go; Hinds et al., 1993Go). In this culture system of mouse hippocampal neurons the development of neuronal morphology, in particular of spine formation, and the electrical network properties were investigated in a combined physiological and morphological approach. These studies suggest a role for FMRP in the development of morphological and functional interactions among the cultured neurons.


    Materials and Methods
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
Tissue Culturing

FVB/NJ strain mice, RD/RD, KO and their WT counterparts were obtained from W.T. Greenough (Comery et al., 1997Go, Irwin et al., 2000Go). Mice were genotyped by polymerase chain reaction to confirm the lack of the Fmr1 gene, and housed in a local animal house. The results reported below were collected from five different litters, each experiment was made with WT and KO newborn mice of both sexes, born and prepared for culture on the same day. KO and WT mice were decapitated on the day of birth, and their brains were removed and placed in a chilled (4°C) dissecting medium, consisting of oxygenated Leibovitz L15 medium (Biological Industries, Beit Haemek, IL) enriched with 0.6% glucose and Gentamicin (Sigma, Israel, 20 µg/ml). The complete hippocampi of 8–10 pups in each experiment were dissected out and collected in the same medium. Tissue was mechanically dissociated with a fire polished Pasteur pipette and passed to the plating medium consisting of 5% heat inactivated horse serum (HS), 5% fetal calf serum (FCS), prepared in minimal essential medium (MEM)-Earl salts (Biological Industries, Illinois), enriched with 0.6% glucose, Gentamicin and 2 mM glutamax. About 0.3–0.5 x 106 cells in 1 ml of medium were plated on 13 mm polylysine-coated glass coverslips placed in each well of a 24-well plate (Papa et al., 1995Go). A monolayer of glia was grown on the glass for 1–2 weeks prior to the plating of the neurons (Papa et al., 1995Go; Murphy and Segal, 1996Go). Cells were left to grow in the incubator at 37°C, 5% CO2, for 4 days, then the medium was changed to 10% HS in enriched MEM plus a mixture of 5'-fluoro-2-deoxyuridine (FUDR)/uridine (Sigma, 20 and 50 mg/ml, respectively) to block glial proliferation. Four days later the medium was changed once again to 10% HS in enriched MEM, and no further changes were made. Cultures were used for physiological and morphological analysis at 1, 2 and 3 weeks after plating.

Electrophysiology

Cultures were removed from the incubator, placed in an inverted phase contrast microscope and superfused with the recording medium containing (in mM) NaCl 130, KCl 5, glucose 30, HEPES 25, CaCl2 2 and MgCl2 1. Osmolarity was adjusted to 320 mOsm with sucrose, pH 7.4. Individual cells were patch-clamped with 1.5 mm external diameter pipettes containing (in mM) K-acetate 120, Mg ATP 2, Na2 phosphocreatine 10, HEPES 10, sucrose 25, EGTA 200 µM, GTP-Tris 300 µM, and QX-314 5. When miniature inhibitory synaptic currents were recorded, the extracellular medium also contained 1 µM tetrodotoxin (TTX), 20 µM DNQX and 20–50 µM 2-aminophosphonovalerate (2-APV). When miniature excitatory synaptic currents were recorded, APV/DNQX were replaced by 50 µM bicuculline. Hyperosmotic medium or glutamate was injected through a 2 µm pressure pipette with brief (10–20 ms) pulses. A hyperosmotic medium contained normal recording medium supplemented with 300 mM sucrose and was used for evoking miniature postsynaptic currents (mPSCs) in the presence of TTX. Glutamate was injected onto cells adjacent to the one being recorded in order to evoke synaptic currents. Experiments were conducted at room temperature. Signals were amplified with Axopatch 200 and stored on an IBM computer for offline analysis, which included analysis of the synaptic currents using PClamp and Minianalysis software.

Double Labeling of Neuronal Morphology and Presynaptic Terminals

After fixation in 4% paraformaldehyde for 20 min the cultures were incubated with an antibody to the synaptic vesicle protein synaptophysin (Sigma), a marker for presynaptic terminals, diluted 1:100 in phosphate-buffered saline (PBS), pH 7.3, for 3 h and then incubated with a secondary anti-mouse antibody conjugated to the fluorescent molecule ALEXA 488 (Molecular Probes), diluted 1:100 in PBS for 45 min at room temperature. Single neurons were labeled with the fluorescent marker DiI prepared in oil and applied in a microdrop through a pressure pipette as detailed elsewhere (Papa et al., 1995Go). After 5–8 h of incubation with the dye the neurons were visualized in a Zeiss 510 confocal laser scanning microscope and 3-D images of neuronal somata, dendrites and spines were taken. The coverslips containing the cultures were coded, so that all steps, i.e. image acquisition and the quantification of neuronal parameters, were performed in a double blind routine. The neurons that were randomly selected for the analysis typically displayed a large and ramified dendritic tree and high densities of spines.

Image Acquisition

All scans were taken at a resolution of 524 x 524 pixels and 0.5 µm steps in the z-direction. From each neuron a low power image of the soma and dendrites (using a x40 oil objective, NA 1.4) was taken, resulting in stacks of between 12 and 30 images. These low power image stacks were used for the measurement of soma area and dendritic length. From each neuron between five and seven primary dendrites and their ramifications, at the level where they emerge from the soma, were zoomed fourfold and scanned again. These image stacks were handled as described below and used for the quantification of spine lengths, spine densities and detection of presynaptic labeling. A scan was taken first to visualize the DiI label alone (excitation wavelength 543 nm), which was later used for the quantification of spine densities. A second scan was then taken from the identical dendritic part using double excitation for DiI and ALEXA 488 (excitation wavelength 488 nm), these images being used for the selective quantification of different spine types (unoccupied ‘empty’, and spines occupied by one, two or more synaptophysin labeled puncta). This sequential ‘double scan’ procedure was performed because we observed bleaching of both fluorescent dyes during the double excitation scan and thus were concerned that we may lose the very thin, weakly labeled spines on our images. However, during the course of these experiments the comparison of the counts in the single DiI images and in the second, double fluorescent images revealed almost identical spine frequencies. For 3-D reconstruction and visualization of neurons (Figure 1A,BGo) and dendrites the software package IMARIS (Bitplane, Zurich, Switzerland), run on a Silicon graphics workstation, and our own programs, which were developed for this purpose (Herzog et al., 1997Go), were used, respectively.



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Figure 1.  (A,B) Comparison between the morphology of hippocampal neurons from WT (A) and Fmr1-KO (B) mice. Compared with normal neurons the Fmr1-KO neurons display shrunken dendrites and reduced spine frequencies. The scale bar is for A and B. (C,D) Single images taken from confocal image stacks of a dendrite from a WT neuron to illustrate double labeling of dendrites and spines (red) and synaptophysin-immunoreactive presynaptic terminals (green). The majority of spines are occupied by one or more synaptophysin-immunoreactive boutons (arrows). The scale bar in D is for C, D and E. (E1,E2) Two consecutive single images taken from a confocal image stack from a dendrite of a WT neuron to demonstrate that long, ‘filopodia-like’ spines are occupied by one or more synaptophysin immunoreactive terminal boutons (arrows).

 
Quantitative Analysis of Neuronal Morphology

The original image stacks were stored on CD, from which they were loaded into the quantification program Neurolucida (MicroBrightfield, Baltimore, MD), with which soma area, spine densities, and the length of dendrites and spines were measured. From each culture 5–9 DiI labeled neurons were collected for quantitative analysis, which resulted for the 1-week-old cultures in a total of 15 neurons for WT and nine neurons for Fmr1-KO, and for the 3-week-old cultures in a total of 24 cells for WT and 24 cells for KO. For each neuron the total length of all dendrites was measured at low magnification (Figures 2 and 3GoGo). Spine densities in the 7-day-old cultures were analyzed on a total of 11 204 µm dendritic length for WT neurons and 4607 µm dendritic length for Fmr1-KO neurons. Spine densities in the 21-day-old cultures were analyzed on a total of 19 412 µm dendritic length for WT neurons and 16 416 µm dendritic length for Fmr1-KO neurons. Spine length was measured in the confocal image stacks for an average of 50 spines per neuron in both age groups.



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Figure 2.  Quantitative comparison of morphological parameters of cultured hippoampal neurons from WT and Fmr1-KO mice at 7 DIV. The values shown are means with SEM. (A) Spine densities are reduced in neurons from Fmr1-KO mice. *P < 0.05, t-test. (B) No difference in mean spine length was found between neurons from WT and Fmr1-KO mice. (C) No difference in the relative distribution of spine lengths was found between neurons from WT and Fmr1-KO mice. Spines were grouped according to their length: 0/1 = from 0 to 1 µm, 1/2 = from 1 to 2 µm etc., and the percent distribution of spines in each length group in relation to the total spine number is shown. (D) Mean dendritic length is significantly reduced in neurons from Fmr1-KO mice. *P < 0.05, t-test. (E) No difference in mean soma size was found between neurons from WT and Fmr1-KO mice.

 


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Figure 3.  Quantitative comparison of morphological parameters of cultured hippocampal neurons from WT and Fmr1-KO mice at 21 DIV. The values shown are means with SEM. (A) Spine densities are reduced in neurons from Fmr1-KO mice. *P < 0.05, t-test. (B) No difference in mean spine length was found between neurons from WT and Fmr1-KO mice. (C) No difference in the relative distribution of spine lengths was found between neurons from WT and Fmr1-KO mice. Spines were grouped according to their length: 0/1 = from 0 to 1 µm, 1/2 = from 1 to 2 µm etc., and the percent distribution of spines in each length group in relation to the total spine number is shown. (D) Spines were further subdivided according to their association with synaptophysin-immunoreactive presynaptic terminals into empty (unoccupied) spines and spines occupied by one, two or three synaptophysin-immunoreactive terminals. Neurons from Fmr1-KO mice show significantly reduced total numbers of synaptophysin-immunoreactive terminals, reduced densities of spines which are associated with one synaptophysin-immunoreactive terminal and reduced numbers of unoccupied spines. The density of shaft synapses, which were defined as synaptophysin-immunoreactive terminals contacting the dendritic shaft, was not different between the two groups. (E) Mean dendritic length is significantly reduced in neurons from Fmr1-KO mice. *P < 0.05, t-test. (F) No difference in mean soma size was found between neurons from WT and Fmr1-KO mice.

 
Statistical Analysis

After testing for normal distribution, Student's t-test was applied for the group comparison of spine densities, dendritic length and soma size. Spine lengths, which showed a skewed distribution, were analyzed by the Mann-Whitney U-test. Differences in the distribution of spine groups, sorted in groups of 1 µm length, between Fmr1-KO and WT neurons were analyzed with a {chi}2 test.


    Results
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
Morphology

Cell density and survival in cultures at the age of 1, 2 or 3 weeks were about the same for the WT and KO cells, following plating of the same number of cells in a culture dish. This lack of difference in cell survival between the two cell types indicate that the results, reported below, are not merely a function of selective survival of certain types of cells or their selective vulnerability to neurotoxic insults. Both culture types contained primarily medium size, pyramidal-like neurons, likely to be of CA1 origin (Papa et al., 1995Go), but also clumps of smaller cells with round somata, likely to be dentate granular neurons. There was no apparent difference in the proportion of GABAergic cells stained for the enzyme glutamate decarboxylase (GAD, data not shown). Figure 1A,BGo illustrates the characteristic features of 3-week-old cultured hippocampal neurons from WT (Figure 1AGo) and Fmr1-KO mice (Figure 1BGo). In general, the overall spine density on 7-and 21-day-old cultured hippocampal neurons from both WT and Fmr1-KO mice was ~2–3 times higher than those observed in rat hippocampal neurons in culture (Papa et al., 1995Go). Spines in both KO and WT cells exhibited a variety of lengths and shapes as reported in vivo. Furthermore, synaptophysin-immunoreactive boutons were found to occupy all morphologically distinguishable spine types (Figure 1C,DGo), including the long, thin ‘filopodia-like’ spines (Figure 1GoE1,E2), indicating that they may possess functional synaptic connections.

At the age of 7 and 21 days in vitro (DIV) significant differences of neuronal morphology were observed. At both ages the neurons from the Fmr1-KO mice displayed significantly shorter dendrites (Figures 2DGo and 3E) and lower spine densities (Figures 2AGo and 3A) compared with the WT neurons, whereas the soma size remained unchanged (Figures 2EGo and 3F). Spine densities and dendritic length/cell and soma size did not show significant differences between 7 and 21 DIV in WT and KO neurons. The length of dendritic spines showed a significant decrease between 7 and 21 DIV (Kruskal-Wallis analysis of variance, P <= 0.001; Dunn's test, P < 0.05) in WT and KO neurons (compare Figures 2BGo and 3B). The {chi}2 test revealed no difference in the relative distribution of spine sizes between KO and WT neurons at 7 and 21 DIV (compare Figures 2CGo and 3C).

In order to further distinguish between unoccupied (‘empty’) spines and spines which are contacted by one or several presynaptic boutons, thus representing morphologically complete and most likely physiologically functional synapses, we used a double labeling technique which allows the simultaneous visualization of postsynaptic and presynaptic elements (Figure 1GoC-E). This analysis, which was done only in the 21-day-old cultures, showed that the neurons from the Fmr1-KO mice display significantly lower densities of synaptophysin-immunoreactive puncta, a lower density of unoccupied ‘empty’ spines and also lower densities of the predominant spine type, contacted by one terminal bouton (Figure 3DGo). No quantitative differences in the density of shaft synapses, counted as synaptophysin-immuno-reactive boutons located on the shaft of the DiI-labeled dendrite, was observed (Figure 3DGo).

Electrophysiology

A total of 76 WT cells and 90 KO cells, studied in five different experiments, were selected for analysis. In each experiment, cells were recorded at 1, 2 and 3 weeks in culture, and the results of the five experiments were pooled. For each 13 mm cover-glass, 2–5 cells were recorded over 1–2 h after removal from the incubator. Only ‘healthy’ cells with a low resting membrane potential (~–60 mV) and good seal resistance were included in the data analysis. There were no apparent differences in passive properties, voltage gated conductances or action potentials between the groups of cells (data not shown).

The formation of synaptic connections among cells was studied in 1-to 3-week-old WT and KO cultures. Spontaneous synaptic activity was recorded in patch-clamped neurons at about resting potential (-60mV) and at a potential where there is a clear distinction between excitatory, inward synaptic currents and inhibitory, outward synaptic currents (-30 mV; Figure 4AGo). Most of control cells express spontaneous synaptic activity already at 1 week in culture (12/16 cells, recorded in five different experiments), whereas most of the KO cells were silent within the first week in culture (6/21 cells active, five experiments, {chi}2 test, P < 0.005; Figure 5Go). Cells of the KO group expressed spontaneous activity at 2 weeks in culture in a manner that was not significantly different from controls (9/15 cells in the KO group, three experiments, and 10/13 cells in the WT, three experiments, P > 0.4; Figure 4AGo). Thus, it seems that the KO cells are slower in establishing synaptic connections than are controls. Once the connections were established, there were no apparent differences between the two cell groups in the frequency of occurrence of spontaneous activity.



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Figure 4.  Electrical activity recorded from 3-week-old, voltage-clamped hippocampal neurons in culture. (A,B) Spontaneous activity recorded from a cell held at -30 mV. The burst consists of inward deflections representing spontaneous EPSCs and outward deflections representing IPSCs. A is shown at a slow time scale, B at a fast time scale, same cell. Note that the outward currents have a much slower decay time constant than the inward currents, which can in fact show up during the decay of the slow events. (C,D) Recording of evoked responses to stimulation of a neuron that is afferent to the one recorded from. (C) Recording of a cell from a WT mouse. (D) A cell from an FMRP-deficient mouse. The top record is a response taken when the cell is held at -30 mV, the bottom one is from a cell held at -60 mV. Note that in the bottom record all deflections are downward, whereas in the top record the fast events are downward (= EPSCs) and the slow events are upward going (= IPSCs). In both C and D the left records are at a slow time scale and the right record is the first event seen in the left record, but at a fast time scale.

 


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Figure 5.  Age-dependent increase in synaptic connectivity in KO compared with WT cells. (A) Presence of spontaneous activity in the two cell groups at two ages. Ordinate, proportion of cells demonstrating spontaneous synaptic currents during a standard 1 min recording time. Asterisks, significant difference at the P < 0.01 level. See text for further details. (B) Proportion of effective synaptic connections evoked by stimulation of afferent neurons. Same groups as in A. (C) Postsynaptic currents evoked in WT and KO cells in response to topical application of glutamate. Cells were held at the designated membrane potentials (abscissa, mV). There was no difference between the two groups of cells in response to glutamate, and both showed the characteristic N-methyl-D-aspartate-mediated rectification at low membrane potentials.

 
A method for assessing more precisely the presence of synaptic connections among relatively quiet cells involves stimulation of putative afferents, within a perimeter of ~250 µm from the recorded cell (Vicario-Abejon et al., 1998Go). Short pulses of glutamate can evoke a barrage of excitatory/inhibitory post-synaptic currents (EPSCs/IPSCs) that represent mono-as well as polysynaptic connections between the stimulated and the recorded cell. Routinely, we tested for the presence of connections from 3–7 putative afferents in different areas of the field of view. This method of comparison (e.g. Figure 4C,DGo) has also yielded a significant difference between the KO and the WT cells at 1 week in culture. In the former group, only ~40% of tested connections yielded significant responses (n = 11 postsynaptic cells) compared with 79% of the WT cells (t = 3.22, P < 0.004; Figure 5BGo). The magnitude of individual postsynaptic currents was not significantly different between the two groups at this age (29 ± 3.9 and 21.5 ± 4.8pA in the WT and KO, respectively). The difference in abundance of synaptic currents was annealed at 2 weeks in culture (eight KO cells, 32 connections tested and seven WT cells, 26 connections tested, five experiments; Figure 5BGo).

The difference in effective connectivity between the WT and KO cells at 1 week in culture can derive from lack of afferent innervation or from a delayed maturation of postsynaptic receptors. This was tested by measuring the responses of 1-week-old cells to pulse application of glutamate. There were no apparent differences between the two groups in response to glutamate, measured at different membrane potentials (four cells each group; Figure 5CGo).

Both spontaneous and evoked synaptic events consist of EPSCs and IPSCs. We could easily distinguish between them by holding the cell at -30 mV. At this potential, the former is inward going and the latter outward (Figure 4AGo). While there were no significant differences in connectivity between the two groups at 2–3 weeks in culture, the duration of the response to afferent stimulation was longer in the WT than in the KO cells (1.4 ± 0.16 versus 0.91 ± 0.14 s, respectively, n = 8 cells in each group, P < 0.02). The total number of IPSCs was not different between the WT and KO cells (n = 7.9 ± 1.4 and 8.0 ± 1.52 events in the two groups, respectively), but the total number of excitatory events per response was significantly smaller in KO cells (9.9 ± 2.07 versus 3.7 ± 1.05 EPSCs per response, respectively, n = 8 in each, P < 0.004). Likewise, the size of the excitatory events was larger for the WT than the KO cells (113 ± 22 versus 25 ± 8 pA, respectively, P < 0.01). Thus, KO cultures express less active mono-and polysynaptic connections among cells than WT cultures even at 2–3 weeks in vitro.

Finally, the difference in size and frequency of excitatory synaptic currents can reflect a difference in individual postsynaptic responses, being larger in the WT than the KO cells, or in release probability from presynaptic terminals. In order to distinguish between the two possibilities, we measured properties of miniature synaptic currents (mEPSCs) in 10 WT and nine KO cells recorded in presence of TTX. These experiments were conducted with 2-to 3-week-old cultures. Synaptic currents were measured spontaneously over standard periods of 1 min. In an additional set of cells, not analyzed herein, mEPSCs evoked in response to a hyperosmotic stimulus were measured. In addition to the rate of mEPSCs and peak amplitudes, we also measured rise time and half-decay time. There was no difference between the two groups of cells in any of the parameters analyzed (Figure 6Go). This indicates that the difference in evoked synaptic currents is not due to a change in the morphology of a single synaptic current but in their coordinated generation by afferent activity.



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Figure 6.  There is no significant difference between miniature spontaneous synaptic currents of WT and KO cells. (A) Sample illustration of mEPSCs recorded over a period of 15 s. Inward currents above noise level are detected and measured automatically using the Minianalysis software. (B-E) Averages of 10 WT and nine KO cells are presented for each of the following parameters: (B) frequency of mEPSCs recorded in a 1 min observation time; (C) rise time (in ms); (D) mEPSC amplitude (in pA); and (E) decay time (ms). Interestingly, both rise time and decay time tend to be shorter in the KO cells, but this is not significant statistically.

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
The present study demonstrates that FMRP-deficient neurons taken from the newborn hippocampus and grown in culture for up to 3 weeks are slower to develop mature neuronal and spine morphology than normal mouse neurons, and that this lag is also expressed in their synaptic physiology. The neurons from the KO mice, which display reduced dendritic length and spine density, express age dependent lower abundance and smaller excitatory synaptic currents than those expressed by the WT cells.

The presence of significantly lower spine densities in the FMRP-deficient neurons in hippocampal cultures is somewhat different from the in vivo observations, which report a non-significant tendency towards higher spine densities in the visual cortex of adult FMRP-deficient mice (Comery et al., 1997Go; Irwin et al., 2000Go). Furthermore, in contrast with the in vivo studies, where dendritic length was not different between neurons in the visual cortex or Fmr1-KO mice and WT mice (Irwin et al., 2000Go), we found significantly reduced dendritic length in the 7-and 21-day-old hippocampal neurons from the Fmr1-KO mice compared with those from the WT controls. Finally, we did not find significant differences of spine length and relative spine size distribution between KO and WT neurons at both ages analyzed. These discrepancies between our results and those obtained in in vivo studies may be due to their different origin: cortical neurons were analyzed in the in vivo study, whereas in our culture system we analyzed hippocampal neurons. The discrepancies may also be explained by a different maturation state, with the neurons in culture being examined at 1–3 weeks of age, whereas those studied in vivo were adults. Our observation of reduced dendritic length and spine frequency in KO hippocampal neurons at the ages of 7 and 21 days in culture indicates that the rate of dendritic and synaptic development may differ between the WT and KO neurons, being slower in the Fmr1-KO neurons. Indeed, the physiological experiments revealed a clear developmental lag in the formation of functional synapses between the two neuron groups. Assuming that the difference in spine density observed in vitro reflects the situation in vivo at an early developmental stage, it can be assumed that the normal process of maturation, which involves spine pruning, will not be seen in the KO mice and thereby may cause a reversal of the spine density difference at older ages, as is seen in vivo. This hypothesis needs to be examined both in vivo and in vitro.

The difference in synaptic activity between cells in the KO and WT neurons indicates that functional connections among the cells may be different; while the inhibitory synaptic currents appear to have the same abundance, the excitatory ones are less numerous in the 2-to 3-week-old KO cells. This reduced excitatory drive may result from a less efficient postsynaptic effector mechanism or a reduced release of neurotransmitter substance from presynaptic terminals. Apparently, in 2-week-old cells, there are no differences in the frequency or magnitude of mEPSCs between the two cell groups. This suggests that the individual synapse may not be different between the WT and the KO mice, but that there are more of them in the WT cells, as suggested by the morphological data. It is not yet known if there is a correlation between subtypes of synaptic currents and shapes of spines, and this will require a more detailed analysis. The large heterogeneity among spines in a given neuron makes such an analysis extremely tedious.

How are these morphological changes in spines and their functional correlates translated into mental retardation? In the behavior and cognitive functions of FMRP-deficient mouse strains no severe impairments were found in operant conditioning and discrimination tasks (Fisch et al., 1999Go). A mildly impaired performance in the Morris water maze was found (D'Hooge et al., 1997Go), with the Fmr1-KO mice showing reduced performance in the reversal test (Kooy et al., 1996Go; Bakker et al., 1999Go). The hallmark of mental capacity is the ability to express long term plasticity, and the expression of long-term potentiation (LTP) of reactivity to afferent stimulation has been correlated with performance in cognitive tasks in normal, drugged and mutated rodents. Data in this field are still scarce, and no evidence for a function of FMRP in LTP or short-term potentiation has been found (Godfraind et al., 1996Go). Finally, there is no direct link between the size of evoked synaptic currents and the ability to change them in a plasticity-producing protocol. Thus, this ability has to be tested more directly in future experiments.

Our results provide an encouraging basis for the further analysis of plasticity of neurons in culture and the role of FMRP in this process. The quantitative differences in neuronal morphology between the Fmr1-deficient and normal neurons, which has already been found in culture and which appears to be linked to differences in their physiological properties and network activity, makes the culture a promising in vitro system for the analysis of the mechanisms involved in activity-induced spine formation and/or elimination.


    Notes
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
The authors wish to express their thanks to Dr W.T. Greenough for the generous supply of the mice and for constructive consultations, to Ms V. Greenberger for preparing the cultures and to Ute Kreher for assistance in the quantification of neuronal morphology. This work was supported by a grant from the FRAXA Foundation.

Address correspondence to Menahem Segal, Department of Neurobiology, The Weizmann Institute, Rehovot 76100, Israel. Email: menahem.segal{at}weizmann.ac.il.


    References
 Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Notes
 References
 
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