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Mol. Cell. Neurosci. 34 (2007) 366 – 377
Impaired hippocampal synaptic transmission and plasticity in mice
lacking fibroblast growth factor 14
Maolei Xiao, a Lin Xu, b Fernanda Laezza, a Kelvin Yamada, b
Sheng Feng, c and David M. Ornitz a,⁎
a
Department of Molecular Biology and Pharmacology, Campus Box 8103, Washington University Medical School, 660 S. Euclid Avenue,
St. Louis, MO 63110, USA
b
Department of Neurology, Washington University School of Medicine, St. Louis, MO 63110, USA
c
Division of Biostatistics, Washington University School of Medicine, St. Louis, MO 63110, USA
Received 1 August 2006; revised 14 November 2006; accepted 14 November 2006
Available online 8 January 2007
Humans with an autosomal dominant missense mutation in fibroblast
growth factor 14 (FGF14) have impaired cognitive abilities and slowly
progressive spinocerebellar ataxia. To explore the mechanisms that
may account for this phenotype, we show that synaptic transmission at
hippocampal Schaffer collateral-CA1 synapses and short- and longterm potentiation are impaired in Fgf14−/− mice, indicating abnormalities in synaptic plasticity. Examination of CA1 synapses in Fgf14−/−
mice show a significant reduction in the number of synaptic vesicles
docked at presynaptic active zones and a significant synaptic fatigue/
depression during high/low-frequency stimulation. In addition,
mEPSC frequency, but not amplitude, is decreased in hippocampal
neurons derived from Fgf14−/− mice. Furthermore, expression of
selective synaptic proteins in Fgf14−/− mice was decreased. These
findings suggest a novel role for FGF14 in regulating synaptic plasticity
via presynaptic mechanisms by affecting the mobilization, trafficking,
or docking of synaptic vesicles to presynaptic active zones.
© 2006 Elsevier Inc. All rights reserved.
Keywords: Fibroblast growth factor 14 (FGF14); Synaptic transmission;
Synaptic plasticity; LTP; Synaptic vesicles
Introduction
The discovery of a mutation in FGF14 (F145S) in a Dutch
family with impaired cognitive function and progressive spinocerebellar ataxia suggests that FGF14 may play a role in mediating
human cognition and neuromuscular function (Brusse et al., 2006;
Van Swieten et al., 2003). An unrelated patient with mild mental
retardation (IQ 70) and ataxia was also found to carry a frameshift
mutation, FGF14 (D163fsX12) (Dalski et al., 2005), further
supporting a role for FGF14 in human cognition. The sensorimotor
deficits in humans are strikingly similar to the phenotype of Fgf14
⁎ Corresponding author. Fax: +1 314 362 7058.
E-mail address: dornitz@wustl.edu (D.M. Ornitz).
Available online on ScienceDirect (www.sciencedirect.com).
1044-7431/$ - see front matter © 2006 Elsevier Inc. All rights reserved.
doi:10.1016/j.mcn.2006.11.020
knockout (Fgf14−/−) mice (Wang et al., 2002), and the phenotype
of Fgf14−/− mice directly led to the identification of the human
mutation (Van Swieten et al., 2003). To further explore underlying
neurophysiological mechanisms, we examined hippocampal function in Fgf14−/− mice.
FGF14 (FHF4), a member of the FGF homologous factors
subfamily (FHF1–4), is widely expressed in the developing and
adult central nervous system (CNS), including the cerebellar granule
cell layer and the pyramidal cell layer of the Cornu AmmonTs (CA)
horn in the hippocampus (Smallwood et al., 1996; Wang et al., 2002;
Yamamoto et al., 1998). In humans, FGF14 is expressed in similar
CNS regions, including the hippocampus, cerebral cortex (temporal
lobe), putamen and cerebellum (Wang et al., 2002). The functions of
FGF14 are not understood; however, expression patterns suggest
that FGF14 may regulate CNS development and/or neuronal
physiology (Munoz-Sanjuan et al., 2000; Smallwood et al., 1996;
Wang et al., 2002; Yamamoto et al., 1998). FGFs 11–14 lack
recognizable amino-terminal signal sequences, are not secreted from
cells, and do not bind or activate classical tyrosine kinase FGF
receptors (Olsen et al., 2003; Ornitz and Itoh, 2001; Smallwood
et al., 1996). Several members of the FGF11–14 subfamily interact
with the mitogen-activated protein kinase (MAPK) scaffolding
protein islet brain 2 (IB2, also called JIP2) and the carboxy terminal
tail of several voltage-gated sodium (Nav) channels (Liu et al., 2001,
2003; Lou et al., 2005; Schoorlemmer and Goldfarb, 2002;
Wittmack et al., 2004). It is not known whether interactions between
FGF14 and IB2 contribute to its role in neurophysiologic function.
However, the interaction of FGF14 with Nav channels may regulate
neuronal excitability (Lou et al., 2005). To understand further the
neurophysiological function of FGF14 and to gain insight into the
mechanism of the human mutations in FGF14, we investigated
synaptic transmission and synaptic plasticity in Fgf14−/− mice. Here
we show reduced hippocampal short-term and long-term potentiation (LTP) in Fgf14−/− mice, a decreased number of synaptic vesicles
docked at presynaptic CA1 synapses, a significant synaptic fatigue/
depression during high/low-frequency stimulation and a reduction
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
in mEPSC frequency, but not amplitude, in cultured hippocampal
neurons derived from Fgf14−/− mice. Taken together, these findings
provide new insights into the etiology of cognitive deficits in
humans with mutations in FGF14 and suggest a novel role for
FGF14 in regulating synaptic plasticity in hippocampal neurons.
Results
Expression of FGF14 in the hippocampus
Previous reports showed that Fgf14 is widely expressed in the
developing and adult CNS (Smallwood et al., 1996; Wang et al.,
2002). To identify the precise location of Fgf14 expression in
the adult hippocampus, in situ hybridization and β-galactosidase
(β-gal) staining were used on thick floating brain slices of 2- to
3-month-old WT and Fgf14−/+ mice. In the hippocampus, the
FGF14-β-gal expression pattern was consistent with expression in
both principal cells and interneurons of CA1, CA3 and the dentate
gyrus (DG). Most intense staining was localized to the CA
pyramidal cell layers (stratum pyramidal, s.p.), granule cell layer of
the dentate gyrus and mossy cells or interneurons in the dentate
hilus (h) (Fig. 1A). A few scattered interneurons in the stratum
oriens (s.o.), stratum radiatum (s.r.) and stratum lacunosum
moleculare (s.l-m.) of CA1 through CA3 and in the stratum
moleculare (s.m.) and stratum granulosum (s.gr.) of the DG were
also stained (Fig. 1A). High resolution in situ detection of the
Fgf14 mRNA in 2- to 3-month-old WT mice showed a regional
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distribution pattern closely paralleling the β-gal expression pattern
seen in Fgf14−/+ mice (Fig. 1B).
Impaired LTP in Fgf14−/− mice
Because a major function of the hippocampus involves learning
and memory, the expression pattern of FGF14 and the human
cognitive impairments associated with mutation in FGF14
suggested a role for this protein in hippocampal function. We thus
investigated whether mice lacking FGF14 showed alterations in
hippocampal LTP. LTP was measured following two protocols to
induce conditioning in Schaffer collateral-CA1 synapses in
hippocampal slices from adult WT and Fgf14−/− mice. Successful
induction and maintenance of LTP were considered achieved when
the average field excitatory postsynaptic potentials (fEPSP) slope
at 55 to 60 min after tetanic stimulation was at least 20% above
baseline. In the first set of stimulus conditions, Schaffer collateralCA1 fEPSPs were recorded in hippocampal slices after a single 1-s
tetanus stimulation at 100 Hz. The average fEPSP slope, as a
percentage of baseline over the 55- to 60-min interval posttetanization, demonstrated impaired maintenance of LTP in Fgf14−/−
mice (109 ± 12%; n = 8 slices; 4 mice) compared to control mice
{129 ± 9%; n = 7 slices; 4 mice; F(1,6) = 7.2, p = 0.02} (Figs. 2A, B).
Even following maximal induction stimuli, successful maintenance
of LTP 55–60 min after induction was not achieved in Fgf14−/−
slices {fEPSP slope 55–60 min after LTP induction: Fgf14−/−;
92 ± 8%; n = 5 slices; 5 mice; WT, 165 ± 24%; n = 7 slices; 5 mice;
F(1,58) = 37, p < 0.001} (Figs. 2C, D). These data show that
FGF14 deficiency is associated with impaired synaptic plasticity,
suggesting that impaired LTP may account for the cognitive deficits
in humans harboring mutations in FGF14.
Altered basal synaptic transmission in Fgf14−/− mice
To characterize whether basal synaptic function contributed to
the impairment of induction of LTP in Fgf14−/− mice, presynaptic
fiber volley (FV) and fEPSPs from hippocampal Schaffer
collateral-CA1 synapses were recorded following different stimulation intensities. The amplitude of the FV, which is a measure of
the number of recruited presynaptic neurons (axons), was
examined to provide an accurate indication of basal synaptic
transmission in acute hippocampal slices. In Fgf14−/− hippocampal
slices, the amplitude of the FV was not significantly different from
that of WT mice (p = 0.9) (Fig. 3A), suggesting that lack of Fgf14
does not significantly affect the number of afferent axons. Input/
Output (I/O) curves, determined by plotting the amplitude of FV
versus the fEPSP slope (Fig. 3B) and I/O curves, measured by
plotting the stimulation intensities versus the corresponding fEPSP
slope (Fig. 3C), were significantly different between Fgf14−/− and
WT mice; {F(1,162) = 4.6, p = 0.03} and {F(1,155) = 5.5, p = 0.02},
respectively, suggesting that a lack of FGF14 alters basal synaptic
transmission in Schaffer collateral-CA1 synapses.
Impaired short-term plasticity in Fgf14−/− mice
Fig. 1. FGF14 expression in the hippocampus. (A) FGF14-β-gal expression
pattern in a hippocampal coronal section showing expression in both
principal cells and interneurons of CA1 through CA3 and the dentate gyrus
(DG) in Fgf14−/+ mice. (B) In situ detection of the Fgf14 mRNA in WT
mice showing similar patterns of expression as seen for FGF14-β-gal. h,
Mossy cells in dentate hilus; s.gr., stratum granulosum; s.l-m., stratum
lacunosum moleculare; s.m., stratum moleculare; s.o., stratum oriens; s.p.,
stratum pyramidal; s.r., stratum radiatum. Scale bar = 200 μm.
The observed impairment of LTP and basal synaptic transmission
in Fgf14−/− mice suggested that FGF14 is required for the normal
function of hippocampal synapses. To further address this, we tested
the effect of lack of FGF14 on short-term potentiation (STP).
Paired pulse facilitation, one form of STP that measures
transient enhancement of neurotransmitter release induced by two
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Fig. 2. Impaired LTP in Fgf14−/− mice. LTP and PTP in Fgf14−/− and WT mice measured in hippocampal slices at Schaffer collateral/CA1 synapses. (A) LTP
was induced with a 1-s tetanic stimulus at 100 Hz and an intensity that evoked 50% of a maximal EPSP. The fEPSPs slopes show significant differences between
Fgf14−/− mice (●) and WT controls (○). Representative traces are shown for WT (+/+) and Fgf14−/− (−/−) mice at baseline (Trace 1) and 60 min (Trace 2) after
tetanic stimulation. Scale bars equal 1 mV and 10 ms. (B) PTP and LTP in Fgf14−/− mice (solid bar) and WT control mice (open bar) measured at 1–7 min and
55–60 min, respectively. At 1–7 min PTP was significantly lower in Fgf14−/− mice (130 ± 11%; n = 8 slices; 4 mice) compared to control mice (166 ± 6%; n = 7
slices; 4 mice; p = 0.03, data were analyzed with a nonlinear exponential mixed model). At 55–60 min LTP was significantly lower in Fgf14−/− mice (109 ± 12%;
n = 8 slices; 4 mice) compared to control mice (129 ± 9%; n = 7 slices; 4 mice; p = 0.002, data were analyzed with a nonlinear exponential mixed model). (C)
Saturating LTP induction stimuli, consisting of two 1-s long 100-Hz stimuli 20 s apart (each arrow), repeated 4 times at 6-min intervals, results in LTP
maintenance in WT slices (○), but not in Fgf14−/− slices (●) (WT, 7 slices, 5 mice; Fgf14−/−, 5 slices, 5 mice). (D) Cumulative data from C demonstrate
significantly greater LTP 55–60 min after LTP induction in WT compared to Fgf14−/− slices (p < 0.001, ANOVA). *p < 0.05, **p < 0.01, ANOVA.
closely spaced stimuli, is thought to be presynaptically mediated
(Schulz et al., 1994). Paired pulse facilitation was similar in
Fgf14−/− mice and WT controls given an interstimulus interval
(ISI) of 10 to 100 ms (p = 0.8) (Fig. 3D). This suggests that FGF14
does not affect transient enhancement of neurotransmitter release
in the early STP facilitation phase (ten to hundreds of
milliseconds). However, post-tetanic potentiation (PTP), another
form of STP that is thought to rely on presynaptic function, may
represent transient increases in neurotransmitter release caused by
the loading of the presynaptic terminals with calcium ions after
tetanic conditioning (Kamiya and Zucker, 1994; Wang et al.,
2004). The magnitude of PTP (measured over a 1- to 7-min
interval following tetanization) was significantly attenuated in
Fgf14−/− mice (130 ± 11%; n = 8 slices; 4 mice) compared to
control mice {166 ± 6%; n = 7 slices; 4 mice; F(1,6) = 8.7, p = 0.02}
(Figs. 2A, B), suggesting a decrease in neurotransmitter release in
the later STP period (several minutes).
Together, these data show that FGF14 is required for synaptic
plasticity, suggesting that an impairment of presynaptic neuro-
transmitter release could be responsible for LTP deficits observed
in Fgf14−/− mice.
Normal hippocampal anatomy in Fgf14−/− mice
Changes in hippocampal function could result from changes in
hippocampal anatomy. Hippocampal anatomy was examined with
histochemical stains and antibodies, respectively. Nissl-stained
sections showed normal patterns, cytoarchitecture, and cell/neurite
density both in CA1 through CA3 and the dentate gyrus in WT and
Fgf14−/− mice (Figs. 4A, A′). Furthermore, immunohistochemical
analysis showed no differences between Fgf14−/− and WT mice in
the distribution of synaptophysin (Figs. 4B, B′), which preferentially stains synapses of the Schaffer collateral pathway from CA3
to CA1 (Pittenger et al., 2002), GAP43 (Figs. 4C, C′), which
preferentially stains the perforant pathway from the entorhinal
cortex to CA1 (Pittenger et al., 2002), or calbindin (Figs. 4D, D′),
which preferentially stains mossy fiber projections to CA3
(Minichiello et al., 1999).
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
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Fig. 3. Abnormal basal synaptic transmission and STP in Fgf14−/− mice. (A–C) Effects of FGF14 deficiency on excitatory synaptic transmission in hippocampal
CA1 synapses in hippocampal slices from 2- to 3-month-old mice. Fiber volleys and fEPSPs were recorded in the CA1 region after stimulation of Schaffer
collateral neurons. The data (mean ± SEM) from multiple recordings of Fgf14−/− mice (●) and WT control mice (○) were pooled. (A) Plot of presynaptic fiber
volley amplitude versus stimulus intensity (WT, 16 slices, 3 mice; Fgf14−/−, 14 slices, 3 mice). No significant difference was seen between Fgf14−/− and WT
mice (linear mixed model analysis, p = 0.9). (B) Input/Output plot of fEPSP slope versus presynaptic fiber volley (WT, 12 slices, 3 mice; Fgf14−/−, 8 slices, 3
mice). Each point represents data from individual slices. The lines represent the best fit linear regression (Fgf14−/−, y = 1.06x; WT, y = 1.29x). A significant
difference is observed between Fgf14−/− mice and WT (linear mixed model, p = 0.03). (C) Input/Output plot of fEPSP slope versus stimulus intensity (WT, 15
slices, 3 mice; Fgf14−/−, 15 slices, 3 mice). A significant difference was observed between Fgf14−/− and WT mice (linear mixed model, p = 0.02). (D) Paired
pulse facilitation showing no significant difference between Fgf14−/− and WT mice (WT, 10 slices, 3 mice; Fgf14−/−, 11 slices, 3 mice; linear mixed model,
p = 0.9). Traces show responses during paired pulse facilitation at 50-ms interpulse intervals in WT (+/+) and Fgf14−/− (−/−) mice. Scale bars equal 1 mV and
10 ms.
Decreased docked and reserve pool synaptic vesicles in the
hippocampus of Fgf14−/− mice
number of docked synaptic vesicles and the size of the reserve
pool.
To understand the mechanisms underlying impaired synaptic
function in Fgf14−/− mice, synapse morphology was analyzed by
TEM. The general appearance of presynaptic terminals, spines, and
postsynaptic densities at excitatory asymmetric synapses in the
stratum radiatum (s.r.) of CA1 were similar in Fgf14−/− and WT
mice (Figs. 5A, B), as were the numbers of excitatory synapses and
the postsynaptic densities (Table 1). However, there was a trend
towards increased active zone width in Fgf14−/− synapses
compared to controls (Table 1). Interestingly, quantization of
synaptic vesicle numbers showed a significant reduction in the
number of docked synaptic vesicles per micrometer of active zone
length (p = < 0.0001), and a significant reduction in the number of
reserve pool synaptic vesicles per terminal (p = 0.002) in Fgf14−/−
mice (Figs. 5C, D; Table 1). In summary, FGF14 affects the
Synaptic fatigue/depression in Fgf14−/− mice
Impairments of basal synaptic transmission and decreases in the
number of reserve pool and docked synaptic vesicles in Fgf14−/−
mice suggest that FGF14 may affect synaptic transmission by a
presynaptic mechanism. To test this hypothesis, release probability
of the readily releasable pool (RRP) and the reserve pool of
synaptic vesicles at CA1 synapses were investigated, and
frequency and amplitude of miniature AMPA-mediated mEPSCs
were measured in hippocampal neuronal cultures isolated from
Fgf14−/− mice.
The synaptic response to a high-frequency repetitive stimulation (HFS) (100 Hz, 40 pulses), which is thought to relate to RRP
quanta and correspond to morphologically defined docked vesicles
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M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
Fig. 4. Normal gross hippocampal anatomy of Fgf14−/− mice. (A, A′) Normal hippocampal cytoarchitecture revealed by Nissl staining (coronal sections) in
Fgf14−/− and WT mice. Scale bar = 500 μm. (B, B′) Synaptophysin immunoreactivity in Fgf14−/− and WT mice (sagittal sections), highlighting the stratum
radiatum of CA1. Schaffer collateral projection from CA3 is indicated by the arrow. (C, C′) GAP43 immunoreactivity in Fgf14−/− and WT mice (coronal sections)
to preferentially stain the perforant pathway projections from the entorhinal cortex to the stratum lacunosum moleculare of CA1 (arrow). (D, D′) Calbindin-D28 k
immunoreactivity in Fgf14−/− and WT mice (coronal sections) to highlight mossy fiber projections from the dentate gyrus to CA3 pyramidal cells (arrow).
(Cabin et al., 2002), was determined. Fgf14−/− and WT control
mice differed in the amplitude of the fEPSPs slope {t(8) = −9.4,
p < 0.0001} but exhibited a similar continuous reduction in fEPSPs
rate in CA1 synapses during HFS {t(8) = − 0.15, p = 0.9} (Figs. 6A,
B). These data not only suggest RRP impairment resulting in
synaptic fatigue in Fgf14−/− mice, but also are consistent with the
morphological data that Fgf14−/− hippocampal synapses have a
decreased number of docked vesicles in CA1. Next, synaptic
responses to a prolonged repetitive low-frequency stimulation train
(14 Hz, 300 pulses), which are typically used to test the size of the
reserve pool of vesicles (Bamji et al., 2003), were determined.
When prolonged repetitive low-frequency stimulation at CA1
synapses was administered, the depletion of RRP vesicles is
usually faster than the replenishment by the reserve pool of
vesicles, leading to a gradual decline in fEPSP slope. Synaptic
responses to prolonged repetitive low-frequency stimulation
gradually decreased over time in both WT and Fgf14−/− mice;
however, in Fgf14−/− hippocampal slices, the decline was
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
371
Fig. 5. Analysis of asymmetric synapses in Fgf14−/− mice. (A–B) Transmission electron microscopy (TEM) images of asymmetric excitatory synapses in the
CA1 stratum radiatum showing normal gross structural features in both WT (A) and Fgf14−/− mice (B). Scale bar = 500 nm. (C–D) TEM images displaying fewer
docked vesicles at the active zones on CA1 asymmetric synapses in Fgf14−/− mice (D) compared to WT mice (C). Scale bar = 250 nm.
significantly greater than in WT slices following the 50th stimulus
trial {0–50: not significant, p = 0.4; 50–150: significant difference,
F(1,264) = 11, p < 0.001; 150–300: significant difference, F(1,404) =
5.5, p = 0.02} (Figs. 6C, D). These results are consistent with the
morphological data showing that Fgf14−/− CA1 synapses have
fewer non-docked vesicles, and suggest that a function of FGF14
may be to regulate the size of the reserve vesicle pool, vesicle
mobilization, or the trafficking of synaptic vesicles from the reserve
vesicle pool to the RRP at presynaptic active zones.
Whole-cell patch-clamp recordings were then used to examine
frequency and amplitude of AMPA-mediated mEPSCs derived
from control and Fgf14−/− cultured hippocampal neurons (Figs.
6E–H). No difference in control versus Fgf14−/− neurons was
observed in mEPSC amplitude (p = 0.9, Fig. 6G), rise time (1.7 ±
0.1 ms, n = 8 WT cells, 1.5 ± 0.1 ms, n = 13 Fgf14−/− cells; p = 0.3)
or decay time (6.1 ± 0.3 ms, 5.4 ± 1.7 ms; p = 0.3). However,
Fgf14−/− hippocampal neurons showed a significant reduction in
Table 1
Morphological analyses of CA1 synapses in Fgf14−/− mice
Parameter
WT a
Fgf14−/− a
P value b
Synapse density c
Active zone width d
Docked SV e
Reserve pool vesicles f
41.2 ± 3.9
275 ± 24
18.6 ± 1.5
19.7 ± 1.7
40.3 ± 3.2
291 ± 21
12.5 ± 1.2
15.8 ± 1.0
ns
ns
<0.0001
<0.003
a
Mean ± SD.
Student's t test, 251 synapses for WT mice (n = 6), 255 synapses for
Fgf14−/− mice (n = 5), ns, not significant.
c
Number of postsynaptic densities (PSDs) per 100 μm2. Total area from
WT (n = 6), 4694 μm2; Fgf14−/−(n = 5), 6035 μm2.
d
Width of the PSD (nm).
e
Docked SV within 50 nm of the presynaptic active zone normalized to
the active zone width (vesicle number/μm).
f
Reserve pool SV located 50–550 nm from the presynaptic active zone
(vesicle number/terminal).
b
mEPSC frequency compared to WT hippocampal neurons
(p = 0.004, Fig. 6H). Together, these results are consistent with a
major presynaptic contribution to the underlying mechanism of
defects in excitatory synaptic transmission in Fgf14−/− mice.
Alteration of synaptic protein distribution in Fgf14−/− mice
To further investigate the possibility that synaptic vesicle
mobilization and docking could be affected by FGF14, the level
of several SNARE (soluble N-ethylmaleiamide-sensitive factor
attachment protein receptor) complex proteins involved in vesicle
docking and fusion (O’Connor et al., 1994; Sudhof, 2004) were
examined in whole hippocampal lysates and in hippocampal
synaptoneurosomes. These included proteins localized to synaptic
vesicles (v-SNAREs, synaptobrevin and synaptotagmin), the
presynaptic plasma membrane (t-SNAREs, syntaxin-1 and SNAP25), and synaptophysin, a major integral membrane protein on
synaptic vesicles. Whole hippocampal lysates showed a significant
decrease in the level of synaptobrevin in Fgf14−/− mice (68 ± 4% of
WT control, p < 0.0001, n = 4, StudentTs t test) (Figs. 7A,B). In
synaptoneurosomes, synaptobrevin, synaptophysin and syntaxin-1
were significantly decreased in Fgf14−/− compared to control mice
(76 ± 5%, p = 0.003; 68 ± 9%, p = 0.01; 82 ± 5%, p = 0.01, respectively, n = 4, StudentTs t test) (Figs. 7A, B). Together, these data
support a model in which, in the absence of FGF14, there is selective
reduction in synaptobrevin in cell bodies, axons and nerve terminals
of hippocampal neurons of Fgf14−/− mice.
Discussion
FGF14 is expressed in the hippocampus and neocortex, and
humans with mutations in Fgf14 exhibit impaired cognitive
functions. We therefore hypothesized that Fgf14−/− mice may also
have compromised cognitive capabilities. In this study we showed
that Fgf14−/− mice have deficits in basal synaptic function and
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plasticity, further extending the similarities between Fgf14−/− mice
and FGF14 (F145S) and FGF14 (D163fsX12) humans. In parallel
studies, we have shown that Fgf14−/− mice have deficits in learning
and memory when tested in the Morris Water maze (Wozniak et al.,
in press). To begin to address the neurophysiological function of
FGF14, we showed that Fgf14−/− mice have decreased LTP at
Schaffer collateral-CA1 synapses and have impaired synaptic
transmission and altered presynaptic vesicle trafficking, docking
and synaptic protein expression. These studies implicate FGF14 as
an important regulator of neuronal function that could contribute to
both human cognition and sensorimotor function.
The predominantly presynaptic abnormalities most likely
account for the impaired long-term synaptic potentiation (LTP)
that we observed in Fgf14−/− hippocampal slices. Although
postsynaptic mechanisms are critical for normal Schaffer collateral
LTP, presynaptic mechanisms are also important (Choi et al., 2003;
Lisman, 2003; Luscher et al., 2000; Sanes and Lichtman, 1999;
Zakharenko et al., 2001). Because normal hippocampal synaptic
function and plasticity are important for learning and memory
(Bliss and Collingridge, 1993; Milner et al., 1998), we speculate
that these physiological alterations could contribute to the
cognitive impairments in humans with mutations in FGF14.
Fgf14−/− mice enable a direct test of this hypothesis and underscore
the relevance of this mouse model in understanding the role of
FGF14 in neuronal function. Synaptic dysfunction in other brain
regions, such as the cerebellum or basal ganglia, may explain the
ataxia/dyskinesia phenotype displayed by humans with mutations
in FGF14 (Brusse et al., 2006; Dalski et al., 2005; Van Swieten
et al., 2003) and mice lacking Fgf14 (Wang et al., 2002).
The synaptic vesicle cycle involves transport of vesicles to
nerve terminals, docking, priming, Ca2+-triggered fusion, endocytosis and the formation of new vesicles (Sudhof, 2004). Fgf14−/−
hippocampal slices showed a significant synaptic fatigue/depression during high/low-frequency stimulation, suggesting insufficient
numbers of available vesicles. TEM analysis showed fewer docked
and reserve vesicles, suggesting that the total numbers of releasable
vesicles (both RRP and reserve pool) were decreased in CA1 in
Fgf14−/− mice. Analysis of AMPA-mediated mEPSCs has often
been used to determine the relative pre- and postsynaptic
contribution to synaptic transmission (Bekkers and Stevens,
1990; Isaac et al., 1996). Changes in the amplitude of mEPSCs
are usually attributed to postsynaptic modulation in receptor
number and/or function, while changes in the frequency of
mEPSCs have traditionally been associated with presynaptic
variables, such as the probability of quanta release (Pr) or number
of vesicles released. Fgf14−/− cultured hippocampal neurons
showed a decrease in mEPSC frequency, whereas mEPSC
amplitude was unchanged, supporting a presynaptic defect in
Fgf14−/− mice.
Also consistent with a major presynaptic defect in Fgf14−/−
mice, protein expression analysis suggested reduced levels of
several synaptic proteins involved in presynaptic membrane fusion.
Knockout studies in mice show that deletion of multiple SNARE
proteins can lead to decreased LTP (Janz et al., 1999). Furthermore,
synaptobrevin has been shown to be important for vesicle
endocytosis (Deak et al., 2004). Thus, consistent with classical
quantal theory (Del Castillo and Katz, 1954), decreased numbers of
docked synaptic vesicles correlate with the decreased vesicle
release observed in Fgf14−/− mice. Additionally, and as we
observed in Fgf14−/− hippocampal slices, this would impair posttetanic synaptic potentiation (PTP), a form of short-term synaptic
plasticity that depends on the probability of vesicle release (Zucker
and Regehr, 2002). In LTP, the probability of glutamate release is
increased (Schulz, 1997), suggesting increased quantal size,
increased probability of quanta release (Pr) or increased numbers
of release sites (N) following LTP induction/maintenance. The
underlying mechanism may involve enhanced synaptic vesicle
fusion pore kinetics (Kaneko and Takahashi, 2004), or enhanced
glutamate release at individual synapses (Zakharenko et al., 2001).
Taken together, these data suggest that FGF14 contributes to shortand long-term synaptic plasticity through a presynaptic mechanism
by modulating vesicle release probability.
The molecular mechanism(s) by which FGF14 affects presynaptic
function is not known. However, recent studies show that FGF14
interacts with and regulates neuronal voltage-gated sodium channels
(Lou et al., 2005), and related FGFs bind to and regulate TTXsensitive and -insensitive voltage-gated sodium channels (Liu et al.,
2001, 2003). Therefore, one possibility is that altered neuronal activity
due to sodium channel dysfunction in Fgf14−/− mice may influence
presynaptic protein expression and synaptic vesicle function.
Interestingly, some of the presynaptic alterations we observed in
Fgf14−/− mice resemble those seen in mice lacking BDNF. BDNF
facilitates tetanus-induced LTP by reducing presynaptic fatigue, and
BDNF-deficient mice exhibit reduced synaptic vesicle associated
protein expression, reduced synaptic vesicle docking and impaired
early-phase LTP at Shaffer collateral synapses (Pozzo-Miller et al.,
1999). Finally, BDNF may interact with TTX-insensitive sodium
channels through its receptor TrkB (Blum et al., 2002). Whether or not
FGF14 possesses functions converging or complementing BDNF
functions is an important area for future studies.
Experimental methods
Mice
Fgf14−/− mice (Wang et al., 2002) were maintained on an inbred C57/
BL6J background (greater than ten generations of backcrossing to C57/
BL6J). All genotypes described were confirmed by PCR analysis (Wang
Fig. 6. Impaired synaptic vesicles function and reduced miniature EPSCs frequency in Fgf14−/− mice. (A–B) Synaptic fatigue in CA1 synapses assessed during a
brief high-frequency stimulation (HFS, 100 Hz, 40 pulses) (WT, 6 slices, 3 mice; Fgf14−/−, 8 slices, 3 mice). (A) In Fgf14−/− mice (●) and WT controls (○), the
fEPSPs slope in CA1 synapses shows a similar continuous reduction in during HFS {nonlinear exponential mixed model, p = 0.9}, however, Fgf14−/− mice show
a reduced amplitude relative to WT controls (p < 0.0001). (B) Examples of EPSPs elicited by a train of HFS showing significant synaptic fatigue in Fgf14−/−
mice. Scale bars equal 1 mV and 50 ms. (C–D) Synaptic depression in CA1 synapses during prolonged repetitive stimulation (14 Hz, 300 pulses) (WT, 7 slices,
3 mice; Fgf14−/−, 7 slices, 3 mice). (C) fEPSPs slope plotted against stimulus number. Fgf14−/− mice (●) have a significantly decreased fEPSPs slope compared
to WT controls (○) following the 50th stimulus. (D) Representative single EPSPs at the first stimulus (1), the tenth stimulus (2), the one hundredth stimulus (3)
and the three hundredth stimulus (4) are shown. The slopes of field EPSPs (mean ± SEM) during the entire recording were normalized to the first EPSP slope in
each recording. Scale bars equal 0.5 mV and 5 ms. (E–F) Voltage-clamp recordings at − 80 mV showing AMPA-mediated mEPSCs in WT (E) and Fgf14−/− (F)
cultured neurons at DIV 11–13. (G–H) Average mEPSC amplitude (G) and frequency (H) isolated from WT (n = 15 neurons, 763 mEPSCs) and Fgf14−/− (n = 20
neurons, 967 mEPSCs) neurons (mean ± SEM, **p < 0.01, T-test).
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
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M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
Fig. 7. Synaptic protein expression in the hippocampus and in hippocampal synaptoneurosomes. (A) Immunoblot detection of synaptic proteins in whole
hippocampal homogenates (HOM) and in hippocampal synaptoneurosomes (SNS) from Fgf14−/− mice and WT controls. (B) Quantization of expression levels of
synaptic protein in HOM and SNS (WT, 4 mice; Fgf14−/−, 4 mice). Mean band intensity for Fgf14−/− extracts is plotted as a percentage of mean WT band
intensity (mean ± SEM). Note the significant reduction in the level of synaptobrevin in HOM (68 ± 4% of control, *p = 0.0007). In SNS, the levels of
synaptobrevin, syntaxin 1 and synaptophysin are significantly decreased in Fgf14−/− mice compared to WT control mice {76 ± 4% (**p = 0.003), 76 ± 4%
(*p = 0.014) and 67 ± 7% (*p = 0.01), respectively}.
et al., 2002). Littermate or age-matched controls were used for all
experiments.
Perfusion and histochemical analysis
Fgf14−/− and wild-type (WT) mice were anesthetized with sodium
pentobarbital (60 mg/kg, i.p.) and transcardially perfused with a vascular
rinse of 0.9% NaCl followed by ice-cold 4% paraformaldehyde in 0.1 M
phosphate buffer (pH 7.4). Brains were dissected, postfixed in the same
solution overnight at 4 °C, and cryoprotected in 30% sucrose in 0.1 M
phosphate buffer until they sank. After embedding in O.C.T. compound,
brains were cryosectioned at 14 μm or 30 μm and collected in PBS for
immunostaining and in situ hybridization. For general histology, sections
were stained with 0.1% cresyl violet according to standard procedures.
X-gal staining
To analyze the expression of the reporter protein, FGF14-β-gal, 500-μm
sections were cut with a vibratome in ice-cold X-gal rinse buffer (0.1 M
PBS and 2 mM MgCl2). Sections were fixed for 1 h in 0.5% glutaraldehyde
in X-gal rinse buffer at 4 °C. The sections were then immersed in X-gal
staining solution (20 mM potassium ferricyanide, 20 mM potassium ferrocyanide, 0.02% Nonidet P-40, 0.01% sodium deoxycholate, and 1 mg/ml 5bromo-4-chloro-3-indoyl-β-D-galactosidase made up in rinse buffer,
pH 7.6) overnight at 37 °C to reveal transgene-expressing cells in the
brain. After X-gal staining, the sections were cryoprotected in 30% sucrose
in 0.1 M phosphate buffer until they sank, embedded in O.C.T. compound,
and cryosectioned at 14 μm. After rinsing briefly in PBS, sections were
dehydrated in an ethanol series (70%, 95%, and 100% ethanol), then in
xylene, and then mounted in Permount (Fisher Scientific) or DPX mount
(BDH Chemicals).
Immunohistochemistry and RNA in situ hybridization
For immunohistochemistry, free-floating brain sections (14 μm) were
washed in PBS, blocked in a solution of 7.5% goat serum (Sigma) and 0.25%
Triton X-100 (TX-100; Sigma) in PBS. Sections were then incubated at 4 °C
overnight with the primary antibodies diluted in 1% goat serum/0.25% TX100 in PBS. The following specific antibodies were used: mouse anticalbindin D-28K (Sigma) at 1:5000, rabbit anti-GAP43 (Chemicon) at
1:1000, mouse anti-synaptophysin (Chemicon) at 1:1000. Sections were then
washed three times for 5 min at room temperature in PBS before application
of secondary antibodies. Sections were then incubated in the appropriate
secondary goat antibodies (labeled with Alexa 488 or Texas Red) against
either mouse or rabbit at 1:100 (Molecular Probes) dilution in the same
solution as the primary antibodies. For ABC immunohistochemistry,
immunoreactivity was performed using the Elite ABC kit (Vector
Laboratories). The immunoreactivity was visualized with DAB.
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
For in situ hybridization, a 550-bp RNA probe (Wang et al., 2000) was
labeled with digoxigenin following the manufacturerTs protocol (Roche
Diagnostics, IN). Free-floating brain sections (30 μm) were washed twice in
PBS and treated with freshly prepared 10 μg/ml proteinase K (Invitrogen,
Carlsbad, CA) at 37 °C. After acetylation, sections were incubated in
hybridization buffer containing 0.2 μg/ml digoxigenin-labeled riboprobes at
43 °C overnight. Hybridized sections were washed by successively
immersing them in 4× SSC (150 mM NaCl, 15 mM sodium citrate, pH
7.0, room temperature), 2× SSC containing 50% formamide (50 °C,
30 min), 2× SSC (37 °C, 10 min), 2× SSC containing 20 μg/ml RNase A
(37 °C, 30 min), 2× SSC (37 °C, 20 min), and 0.1× SSC (room temperature,
10 min). Controls hybridized without primary probe showed no signal. The
hybridization signals were detected with digoxigenin detection reagents
(Roche Diagnostics, Indianapolis, IN). After rinsing in PBS, sections were
mounted and analyzed with a Zeiss Axioskop microscope.
Transmission electron microscopy
Two to three-month-old mice were used for transmission electron
microscopy (TEM) analysis. Mice were anesthetized with sodium
pentobarbital and perfused with PBS (pH 7.4) followed by 2%
paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer (pH
7.4). The heads were removed and stored overnight at 4 °C in 0.1 M
phosphate buffer. The brains were dissected and coronal sections were cut
at 500 μm with a vibratome in 0.1 M PBS. The hippocampal CA1 region
was dissected from appropriate sections and then was postfixed for 1 h with
1% osmium tetroxide in 100 mM cacodylate buffer (pH 7.4), dehydrated
using a series of ethanol dilutions, rinsed in propylene oxide and embedded
in Epon 812. Ultrathin sections (60–80 nm) were cut and stained with
uranyl acetate and lead citrate. Complete profiles of nonperforated
asymmetric synapses on dendritic spines in the stratum radiatum of CA1
were photographed using a digital camera in a Hitachi 7500 electron
microscope operated at 80 kV at a final magnification of 8000× or 50,000×.
A total area of 4694 μm2 from WT and 6035 μm2 from Fgf14−/− were
counted for quantification of synapse density. A total of 251 asymmetric
synapses from 6 WT and 255 synapses from 5 Fgf14−/− mice were
analyzed. Synapses were quantified for length of the active zone, the
number of reserve pool small synaptic vesicles (∼ 50 nm vesicles located
50–550 nm from the presynaptic active zone) and the number of docked
vesicles (∼50 nm vesicles located within 50 nm of the presynaptic active
zone) (Bamji et al., 2003; Pozzo-Miller et al., 1999; Schoch et al., 2002).
Western blot and biochemical analysis of synaptic proteins
Wild-type and Fgf14−/− mice were anesthetized, hippocampal tissues
were dissected and homogenized in 1% SDS. Synaptoneurosomes from at
least three pairs of 2- to 3-month-old WT and Fgf14−/− mice were prepared
as described (Scheetz et al., 2000). Twenty micrograms of soluble protein
per lane was subjected to Western blotting and detected with antibodies
against synaptotagmin (Sigma) at 1:5000, synaptophysin (Chemicon) at
1:2000, synaptobrevin (Chemicon) at 1:1000, SNAP-25 (Chemicon) at
1:2000 and syntaxin-1 at 1:1000 (Sigma). The blots were developed using
horseradish peroxidase-conjugated secondary antibodies at 1:5000 and
enzyme-linked chemiluminescence (PerkinElmer or Pierce). An antibody
against β-actin (Sigma) at 1:2000 was used sequentially on the same blot to
control for loading. Signal intensities were quantified using the Quantity
One Software (BioRad). All data are presented as mean ± SEM and statistical
significance was determined either by StudentTs t test or ANOVA.
Preparation of hippocampal slices and electrophysiology
Two- to three-month-old WT and Fgf14−/− littermates were used in all
electrophysiological experiments. After halothane anesthesia, decapitation,
and removal of the brain, transverse acute hippocampal slices (400 μm)
were cut with a vibratome in ice-cold artificial CSF (ACSF) containing
124 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 2 mM MgSO4, 1.25 mM
NaH2PO4, 22 mM NaHCO3, 10 mM glucose. The protocol for electrical
375
stimulation and recordings was as described (Yamada et al., 2004). The
slices were kept for at least 1 h before recording at room temperature in
95% O2 and 5% CO2 bubbled ACSF. Recordings were performed in the
same solution, in a submerged chamber, at 30 °C. Recordings of field EPSP
(fEPSP) were performed in the medial CA1 subfield with glass pipettes
filled with 2 M NaCl (5–10 MΩ DC resistance), connected to an
Axoclamp2A amplifier (Molecular Devices, Union City, CA). Schaffer
collaterals were stimulated with a bipolar electrode placed in the lateral
CA1 subfield. The duration of the pulses was 0.1- to 0.2-ms constant
current, and the stimulation strength was set to provide baseline fEPSPs
with an amplitude of ∼ 50% of the maximum amplitude. For induction of
LTP experiments, the average slope of the fEPSPs during a 20-min baseline
recording period was used to normalize the fEPSP slopes for comparison
among slices. Average responses (means ± SEM) are expressed as percent of
baseline response. Two protocols were used to induce LTP: 100 Hz tetanus
for 1 s (100 Hz) and paired 1 s (100 Hz) stimulation (20-s intervals)
repeated four times, with each pair of stimulations separated by 6 min. The
traces were digitized using Clampex 9 and a Digidata 1322A interface and
the data were analyzed using Clampfit 9.0 (PClamp software, Molecular
Devices, Union City, CA).
Neuronal cultures and electrophysiology
Time mated female mice were killed by carbon dioxide asphyxiation
using a protocol approved by the Washington University Animal Studies
Committee. Mouse hippocampal cultures were prepared from E17.5 mouse
embryos, which were isolated, placed on ice, and decapitated. Hippocampi from each embryo were separately dissected and dissociated using
papain (20 U/ml) and trituration through Pasteur pipettes. Neurons were
plated at low density (5 × 105 cells/dish) on poly-D-lysine-coated coverslips
(1 mg/ml) in 60-mm culture dishes in serum-free minimum essential
medium (MEM). After 3–5 h, coverslips containing neurons were inverted
over a glial feeder layer in serum-free MEM with N2 supplements, 0.1%
ovalbumin, and 1 mM pyruvate (N2.1 medium; components from
Invitrogen). The neurons grew over the feeder layer but were kept separate
from the glia by wax dots on the neuronal side of the coverslips. To prevent
the overgrowth of the glia, cytosine arabinoside (5 μM; Calbiochem, La
Jolla, CA, USA) was added to neuron cultures after 1–2 days in vitro (DIV).
Cultures were maintained in N2.1 media for up to 15 DIV, feeding the cells
once per week by replacing one-third of the media per dish.
Whole-cell patch-clamp recordings were made at room temperature
from 11 to 13 DIV isolated neurons cultured from E17.5 mice. Patch
pipettes (5–9 MΩ) were prepared from borosilicate glass using a Sutter
Instrument horizontal puller and filled with internal solution containing the
following (in mM): 140 Cs glucuronate, 10 EGTA, 5 CsCl, 5 MgCl2, and
10 HEPES, pH adjusted to 7.4 with CsOH. Culture dishes were perfused
with TyrodeTs solution containing the following (in mM): 150 NaCl, 4 KCl,
2 MgCl2, 2 CaCl2, 10 glucose, and 10 HEPES, pH adjusted to 7.4 with
NaOH. Recordings were performed using a Multiclamp 700B amplifier
(Molecular Devices, Union City, CA) in voltage-clamp mode. The cells
were held at −80 mV throughout the recordings. Miniature excitatory
postsynaptic currents (mEPSCs) were typically filtered at 2 kHz and
digitized at 5–10 kHz using pClamp9 acquisition software (Molecular
Devices, Union City, CA). Series resistance and input resistance were
measured by applying a 10 mV voltage step at the beginning and at the end
of each experiment. Only cells with stable series and input resistances were
included in this study. Drugs were added directly to the perfusion solution.
Agents used were bicuculline methiodide (10 μM), D,L-2-amino-5phosphopentoic acid (100 μM) and tetrodotoxin (1 μM). Experiments were
collected from two independent neuronal cultures including age-matched
littermate WT controls (Fgf14+/+, n = 12) and Fgf14−/− (n = 20), and one
culture including an age-matched WT control group (n = 3).
The first 33 to 60 consecutive mEPSCs for each cell were analyzed off-line
using Clampfit9 software (Molecular Devices, Union City, CA). mEPSC
amplitude, rise time (10–90%) and decay time (10–90%) were analyzed
manually with cursors. The threshold amplitude for the detection of an event
was adjusted to 5 pA (≥2 SD above noise level). Frequencies were expressed as
376
M. Xiao et al. / Mol. Cell. Neurosci. 34 (2007) 366–377
number of events per second (in Hertz). All data are presented as mean ± SEM.
StudentTs t-tests were performed as appropriate to test for statistical significance.
Statistical analysis
Statistical analyses for the electrophysiological studies were conducted
using SAS institute software, which involved applying linear or nonlinear
mixed models to fit the data. In each experiment, the trends of the response
variables (e.g., the “EPSP slope” in HFS study) changing over the predictor
variables (e.g., the “stimulus numbers” in HFS study) in both Fgf14−/− and
WT mice were modeled by either straight lines or nonlinear curves,
depending on shape of the curves shown in mean plots. The key characteristics of the lines or curves were represented by specific parameters with
group differences in these parameters being evaluated in statistical models.
Each individual mouse (coded as m) was considered as the experimental
unit; when appropriate, slides prepared from each mouse (coded as s(m))
were considered as the sub-sampling units. Both m and s(m) were modeled
as random effects. For all tests, statistical significance was denoted by
p < 0.05.
Acknowledgments
We thank L. Li, A. Saharge, A. Meyenburg and K. Johnson for
their excellent technical assistance, A. DiAntonio for critically
reading the manuscript and J. Huettner for sharing equipment. This
work was supported in part by NIH grant CA60673, AG11355,
funds from the Department of Molecular Biology and Pharmacology, Washington University School of Medicine, the McDonnell
Foundation (K.A.Y.) and a generous contribution from the Virginia
Friedhofer Charitable Trust.
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