Bassoon and the synaptic ribbon organize Ca2+ channels and vesicles to add release sites and promote refilling.

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Neuron
Article
Bassoon and the Synaptic Ribbon
Organize Ca2+Channels and Vesicles
to Add Release Sites and Promote Refilling
T. Frank,1,2,3M.A. Rutherford,1,10N. Strenzke,4,5,10A. Neef,3,10T. Pangr? si? c,1D. Khimich,1A. Fetjova,6E.D. Gundelfinger,6
M.C. Liberman,5B. Harke,7K.E. Bryan,8A. Lee,8A. Egner,7D. Riedel,9,* and T. Moser1,2,3,*
1InnerEarLab, Department of Otolaryngology and Center for Molecular Physiology of the Brain, University of Go ¨ttingen Medical Center,
37099 Go ¨ttingen, Germany
2International Max Planck Research School for Neurosciences, Go ¨ttingen Graduate School for Neurosciences and Molecular Biosciences,
37077 Go ¨ttingen, Germany
3Bernstein Center for Computational Neuroscience, 37073 Go ¨ttingen, Germany
4Auditory Systems Physiology Group, Department of Otolaryngology and Center for Molecular Physiology of the Brain,
University of Go ¨ttingen Medical Center, 37099 Go ¨ttingen, Germany
5Eaton Peabody Laboratory, Massachusetts Eye and Ear Infirmary, Boston, MA 02114, USA
6Department of Neurochemistry and Molecular Biology, Leibniz Institute for Neurobiology, 39118 Magdeburg, Germany
7Department of Nanobiophotonics, Max Planck Institute for Biophysical Chemistry, 37077 Go ¨ttingen, Germany
8Department of Molecular Physiology and Biophysics, University of Iowa, Iowa City, IA 52242, USA
9Laboratory of Electron Microscopy, Max Planck Institute for Biophysical Chemistry, 37077 Go ¨ttingen, Germany
10These authors contributed equally to the work
*Correspondence: driedel@gwdg.de (D.R.), tmoser@gwdg.de (T.M.)
DOI 10.1016/j.neuron.2010.10.027
SUMMARY
At the presynaptic active zone, Ca2+influx triggers
fusion of synaptic vesicles. It is not well understood
how Ca2+channel clustering and synaptic vesicle
docking are organized. Here, we studied structure
and function of hair cell ribbon synapses following
geneticdisruption ofthepresynapticscaffoldprotein
Bassoon. Mutant synapses—mostly lacking the
ribbon—showed a reduction in membrane-proximal
vesicles, with ribbonless synapses affected more
than ribbon-occupied synapses. Ca2+
were also fewer at mutant synapses and appeared
in abnormally shaped clusters. Ribbon absence
reduced Ca2+channel numbers at mutant and wild-
type synapses. Fast and sustained exocytosis was
reduced, notwithstanding normal coupling of the
remaining Ca2+channels to exocytosis. In vitro
recordings revealed a slight impairment of vesicle
replenishment. Mechanistic modeling of the in vivo
data independently supported morphological and
functional in vitro findings. We conclude that
Bassoon and the ribbon (1) create a large number
of release sites by organizing Ca2+channels and
vesicles, and (2) promote vesicle replenishment.
channels
INTRODUCTION
Sensory encoding in the auditory and visual system of verte-
brates relies on transformation of graded receptor potentials
into rates of neurotransmitter release at ribbon synapses. The
synaptic ribbon, an electron-dense structure anchored at the
active zone, tethers a halo of synaptic vesicles (Glowatzki
et al., 2008; Nouvian et al., 2006; Sterling and Matthews,
2005).Aside fromits majorcomponent, RIBEYE/CtBP2 (Khimich
et al., 2005; Schmitz et al., 2000; Zenisek et al., 2004), the ribbon
also contains scaffold proteins such as Bassoon and Piccolo
(Dick et al., 2001; Khimich et al., 2005; tom Dieck et al., 2005).
Genetic disruption of Bassoon perturbs the anchoring of ribbons
totheactivezones(AZs)ofphotoreceptors(Dicketal.,2003)and
cochlear inner hair cells (IHCs) (Khimich et al., 2005). At the IHC
synapse, where the functional effects of Bassoon disruption and
ribbon loss are best studied, fast exocytosis is reduced (Khimich
et al., 2005), and sound encoding by the postsynaptic spiral
ganglion neurons impaired (Buran et al., 2010). Moreover, IHCs
of these Bassoon mouse mutants (BsnDEx4/5) show smaller
Ca2+currents. However, matching Ca2+currents by reducing
the driving force for Ca2+in wild-type IHCs does not equalize
fast exocytosis between wild-type and mutant IHCs. This,
together with an unaltered rate constant of fast exocytosis in
mutant IHCs—indicating a normal vesicular release proba-
bility—led to the previous hypothesis that the defect primarily
reflects a reduction of the readily releasable pool of vesicles
(RRP) due to the loss of the ribbon (Khimich et al., 2005).
However, the exact structural and functional correlates of the
RRP reduction remained unclear. For example, potential differ-
ences between mutant AZs that still have a ribbon (ribbon occu-
pied) and their ribbonless counterparts have not yet been inves-
tigated. Moreover, it is not known to which degree and by which
mechanism Ca2+influx is affected at the level of individual
synapses and how this might contribute to the exocytic deficit.
Several mechanisms may explain the impairment of fast exocy-
tosisinIHCs ofBsnDEx4/5mutants.First, mutantAZsmaycontain
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fewer vesicular docking sites and/or closely colocalized Ca2+
channels. Together, they have been suggested to constitute
the numerous release sites of the IHC AZ at which vesicle fusion
iscontrolled bytheCa2+nanodomain ofoneorfewnearbyactive
Ca2+channels (Brandt et al., 2005; Moser et al., 2006; Goutman
and Glowatzki, 2007). Vesicles docked and primed in these
‘‘slots’’ probably constitute the RRP, of which the released frac-
tion but not the release kinetics depends on the number of slots
recruited by a given stimulus (Brandt et al., 2005; Furukawa and
Matsuura, 1978; Wittig and Parsons, 2008). Therefore, fewer
release sites, because of fewer Ca2+channels (Neef et al.,
2009) and/or fewer docking sites, could explain impaired fast
exocytosis as a deficit of RRP size. Second, even if the number
of release sites was unchanged, the standing RRP would be
diminished if vesicle occupancy at each of these sites was
reduced in BsnDEx4/5IHCs, e.g., because of impaired replenish-
ment or enhanced undocking of vesicles. Third, the coupling
between Ca2+influx and Ca2+sensors of the exocytosis
machinery could be altered, such that not all vesicles can
contribute to fast exocytosis, even after proper docking and
biochemical priming. This point subsumes changes in diffusion,
buffering, or homeostasis of [Ca2+]i, as well as an increased
distancebetweenchannelsandCa2+sensors,positionalpriming
(Neher and Sakaba, 2008), as it was reported at the Drosophila
neuromuscular junction after disruption of the presynaptic scaf-
fold protein Bruchpilot (Kittel et al., 2006). Finally, the intrinsic
Ca2+sensitivity of exocytosis could be altered.
The availability of a number of techniques such as improved
stimulated emission depletion (STED) microscopy, and fast
confocal imaging of Ca2+influx, as well as the generation of
another Bassoon-deficient mouse line (Bsngt) now allowed us
to address these questions. Here, we used in vitro and in vivo
physiology in combination with light and electron microscopy
and computational modeling to study in detail structural and
functionaleffectsofBassoondisruptionatbothribbon-occupied
and ribbonless AZs. Our results indicate that both functional
inactivation of Bassoon and ribbon loss reduce the number of
synaptic Ca2+channels. Membrane tethering of vesicles was
improved but not fully normal at ribbon-occupied mutant AZs,
suggesting a partial function of these ribbons. Mutant IHCs
showed a reduction in the number of release sites while main-
taining an intact coupling of Ca2+influx to exocytosis. Vesicle
replenishment was slightly impaired in in vitro experiments. We
conclude that the multiprotein complex of the synaptic ribbon
and Bassoon organize Ca2+channels and synaptic vesicles at
the AZ, thereby creating a large number of release sites.
RESULTS
The most prominent morphological phenotype of IHCs associ-
ated with the disruption of Bassoon function in mouse mutants
with partial gene deletion (BsnDEx4/5) is the loss of synaptic
ribbons from their AZs (Buran et al., 2010; Khimich et al.,
2005). In IHCs of immunolabeled whole-mounted organs of Corti
from 3-week-old mice, we used confocal microscopy to count
ribbon synapses as juxtaposed spots of presynaptic CtBP2/
RIBEYE (labeling ribbons) and postsynaptic GluR2/3 (labeling
glutamate receptor clusters). Per IHC in BsnDEx4/5, we found on
average 2.5 ribbon-occupied synapses (22% of 1240 synapses,
n=112IHCs) insteadof 11.9ribbon-occupied synapsesinBsnwt
(97% of 1028 synapses, n = 84 IHCs). Consistent with observa-
tions at retinal photoreceptor ribbon synapses (Dick et al., 2003),
we detected expression of the N-terminal Bassoon fragment in
IHCs of BsnDEx4/5mice (Figure S1A, available online) but found
that it was not localized to afferent IHC synapses, arguing
against a residual function at the AZ. This observation and the
absence of an auditory deficit in 8-week-old heterozygous
BsnDEx4/5mice (data not shown) do not support the idea of
a dominant negative effect of the N-terminal Bassoon fragment.
We also observed fewer ribbon-occupied synapses in IHCs of
the newly generated Bassoon-deficient mouse line Bsngt(4.8
versus 9.6 ribbon-occupied synapses per IHC in wild-type),
which, like BsnDEx4/5mice, showed a mild hearing impairment
(threshold increase by 23 dB for click stimuli in four Bsngtmice
compared to three wild-type littermates versus 37 dB increase
in BsnDEx4/5; Pauli-Magnus et al., 2007). A weak Bassoon immu-
nolabeling was observed at a small subset (approximately 10%)
of synapsesin BsngtIHCs (Figure S1B),potentially explaining the
higher number of ribbon-occupied AZs in BsngtIHCs.
Reduction of Membrane-Proximal Vesicles at Hair Cell
Synapses of Bassoon Mutants
We studied effects of Bassoon disruption and ribbon loss on
synaptic ultrastructure in electron micrographs of 80 nm
sections (Figures 1A and 1B). Membrane-proximal vesicles at
apparently ribbonless BsnDEx4/5AZs showed an altered distribu-
tion. When measuring their lateral position relative to the presyn-
aptically projected center of the postsynaptic density, we
observed a broad and seemingly random distribution of those
vesicles at the AZ (Figure 1C, gray bars). In contrast,
membrane-proximal vesicles at AZs of BsnwtIHCs fell into two
categories: ribbon-associated (red open bars) and non-ribbon-
associated (black open bars). The latter population was indistin-
guishable from membrane-proximal vesicles at ribbonless
BsnDEx4/5AZs (p = 0.27, Kolmogorov-Smirnov test). We then
counted the total number of those vesicles in single 80 nm
sections and observed significantly fewer vesicles at apparently
ribbonless (1.5 ± 0.2 vesicles, 53 AZs) and ribbon-occupied
BsnDEx4/5synapses (2.0 ± 0.4 vesicles/AZ section, 10 AZs)
than at ribbon-occupied Bsnwtsynapses (4.2 ± 0.4 vesicles/AZ
section, 26 AZs, p < 0.01 for both comparisons).
Because the absence of a synaptic ribbon cannot unequivo-
cally be concluded from not seeing a ribbon in a single 80 nm
synaptic section, we used electron tomography to address
potential differences between ribbon-occupied and ribbonless
AZs in Bsn mutant mice (Figures 1E–1H). We used Bsngtmice
for these experiments because of their larger fraction of
ribbon-occupied AZs. In electron tomography, we counted
vesicles that were tethered to the plasma membrane by filamen-
touslinkers(seeFigure1Dforexamples;Ferna ´ndez-Busnadiego
et al., 2010). Indeed, we found a trend toward more membrane-
tethered vesicles when a ribbon was present (6.4 ± 0.8, n = 10
versus 3.7 ± 1.1, n = 6 at ribbonless BsngtAZs; p = 0.1), probably
reflecting the addition of a ribbon-associated vesicle population.
As in the analysis of 80 nm sections of BsnDEx4/5AZs, vesicle
numbers at ribbon-occupied BsngtAZs did not reach Bsnwt
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levels (10.6 ± 0.7, n = 5, p < 0.01). We also observed that, unlike
at Bsnwtsynapses, ribbons of Bsngttended to be farther away
from the plasma membrane (Figure 1I). In fact, we found a spec-
trum of ribbon-anchorage phenotypes: from wild-type-like prox-
imity to loosely anchored ribbons (often accompanied by
a second detached ribbon) to complete ribbon absence. It is
tempting to speculate that loosely anchored ribbons may not
fully promote membrane tethering of vesicles. We note that
AB
E1
D
E2
G1H1
J
I F1
G2H2
coated vesicles
F2
membrane-tethered vesicles
presynaptic density
plasma membrane
ribbon-associated vesicles
ribbon
C
3
2
1
0
Vesicles / section
n.s.
***
wt
Bsn
ΔEX4/5
CaVβ2
KO
150
100
50
0
Ribbon distance (nm)
Bsn
wt
Bsn
gt (1
strib.)
Bsn
gt (all rib.)
p=0.06**
0.6
0.4
0.2
0.0
Vesicles / section
600 400 2000
Distance to PSD center (nm)
Bsn
Bsn
Bsn
ΔEx4/5
wt ribbon-assoc.
wt non-ribbon-assoc.
Bsn
wt
ribbon-occupied
r
IHC
PSD
aff
SV
100 nm
r
IHC
PSD
aff
SV
Bsn
wt
ribbon-occupied
100 nm
Bsn
ΔEx4/5ribbon-occupied
100 nm
►
SV
SC
Bsn
wt
Bsn
gt
SV
SC
►
Bsn
gt
►
SV
SC
Bsn
gt
►
SV
SC
Bsn
gt
ribbon-occupied
100nm
Bsn
gt
ribbon-occupied
100nm
Bsn
gtribbonless
100nm
non-tethered vesicles
Figure 1. Synaptic Ultrastructure and Vesicle Distribution in the Presence and Absence of the Synaptic Ribbon
(A and B) Electron micrographs of single thin sections of Bsnwt(A) and BsnDEx4/5ribbon-occupied IHC ribbon synapses (B). The following abbreviations are used:
r, ribbon; SV, synaptic vesicle; PSD, postsynaptic density; aff, afferent bouton.
(C) Distribution of membrane-proximal synaptic vesicles in Bsnwtand BsnDEx4/5IHCs as a function of distance from the PSD center. The histogram was normal-
ized to the number of sections analyzed in the respective genotype (Bsnwt, n = 58 SVs, 18 sections; BsnDEx4/5, n = 74 SVs, 36 sections).
(D)Example slices fromsingle-axis electron tomograms showing membrane-tetheredsynapticvesicles.Tethers aremarked byarrowheadsSCdenotes synaptic
cleft. The scale bars represent 40 nm.
(E–H) Single slices from tomograms for Bsnwt(E1), Bsngtribbon-occupied (F1 and G1), and Bsngtribbonless synapses (H1). (E2–H2, upper) Tomogram-based
model of Bsnwt(E2), Bsngtribbon-occupied (F2 and G2), and Bsngtribbonless synapses (H2). Vesicles distant from the ribbon and the plasma membrane are
not shown. (Lower) Same models as in upper but only showing membrane-proximal SVs used for analysis (see Results).
(I)Barplotshowing mean minimal distance betweenribbonand plasma membrane asmeasured inelectron tomograms ofBsnwtAZs(black;n= 5ribbons/5 AZs),
of just the proximal ribbons at BsngtAZs (red; n = 10 ribbons/10 AZs), and of all ribbons at BsngtAZs (light red; n = 16 ribbons/10 AZs). The error bars represent
standard error of the mean (SEM).
(J) Bar plot, showing average number of membrane-proximal SVs per thin section for wild-type (black; n = 46 AZs, pooled data from Bsn and CaVb2wild-type
littermates) and mutant synapses. BsnDEx4/5(red; n = 67 AZs), but not CaVb2knockout synapses (blue; n = 32 AZs), had approximately one-half the numbers of
membrane-proximal SVs. The error bars represent SEM.
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Figure 2. Decreased Immunofluorescence and Altered Shape of CaV1.3 Clusters
(A) Projections of confocal sections of IHCs of apical cochlear coils immunolabeled for synaptic ribbons (CtBP2, red) and Ca2+channels (CaV1.3, green) as used
for analysis in (B and C). (Left) Four BsnwtIHCs (n is an abbreviation for nuclei). (Middle) Enlargement of part of the synaptic layer (white box, left) showing coloc-
alization of CtBP2 and CaV1.3. (Right) In the partial gene deletion mutant (BsnDEx4/5), Ca2+channels still cluster but few ribbons remain (P28). Arrowheads point to
ribbonless CaV1.3 clusters in wild-type and mutant. Arrow points to a ribbon-occupied CaV1.3 cluster. Asterisk labels a floating ribbon.
(B) CaV1.3 immunofluorescence intensity (mean ± SEM, a.u.) was less at BsnDEx4/5synapses when analyzing only CaV1.3 clusters that colocalized with GluR2
(gray; BsnDEx4/5versus Bsnwt, p < 0.0005) or when counting the ten brightest clusters per hair cell (black; BsnDEx4/5versus Bsnwt, p < 1e?20). In both Bsnwt
and BsnDEx4/5, the presence of a ribbon (CtBP2 colocalized: ribbon occupied, red) was associated with greater CaV1.3 intensity when compared to ribbonless
synapses (blue). Bsnwtribbon occupied versus Bsnwtribbonless, p < 0.05; BsnDEx4/5ribbon occupied versus BsnDEx4/5ribbonless, p < 0.005).
(C) CaV1.3 cluster intensity histogram for Bsnwt(solid line) and BsnDEx4/5(dotted line). Each distribution is decomposed into ribbon-occupied (red) and ribbonless
(blue) clusters.
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even in the 250 nm tissue sections that were used for tomog-
raphy, the reported vesicle numbers represent underestimates
of the full complement of membrane-proximal vesicles because
synapses were not completely included along one dimension.
However, this error equally affected each synapse type, and
tomograms fully contained the synapse in the other two dimen-
sions. Notably, we found that the electron-dense material lining
the presynaptic plasma membrane (presynaptic density) was
longer and thicker at ribbon-occupied BsnwtAZs than the
spot-likepresynapticdensitiesatBsngtAZs(regardlessofribbon
presence; Figures 1E2–1H2 and Table S1), which sometimes
harbored more than one density (Figures 1F2 and1 H2).
Finally, we also studied AZs in IHCs of mouse mutants that
contain fewer Ca2+channels because of a lack of the b2subunit
(CaVb2; Neef et al., 2009). CaVb2-deficient IHCs display a 70%
reduction of both Ca2+influx and RRP exocytosis despite the
presence of synaptic ribbons. Number (Figure 1J, data from
wild-type littermates of BsnDEx4/5and CaVb2 mutants were
pooled) and distribution (data not shown) of membrane-proximal
vesicles were unaltered in 80 nm sections, suggesting that
proteins of the macromolecular ribbon complex, but not Ca2+
channels, are required for the formation of vesicle docking sites.
Fewer Ca2+Channels and Altered Shape of Ca2+
Channel Clusters
Voltage-gated Ca2+influx is decreased in IHCs of Bsn mutants
(BsnDEx4/5; Khimich et al., 2005). Here, we explored changes of
synaptic Ca2+
signaling by morphological and functional
imaging. First, we studied synaptic Ca2+channel clusters by
confocal and STED microscopy following immunolabeling of
CaV1.3 Ca2+channels. Images of BsnDEx4/5and Bsnwtorgans
of Corti that had been processed for immunohistochemistry in
parallel and following the same protocol were acquired with
identical microscope settings and analyzed for intensity and
shape of CaV1.3 immunofluorescent spots (Figure 2). We esti-
mated the short and long axes of the elliptic fluorescent objects
by fitting 2D Gaussian functions to the background-subtracted
images (see Supplemental Experimental Procedures). The fluo-
rescence integral within this region served as a proxy of the
abundance of synaptic Ca2+channels. In Bsnwtorgans of Corti,
the synaptic location of CaV1.3 clusters (Figure 2A) was readily
confirmed by the colocalization with synaptic ribbons (Brandt
et al., 2005; Meyer et al., 2009) and Bassoon (Figure S1B). In
addition, some lower intensity spot-like immunofluorescence
was present in IHCs (Figure 2). In BsnDEx4/5and BsnwtIHCs,
the synaptic localization of Ca2+channel clusters was identified
by costaining for postsynaptic glutamate receptors (GluR2; Fig-
ure S1C). This confirmed that Ca2+channels remained clustered
at synapses despite both the disruption of Bassoon and, in most
cases, absence of the ribbon. In comparison to Bsnwt, the immu-
nofluorescence of CaV1.3 clusters colocalized with GluR2 was
reduced at BsnDEx4/5synapses (Figure 2B and Figure S1D).
In experiments colabeling for CaV1.3 and the synaptic ribbon
marker RIBEYE/CtBP2, we were able to separate ribbon-occu-
pied AZs from ribbonless AZs in BsnDEx4/5and Bsnwtmice.
Because of the absence of an additional synaptic marker at rib-
bonless BsnDEx4/5AZs, and to exclude nonsynaptic CaV1.3
immunofluorescent spots from analysis, we considered only
the ten brightest spots in each cell for both genotypes. This
approach was justified by knowledge of cochlear location (ten
synapses per cell in apical turn; Meyer et al., 2009) and the
observation that 92.2% and 89.4% of the ten brightest Ca2+
channel clusters were juxtaposed to GluR2 immunofluorescent
spots in BsnDEx4/5and BsnwtIHCs, respectively. In confocal
images, CaV1.3 immunofluorescence was reduced by 42% at
BsnDEx4/5AZs (Figure 2B; p < 1e?20). As quantified in Figures
2B and 2C and Table 1, the CaV1.3 immunofluorescence
decreased in the order: ribbon-occupied Bsnwt> ribbonless
Bsnwt> ribbon-occupied BsnDEx4/5> ribbonless BsnDEx4/5.
CaV1.3 channel clusters of ribbon-occupied BsnDEx4/5AZs
were also more similar to BsnwtAZs in shape than those of rib-
bonless BsnDEx4/5AZs. The altered shape of the latter was
evident in a smaller long-to-short axis ratio (standard STED;
Figures 2D and 2E and Table 1).
To resolve a potential substructure within CaV1.3 clusters, we
used a custom-built STED microscope (STED*; lateral point
spread function [PSF] less than 100 nm at a tissue depth of
15 to 25 mm; yellow range in Figure 2E). Ca2+channel clusters
of BsnwtAZs typically displayed one to three stripes of CaV1.3
immunofluorescence (Figure 2F). Parallel confocal observation
of the associated CtBP2/RIBEYE immunofluorescence sug-
gested that these synapses featured one ribbon regardless of
the number of stripes, although two closely-spaced ribbons
may fall within the confocal PSF and thus may not be resolved
as individual ribbons. In contrast, BsnDEx4/5AZs showed
CaV1.3immunofluorescencespotsratherthanstripes(Figure2F;
full width at half maximum of long and short axes: 120 ± 7.9 nm
and 95 ± 5.5 nm, n = 13) with ribbon-occupied AZs typically
harboring more spots than ribbonless AZs. CaV1.3 immunofluo-
rescent stripes and spots were reminiscent of the patterns of
presynaptic density observed in electron tomography (Figures
1E2–1H2). In summary, the abundance of synaptic Ca2+chan-
nels and the cluster shape are altered upon Bassoon disruption,
which might reflect the loss of a direct Bassoon action on Ca2+
channelclusteringorofthe
anchorage. To test for a potential role of Bassoon in the direct
synaptic anchoring of Ca2+channels, we determined whether
Bassoon and the CaV1.3 channel interacted in a heterologous
expressionsystem.WedidnotfindevidencethatBassooncoim-
munoprecipitated or colocalized with Cav1.3 in transfected
HEK293T cells (Figure S4). Therefore, the role of Bassoon in
Bassoon-mediatedribbon
(D) Single Z sections of CtBP2 (confocal) and CaV1.3 (standard STED) for size analysis in (E) of CaV1.3 clusters at Bsnwt(left) and BsnDEx4/5AZs (right).
(E) Compared to Bsnwt(filled circles: individuals, gray; mean, black) and ribbon-occupied BsnDEx4/5synapses (red open circles; mean, dark red), the ribbonless
CaV1.3 clusters (blue open circles; mean, dark blue) fell closer to unity (dashed line). Apparent sizes of 40 nm beads mounted above and below the organ of Corti
illustrate the PSF range for the two STED microscopes (black, Leica STED; yellow, custom STED*). The error bars represent SEM.
(F) STED* microscopy revealed ribbon-occupied BsnwtCaV1.3 clusters as one or more elongated stripes, not observed at BsnDEx4/5synapses.
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recruiting Ca2+channels to the AZ may not involve a direct asso-
ciation of the two proteins.
Reduced Synaptic Ca2+Influx Results from Fewer
Channels and Lower Open Probability
To study synaptic Ca2+influx in BsnDEx4/5IHCs, we performed
whole-cell patch-clamp recordings of Ca2+current (ICa) and
confocal imaging of presynaptic Ca2+microdomains (Frank
et al., 2009). We found a reduction of peak whole-cell ICaampli-
tudeinBsnDEx4/5IHCsof3-week-oldmice(Figure3,Table1,Fig-
ure S2, and Table S1). It ranged between 62% (ruptured-patch,
5mM[Ca2+]e;Figure3AandTable1)and69%(perforated-patch,
10 mM [Ca2+]e; Figure S2 and Table S1) of Bsnwtamplitude. The
difference from Bsnwtwas alleviated in the presence of the dihy-
dropyridine agonist BayK8644 (77%, ruptured-patch, 10 mM
[Ca2+]e; Figure 3B and Table 1), suggesting that Ca2+channel
open probability is reduced in BsnDEx4/5IHCs in the absence of
BayK8644. Moreover, we found that Ca2+current activation
was slowed in BsnDEx4/5IHCs (Figure 3C and Figure S2), while it
was indistinguishable from wild-type IHCs in the presence of
BayK8644 (Figure 3D). Finally, the Ca2+currents inactivated
slightly more in BsnDEx4/5IHCs (Figure 3E and Table S1).
To test whether the observed reduction in ICawas caused by
changes in channel number (NCa), unitary current (iCa), or open
Table 1. Summary of IHC Physiology in Bsn Wild-Type and Mutant Mice
Bsnwt
Parameter
BsnDEx4/5
p value
CaV1.3 Immunofluorescence
Intensity, confocal (a.u.) Ribbonless:
3.1e5 ± 3e4
(n = 16)
Ribbon-occupied:
4.0e5 ± 1e4
(n = 219)
Ribbonless:
2.1e5 ± 1e4
(n = 196)
Ribbon-occupied:
2.9e5 ± 3e4
(n = 46)
Bsnwt, ribbonless versus
ribbon-occupied, p < 0.05;
BsnDEx4/5, ribbonless versus
ribbon-occupied, p < 0.01;
wild-type versus mutant,
p < 0.001
Bsnwtversus BsnDEx4/5,
ribbonless, p < 0.001; Bsnwt
versus BsnDEx4/5, ribbon-
occupied, n.s.; BsnDEx4/5,
ribbonless versus ribbon-
occupied, p < 0.05
FWHM, STED,
long:short (nm)
Ribbon-occupied:
345.9 ± 12:
230.0 ± 6 (n = 68)
Ribbonless:
300.3 ± 15:
251.4 ± 14
(n = 45)
Ribbon-occupied:
355.6 ± 22:
251.8 ± 15 (n = 16)
Whole-Cell Ca2+Current
Peak ICa(pA): 5Ca2+/?BayK
Peak ICa(pA): 10Ca2+/+BayK
NCa: 10Ca2+/+BayK
Synaptic Ca2+Microdomains
DFavg(a.u.): 5 Ca2+/?BayK
?179.5 ± 9.6 (N = 31)
?417.5 ± 29.0 (N = 29)
1574 ± 92 (N = 27)
?111.1 ± 6.2 (N = 38)
?321.0 ± 33.9 (N = 19)
1227 ± 111 (N = 22)
p < 0.001 (W)
p < 0.05 (W)
p < 0.01 (W)
85.5 ± 9.1
(n = 74/N = 30)
30.5 ± 2.0
(n = 112/N = 45)
p < 0.001 (W)
DFavg(a.u.): 5 Ca2+/+BayK 89.5 ± 9.0
(n = 53/N = 21)
46.9 ± 5.2
(n = 52/N = 20)
p < 0.001 (W)
DFavg(a.u.): ±BayK p = 0.14 (W)p < 0.01 (W)
Exocytosis
DCm,20 ms(fF) ? pp
QCa,20ms(pC) ? pp
DCm,20 ms(fF) ? rp
QCa,20ms(pC) ? rp
DCm,100 ms(fF) ? pp
QCa,100ms(pC) ? pp
DCm,100 ms(fF) ? rp
DCm,100 ms(fF) ? rp
sustained DCm(DCm,100?
DCm,20; fF)
n denotes number of synapses (CaV1.3 immunofluorescence and synaptic Ca2+microdomains) and N number of IHCs (whole-cell Ca2+current, Ca2+
imaging,and capacitance measurements). n.s.denotesnot significant. Forimmunofluorescence,Bsn mutant data were separated into ribbonless and
ribbon-occupied synapses. Statistical comparisons were made with an independent two-sample t test (T) or a Mann-Whitney-Wilcoxon (W) test
(Experimental Procedures). a.u., arbitrary units; FWHM, full width at half-maximum; ICa, whole-cell Ca2+current; NCa, number of Ca2+channels; DFavg,
averageCa2+microdomainamplitude;DCm,exocyticmembranecapacitancechanges;pp,perforated-patchconfiguration;QCa,Ca2+currentintegral;
rp, ruptured-patch configuration. Sustained DCmwas calculated cell-wise, by subtracting the average DCmresponse to 20 ms from the average DCm
response to 100 ms. Data are presented as mean ± SEM.
13.2 ± 1.1 (N = 38)7.9 ± 0.9 (N = 37)p < 0.001 (W)
4.4 ± 0.3 (N = 38)3.0 ± 0.2 (N = 37)p < 0.001 (W)
8.1 ± 1.0 (N = 17) 4.7 ± 0.8 (N = 16)p < 0.01 (W)
3.3 ± 0.2 (N = 17)2.6 ± 0.1 (N = 16)p < 0.05 (T)
39.6 ± 5.2 (N = 41)21.9 ± 4.3 (N = 40)p < 0.001 (W)
20.4 ± 1.4 (N = 41)13.2 ± 0.9 (N = 40)p < 0.001 (W)
35.1 ± 5.3 (N = 13)19.5 ± 5.8 (N = 11)p < 0.01 (W)
15.5 ± 1.0 (N = 13)11.2 ± 0.9 (N = 11)p < 0.01 (T)
26.8 ± 4.6 (N = 38)15.1 ± 4.4 (N = 35)p < 0.001 (W)
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Page 7

probability (popen), we performed a nonstationary fluctuation
analysis on Ca2+tail-currents ([BayK8644]e = 5 mM; Brandt
et al., 2005). In line with the observed reduction in ICaamplitude,
bothvarianceandmeanwerereducedinBsnDEx4/5IHCs(Figures
3F and 3G). The analysis indicated a ?20% decrease in the
number of functional Ca2+channels but statistically indistin-
guishable single-channel currents and maximal open probabili-
ties in the presence of BayK8644 (Table 1 and Table S1). We
note that due to uncertainties associated with the channel prop-
erty estimates from fluctuation analysis (Tables 1 and Table S1),
which also deviate from those obtained from single-channel
recordings in immature IHCs (Zampini et al., 2010), emphasis
isoncomparisonbetweenthegenotypesratherthanonabsolute
values (see also Supplemental Experimental Procedures).
Synaptic Ca2+microdomains, primarily reflecting Ca2+influx
at the AZ (Frank et al., 2009), were visualized with the low-
affinity Ca2+indicator Fluo-5N (400 mM, Kd= 95 mM) in conjunc-
tion with the slow Ca2+chelator EGTA (2 mM). The Ca2+micro-
domain amplitude (DF) measured under these conditions
probably reflects a linear summation of the Ca2+influx contrib-
uted by the individual synaptic Ca2+channels (Frank et al.,
2009). Consistent with the finding of CaV1.3 channel clusters
in immunohistochemistry, we readily observed Ca2+microdo-
mains also in BsnDEx4/5IHCs (Figure 4A). However, their
average amplitude (DFavg; Table 1), measured at ?7 mV in
spot-detection experiments at the center of the Ca2+microdo-
mains, was reduced to 36% of control (Figures 4B and 4C),
exceeding the reduction of whole-cell ICa (to 60%–70%;
A
C
F
E1E2
Normalized ICa
G
D
τactivation (ms)
B
− BayK
− BayK
+ BayK
+ BayK
-150
-100
-50
ICa (pA)
-80-60-40-202040
Vm (mV)
Bsn
Bsn
wt
ΔEx4/5
-400
-200
ICa (pA)
-80 -60-40-202040
Vm (mV)
Bsn
Bsn
wt
ΔEx4/5
300
200
100
0
τactivation (μs)
-40-20020
Vm (mV)
Bsn
Bsn
wt
ΔEx4/5
**
**
***
***
***
1.0
0.5
0.0
-40-200 20
Vm (mV)
Bsn
Bsn
wt
ΔEx4/5
-1.0
-0.5
0.0
<ICa> (nA)
200
0
<VAR> (pA²)
1 ms
Bsn
Bsn
wt
ΔEx4/5
150
100
50
0
<Var> (pA²)
800600 4002000
<ICa> (pA)
Bsn
Bsn
wt
ΔEx4/5
-1.0
-0.5
0.0
Normalized ICa
50 ms
Bsn
wt
Bsn
ΔEx4/5
-1.0
-0.5
0.0
100 ms
Bsn
wt
Bsn
ΔEx4/5
Figure 3. Biophysical Properties of Voltage-
Dependent Whole-Cell Ca2+Current (ICa)
(A) Average steady-state ICa-V for Bsnwtand
BsnDEx4/5IHCs in 5 mM [Ca2+]e(n [Bsnwt] = 31
IHCs, n [BsnDEx4/5] = 38 IHCs). Note the reduction
of max. ICato ?60% of wild-type level in BsnDEx4/5
IHCs (Table 1).
(B) As in (A) but in 10 mM [Ca2+]eand presence
of 5 mM BayK8644 (n [Bsnwt] = 29 IHCs,
n [BsnDEx4/5] = 19 IHCs). Note the smaller differ-
ence in max. ICa between the two genotypes
(BsnDEx4/5: ?80% of Bsnwtlevel; Table 1).
(C) Averagetime-constantofICaactivation in5mM
[Ca2+]eas a function of membrane voltage (Vm),
derived from single exponential fits to the initial
3.5 ms of ICa (see Supplemental Experimental
Procedures; n [Bsnwt]: % 30 IHCs, n [BsnDEx4/5]:
% 35 IHCs). Asterisks indicate Vmat which differ-
ences between genotypes were statistically signif-
icant (a = 0.05; Bonferroni correction). Average
series resistance (RS) was 6.0 ± 2.3 MU for Bsnwt
IHCs, and 6.1 ± 2.3 MU for BsnDEx4/5IHCs
(mean ± SD), respectively.
(D) Same as (C) but in 10 mM [Ca2+]eand 5 mM
extracellular BayK8644 (n [Bsnwt]: % 29 IHCs,
n [BsnDEx4/5]: % 19 IHCs). Average RSwas 4.7 ±
3.1 MU for BsnwtIHCs, and 5.3 ± 3.3 MU for
BsnDEx4/5IHCs (mean ± SD), respectively.
The error bars in (A–D) represent SEM.
(E1and E2) Average paired-pulse ICatraces (depo-
larization to Vmof maximum ICa; [Ca2+]e= 10 mM)
illustrate stronger inactivation in BsnDEx4/5IHCs,
being more evident for longer (100 ms; E2) than
for shorter depolarizations (20 ms; E1).
(F) Example mean Ca2+tail-currents (lower) used
for nonstationary fluctuation analysis (Tables 1
and Table S1), elicited by repolarizing IHCs
from +57 mV to ?68 mV, and corresponding
mean trial-to-trial variance (upper). [Ca2+]e =
10 mM, [BayK8644]e= 5mM.
(G) Grand average (lines with filled circles) of vari-
ance versus mean relationships (n [Bsnwt] = 27
IHCs, n [BsnDEx4/5] = 22 IHCs). Filled areas depict
SD of grand average of variance. Broken lines
represent grand average of parabolic fits (Supple-
mental Experimental Procedures).
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Figure 3) and CaV1.3 immunofluorescence (to 58%; Figure 2).
Augmenting influx through CaV1.3 Ca2+
BayK8644) alleviated the amplitude reduction of synaptic Ca2+
influxinBsnDEx4/5IHCs(to52%ofcontrol;Figure4DandTable1)
and increased amplitude variability among the BsnDEx4/5, but not
the Bsnwtsynapses (Figures 4E and 4F). Kinetics (Figures 4C
and 4D, Figure S3, and Table S1), voltage dependence (Fig-
ure 4G, Figure S3, and Table S1), and spatial extent (Figure 4H
and Table S1) of the Ca2+microdomains in BsnDEx4/5IHCs were
similar to control. There was, however, a tendency toward faster
kineticsandmorenegativeactivationofBsnDEx4/5Ca2+microdo-
mains(TableS1).WhiletheformermayreflectdifferencesinCa2+
channels (5 mM
A
B
1000
500
0
ΔF (a.u.)
2 μm
20 ms
− BayK
+ BayK
20 ms
C
D
E
Cumulative frequency
F
Cumulative frequency
H
G
I1
20 ms
20 ms
400
200
0
ΔF (a.u.)
2 μm
I2
20 ms20 ms
1.0
0.5
0.0
norm. ΔF/F0
-60-40-200
Vm(mV)
Bsn
wt
Bsn
ΔEx4/5
200
100
0
ΔF (a.u.)
Bsn
wt
Bsn
ΔEx4/5
200
100
0
ΔF (a.u.)
Bsn
wt
Bsn
ΔEx4/5
1.0
0.5
0.0
3002001000
ΔFavg.(a.u.)
– BayK (CV=0.91)
+ BayK (CV=0.73)
Bsn
wt
1.0
0.5
0.0
150 100 500
ΔFavg.(a.u.)
– BayK (CV=0.70)
+ BayK (CV=0.79)
Bsn
ΔEx4/5
Bsn
ΔEx4/5
Bsn
wt
Bsn
ΔEx4/5
Bsn
wt
100
50
0
ΔF (a.u.)
-200
0
ICa(pA)
Bsn
wt
Bsn
ΔEx4/5
100
50
0
ΔF (a.u.)
Bsn
ribbon-occupied
(n=92)
wt
wt
Bsn
ribbonless
(n=12)
100
50
0
ΔF (a.u.)
Bsn
ribbon-occupied
(n=46)
ΔEx4/5
ΔEx4/5
Bsn
ribbonless
(n=50)
Figure 4. Reduced Presynaptic Ca2+Influx
(A) Exemplary localized Ca2+influx sites in optical
sections through the basal part of IHCs (Experi-
mental Procedures). Resting fluorescence (F0)
was subtracted (DF images). Ca2+microdomains
at AZs are present in BsnDEx4/5IHCs, albeit of
smaller amplitude.
(B) Exemplary spot-detection responses during
depolarization to ?7 mV (bar, lower; all reported
responses were from the Ca2+
center) and simultaneously acquired whole-cell
ICa(upper). Note the out-of-proportion reduction
of synaptic Ca2+influx.
(C) Grand average of spot-detection responses
from Ca2+microdomains (n [Bsnwt] = 74 AZs/30
IHCs, n [BsnDEx4/5] = 112 AZs/45 IHCs); shaded
areas indicate SD.
(D) Same as (C) but in presence of 5 mM BayK8644
(n [Bsnwt] = 53 AZs/21 IHCs, n [BsnDEx4/5] = 52 AZs/
20 IHCs). Note the peak at the end of the stimula-
tion, corresponding to tail current-mediated Ca2+
influx (prolonged due to BayK8644).
(E and F) Cumulative frequency distributions of
Bsnwt(E) and BsnDEx4/5(F) Ca2+microdomain
amplitudes (averaged over the second half of the
stimulus)ineitherabsence(black/gray)orpresence
(red/light red) of 5 mM BayK8644. CV denotes coef-
ficient of variation (SD/mean).
(G) Normalized steady-state fluorescence-voltage
relationships (n [Bsnwt] = 19 AZs, n [BsnDEx4/5] =
27 AZs). Relative fluorescence changes were aver-
aged over the last 14.6 ms of the 20 ms stimulus
and normalized to the peak response of the given
Ca2+microdomain. Shaded areas depict SEM.
(H) Representative line scans across the Ca2+
microdomain center (5 mM [Ca2+]e). Bar indicates
period of depolarization to ?7 mV.
(I) Grand average of Bsnwt(I1) and BsnDEx4/5(I2)
spot-detection responses, sorted according to
the presence/absence of a colocalized ribbon
(Experimental Procedures). n (Bsnwt) = 104 AZs/
32 IHCs, n (BsnDEx4/5) = 96 AZs/37 IHCs.
microdomain
buffering
may indicate an altered gating of synaptic
Ca2+channels in the absence of Bassoon
and/or the ribbon.
In a second set of experiments, we
studied Ca2+signaling at ribbonless and
AZsin separation
RIBEYE-binding peptide to identify ribbon-occupied AZs (Frank
et al., 2009; Zenisek et al., 2004) in both BsnDEx4/5and Bsnwt
IHCs. While Ca2+microdomains at ribbon-occupied AZs had
larger amplitudes than ribbonless synapses in BsnwtIHCs,
there was no significant difference between ribbonless and
ribbon-occupied AZs in BsnDEx4/5IHCs (Figure 4I). The latter
finding was unexpected given their difference in CaV1.3 immu-
nofluorescence but could reflect limited sensitivity of functional
Ca2+imaging, precluding detection of very dim Ca2+signals at
BsnDEx4/5ribbonless synapses. In summary, the reduced ampli-
tude of Ca2+microdomains and its partial alleviation upon the
and/ordiffusion,thelatter
ribbon-occupiedwithafluorescent
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BayK8644-mediated increase in open probability led us to
conclude that BsnDEx4/5synapses contain fewer Ca2+channels
with a lower open probability. The reduction of synaptic Ca2+
influx beyond the decrease observed in whole-cell ICaindicates
a higher proportion of extrasynaptic channels in BsnDEx4/5
IHCs.
Reduced RRP and Sustained Exocytosis but Intact Ca2+
Influx-Exocytosis coupling
How does the reduction of Ca2+channels and membrane-prox-
imal vesicles—as well as a potential mislocalization of these two
elements—affect hair cell exocytosis? We addressed this ques-
tion in BsnDEx4/5IHCs by measuring exocytic membrane capac-
itance changes (DCm) in response to short (20 ms, DCm, 20 ms)
andlonger(100ms,DCm, 100 ms)depolarizations tothemaximum
Ca2+current potential in native buffering conditions (perforated-
patch configuration; Figure 5). Based on previous work (Gout-
man and Glowatzki, 2007; Li et al., 2009; Meyer et al., 2009;
Neef et al., 2009; Rutherford and Roberts, 2006; Schnee et al.,
2005), we interpret DCm, 20 msas fast (synchronous) exocytosis,
representing release of a standing RRP, and the difference
between DCm, 100 msand DCm, 20 msas sustained exocytosis, re-
flecting vesicle supply to the RRP and subsequent fusion. In this
set of experiments, DCm, 20 mswas reduced to 60% (Figure 5A
and Table 1) and sustained exocytosis to 56% (Figure 5B and
Table 1). These results are consistent with a model in which
RRP size and sustained exocytosis rate are related to the
number of physical docking and release sites at the AZ (reduc-
tion of membrane-proximal vesicles: ?50%, Figure 1J). To test
whether the intrinsic Ca2+dependence of exocytosis differed
between genotypes, we used flash photolysis of caged Ca2+,
but found comparable time constants of the fast component of
the DCm, flashfor elevations of [Ca2+]ito 25–37 mM (BsnDEx4/5:
2.4 ± 0.4 ms, mean postflash [Ca2+]i: 29.0 ± 1.9 mM, n = 6 versus
Bsnwt:2.6±1.1ms,meanpostflash[Ca2+]i:31.5±2.5,n=4;Fig-
ure 5C and Table S1) suggesting an unaltered biochemical Ca2+
sensitivity of exocytosis. Notably, despite the lack of ribbons
from most synapses in BsnDEx4/5IHCs the amplitude of their
flash-evoked Cm rise was statistically indistinguishable from
Bsnwt.
A
ICa (pA)
BC
DE1E2F
ΔCm,PP / ΔCm,EGTA
-500
0
50
0
ΔCm (fF)
100 ms
Bsn
Bsn
wt
ΔEx4/5
-200
0
ICa (pA)
50
0
ΔCm (fF)
100 ms
Bsn
Bsn
wt
ΔEx4/5
Bsn
wt
Bsn
ΔEx4/5
1.0
0.5
0.0
Cm,flash (pF)
Δ
0.20.0
Time (s)
2
1
0
20 ms 100 ms
Bsn
Bsn
wt
ΔEx4/5
Bsn
wt
10
5
0
ΔCm,20ms (fF)
420
QCa (pC)
-32mV
Bsn
ΔEx4/5
-27mV
-22mV
-12mV
+18mV
3210
synaptic QCa (pC)
-32mV
-27mV
-22mV
-12mV
+18mV
1.0
0.5
0.0
G/Gmax
-500
Vm (mV)

Bsn
wt

Bsn
ΔEx4/5
-32mV
-27mV
+18mV
-22mV
-12mV
Figure 5. Reduced Exocytosis but Normal Ca2+Influx-Exocytosis Coupling
(A) Representative membrane capacitance changes (DCm) and Ca2+currents (ICa) in response to 20 ms depolarizations to peak-ICaVm.
(B) Same as (A), but in response to 100 ms depolarizations. See Table 1 for pooled data (A and B).
(C) Average DCmresponses recorded during flash photolysis of caged Ca2+. Dataset comprises IHCs (n [Bsnwt] = 4, n [BsnDEx4/5] = 6) with comparable postflash
[Ca2+]i, (range: 25–37 mM; Table S1); 0 ms indicates time of UV flash delivery.
(D) Normalized conductance (G)-voltage relationships for both genotypes.
(E1) Summary of exocytic DCmresponses to 20 ms depolarizations to the five test potentials depicted in (D) plotted versus the corresponding mean Ca2+current
integrals (QCa).Pulses were applied to21 BsnwtIHCs and 23 BsnDEx4/5IHCsinrandom order at intervals of > 30s. Note thelarger responses at +18 mVcompared
to ?27 mV, despite similar QCa, coinciding with a larger popenof Ca2+channels (see D). (E2) To compare the relation between popenand the efficiency of synaptic
vesicle release in the two genotypes, we applied 3 transformations to the plot shown in (E1): (1) assuming a certain extrasynaptic NCa(Brandt et al., 2005), we
estimated the fraction of synaptic Ca2+channels (out of total NCa; Table 1), and multiplied QCaby the respective ratio (<1) to estimate ‘‘synaptic QCa.’’ (2) We then
doubled the mutant DCmdata to account for the halving of membrane-proximal synaptic vesicles seen at mutant AZs (Figure 1). (3) Last, we accounted for the
apparently reduced number of synaptic Ca2+channels at mutant AZs by multiplying mutant QCaby 1/0.52 (assuming that the Ca2+microdomain amplitude in the
presence of BayK8644 presents the most reliable reflection of synaptic NCa; Table 1).
(F) Ratio of exocytic responses (20 ms and 100 ms depolarizations to peak-ICaVm) between perforated-patch (endogenous Ca2+buffers) and ruptured-patch
([EGTA]i= 5 mM) configurations for Bsnwtand BsnDEx4/5IHCs (Table 1).
The error bars in (C, E, and F) represent SEM.
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The observation that the reduction of Ca2+-influx-triggered
exocytosis did not exceed the reduction in the number of
membrane-proximal and -tethered vesicles (Figure 1) suggests
that the remaining docking sites are equipped with nearby Ca2+
channels (reduction of synaptic Ca2+channels: ?50%, Table 1;
estimated from BsnDEx4/5versus Bsnwtsynaptic Ca2+microdo-
main amplitude in the presence of BayK8644). Yet, a looser
coupling between Ca2+channels and vesicle docking sites
than implied for the Ca2+nanodomain regime suggested for
wild-type IHC AZs could not be excluded (Brandt et al., 2005;
Goutman and Glowatzki, 2007; Moser et al., 2006). Therefore,
we studied the sensitivity of exocytosis to the slow Ca2+
chelator EGTA (Figure 5F). Consistent with the preservation of
nanodomain-controlled vesicle fusion in BsnDEx4/5IHCs, their
DCm, 20 msin the presence of 5 mM [EGTA]iwas reduced to
58%ofcontrollevels(Table1)—closelyresemblingthereduction
in the presence of endogenous Ca2+buffers (see above). Addi-
tionally, we probed RRP exocytosis as a function of Ca2+influx
at different membrane potentials (Figures 5D and 5E). Changing
the membrane potential manipulates open probability and
single-channel current in opposite directions. Thus, exocytosis
can be tested for the same absolute Ca2+influx through either
few open channels with high single-channel current (mild depo-
larizations) or more open channels with low single-channel
current (strong depolarizations). If exocytosis of a given vesicle
was under control of a population of several Ca2+channels
(Ca2+microdomain control), exocytosis should be identical for
the same Ca2+current independent of the membrane potential.
In case of a Ca2+nanodomain control, more exocytosis is
expected for more open Ca2+channels, i.e., at more depolarized
potentials (hysteresis; Zucker and Fogelson, 1986). This was
indeed observed in BsnwtIHCs (Figure 5E1 and Figure S5), as
described before (Brandt et al., 2005), but also in BsnDEx4/5
IHCs (Figure 5E1 and Figure S5), further arguing that Ca2+nano-
domain control of exocytosis is maintained at mutant AZs. As
a further consistency check, we scaled the exocytosis-Ca2+
current integral relationship of BsnDEx4/5IHCs by experimentally
derived factors to normalize the data to the lower number of
membrane-proximal vesicles and synaptic Ca2+channels. This
resultedlargelyinanoverlapwiththewild-typedata(Figure5E2).
In summary, the data indicate that the coupling of Ca2+channels
to release sites remains intact despite Bassoon disruption but
that the rates of initial and sustained exocytosis are reduced to
a similar extent as the number of membrane-proximal vesicles.
In Vitro and In Vivo Analysis of Synaptic Vesicle
Replenishment
Traditionally, the synaptic ribbon has been assigned a conveyor
belt and/or attractor function (Holt et al., 2004; Sterling and
Matthews, 2005), according to which it is responsible for rapid
supply of vesicles to the RRP and enables high rates of tonic
neurotransmitter release (Gomis et al., 1999; Johnson et al.,
2008; Moser and Beutner, 2000; Rutherford and Roberts,
2006; Schnee et al., 2005; Spassova et al., 2004). Hence, we
tested whether the rate of RRP refilling was reduced in the
absence of the ribbon and functional Bassoon protein. Here,
we explored vesicle replenishment in vitro by measuring relative
DCm in paired-pulse protocols, with the stimuli (20 ms or
100 ms long depolarizations) being separated by various
time intervals (98, 198, and 398 ms; Figure 6). The ratio of Ca2+
current integrals was close to one in both genotypes (marginally
smaller in BsnDEx4/5IHCs; Figures 6C and 6D and Table S1) indi-
cating that the Ca2+signals that drive exocytosis were mostly
comparable between both pulses. For 20 ms stimuli at short
inter-pulse-intervals (IPI: 98 ms) we observed stronger depres-
sion of the exocytic response in BsnDEx4/5IHCs, indicating
a slower recovery of the RRP at BsnDEx4/5synapses (p < 0.01).
For longer recovery times (IPI: 198, 398 ms), the difference
did not reach statistical significance. While both Bsnwtand
BsnDEx4/5IHCs showed depression for short stimuli, Bsnwt
IHCsexhibitedatendencytowardfacilitationforlongdepolariza-
tions (100 ms). In contrast, BsnDEx4/5IHCs also showed depres-
sion when challenged with long stimuli (p < 0.01 for 98, 198, and
398 ms IPI).
**
**
**
**
*
**
**
*
**
20 ms100 ms
A
C
Q2/Q1
D
B
ICa(pA)
1.0
0.5
0.0
ΔCm2/ ΔCm1
0.1
246 8
1
246 8
10
2
Inter-pulse-interval (s)
1.0
0.9
20 ms
Bsn
wt
Bsn
ΔEx4/5
1.0
0.5
0.0
ΔCm2/ ΔCm1
0.1
246 8
1
246 8
10
2
Inter-pulse-interval (s)
1.0
0.9
Q2/Q1
100 ms
Bsn
wt
Bsn
ΔEx4/5
-500
0
ICa(pA)
40
20
0
ΔCm(fF)
100 ms
Bsn
wt
(98 ms)
Bsn
ΔEx4/5(98 ms)
-400
-200
0
100
50
0
ΔCm(fF)
100 ms
Bsn
wt
(98 ms)
Bsn
ΔEx4/5(98 ms)
Figure 6. Slowed Vesicle Replenishment
Kinetics
(A) Example DCmresponses and Ca2+currents
(ICa) from Bsnwtand BsnDEx4/5IHCs upon two
20 ms depolarizations to maximum ICapotential,
separated by 98 ms.
(B) Same as (A) but with 100 ms depolarizations.
(C) Summary of paired-pulse DCm recordings
following 20 ms depolarizations. The graph shows
the ratio of response magnitudes between the
second and the first pulse (DCm2/DCm1) for
different inter-pulse-intervals.Notethe depression
in both genotypes, which is, however, more
pronounced in BsnDEx4/5IHCs (p < 0.01 for IPI of
98 ms; n [Bsnwt]: 23 to 32 IHCs; n [BsnDEx4/5]: 20
to 32 IHCs).
(D) Same as (C) but for 100 ms depolarizations.
Note the slight facilitation in BsnwtIHCs, but
consistent depression in BsnDEx4/5IHCs for short
IPIs, respectively (p < 0.01 for IPI of 98, 198, and
398 ms; n [Bsnwt] = 22 to 35 IHCs; n [BsnDEx4/5] =
20 to 39 IHCs).
The error bars in (C and D) represent SEM.
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Page 11

In vivo, we measured the recovery of the auditory nerve fiber
response following a masking sound as a proxy of the recovery
of the presynaptic RRP (Spassova et al., 2004). We used
a forward masking paradigm (Harris and Dallos, 1979; Spassova
et al., 2004) in which a 100 ms masking stimulus was separated
from a 15 ms probe stimulus by a variable silent interval ranging
between 2 and 512 ms (Figure 7A). Onset spike rates and adap-
ted spike rates in response to the masking stimulus were
reduced by a factor of 1.7 and 1.4, respectively, in BsnDEx4/5
(poststimulus time histograms [PSTHs]; Figure 7B). We found
an enhanced forward masking effect in BsnDEx4/5fibers when
comparing each probe response to its masker response (lower
ratio of spike rate for probe over spike rate for masker [averaged
over the first 5 ms of the probe and the masker] at 4-32 ms
interval for mutants, p < 0.05 each). There was also a trend
toward longer time of half-recovery from masking in BsnDEx4/5
(34.9±5.0msin mutantand 23.3±4.9 msin wild-type, p=0.13).
Taken together, the in vitro and in vivo results suggest
a disturbed replenishment of fusion-competent synaptic vesi-
cles in BsnDEx4/5IHCs. To what degree is the impaired sound
coding phenotype in BsnDEx4/5IHCs caused by a reduction in
the number of release sites or by their deficient refilling? To
answer this question, we quantified the forward masking data
by a model of sound-dependent RRP fusion and replenishment
combined with auditory nerve fiber refractoriness. The core
parameters of this model are the number of release sites,
sound-dependent rates (fusion rate constants and refilling rate
constants per release site in the presence and absence of
sound), and absolute and relative refractory periods (Supple-
mental Experimental Procedures). As those parameters are bio-
physically accessible quantities, the model can be used for
quantitative, mechanistic data analysis of auditory nerve fiber
responses in the context of cellular physiology.
A single set of parameters accurately reproduced PSTHs for
all nine recovery periods (Figure 7B and Figure S6). The very
same set of parameters also predicted the ratio of spike counts
(probe/masker)foranalysiswindowsof5msand13msfollowing
sound onset (Figure 7C). Importantly, the dominating difference
between the parameter sets for the two genotypes was a 35%
reduction in release site number for BsnDEx4/5(i.e., the maximal
capacity of the RRP; see dotted line in Figure 7B for a simulation
with wild-type release site number), while the fusion rates and
vesicular release probability remained virtually unchanged
(TableS2;consistentwithcapacitancemeasurements;Figure5).
Additionally, refilling rate constants were slightly reduced. When
assuming wild-type vesicle replenishment kinetics for BsnDEx4/5
fibers—while keeping all other model parameters for this geno-
type—the adapted spike rates were accordingly slightly
improved (see dashed line in Figure 7B).
C
Relative spike count (probe/masker)
A
B
64
ms
15 ms probe100 ms masker
821 ms interval to complete 1s cycle
100 ms
D
13 ms
5 ms
Exp. Sim.
Exp.
Sim.
1.5·Nslots
2.5·krefill, spont
1.2·krefill, stim
r
sv
Ca2+ channel
Bassoon
AMPA receptor
ribbon-occupiedribbonless
1.0
0.9
0.8
0.7
0.6
0.5
2832128512
1.0
0.9
0.8
0.7
0.6
0.5
2832128512
800
400
0
200150 100
500
800
400
0
800
400
0
Spike rate (s )
-1
200150100
500
800
400
0
Time (ms)
Recovery duration (ms)
Exp.
Sim.
BsnΔEx4/5
BsnΔEx4/5
Bsnwt
Bsnwt
Bsn
ΔEx4/5
Bsnwt
Bsn
ΔEx4/5
Figure 7. Comparison of Sound-Evoked Spike Rates In Vivo for
Bsnwtand BsnDEx4/5Mice
Reduced in vivo action potential rates and minimally enhanced forward mask-
inginauditorynervefibers canbeexplainedbyavesicle poolmodel combined
with spike refractoriness.
(A)Illustrationofthestimulusdesign:a100mstoneburst(masker)isseparated
from a 15 ms probe stimulus by a silent interval ranging between 2 and 512 ms
(example: 64 ms). Both stimuli were presented at characteristic frequency,
30 dB above threshold and with 2.5 ms rise-fall times. The interval between
two maskers was always 1 s. Each interval was repeated at least 50 times.
(B) Experimental results (blue [Bsnwt], and red [BsnDEx4/5]) and model-pre-
dicted (black) spike rates for recovery periods of 8 (upper) and 64 (lower) ms
in Bsnwt(left, n = 15) and BsnDEx4/5(right, n = 7) auditory nerve fibers. Dotted
line shows the prediction of a model with the set of BsnDEx4/5parameters
except Bsnwtrelease site number. Dashed line, instead, results from using
BsnDEx4/5parameters with Bsnwtvesicle replenishment kinetics.
(C) Experimental (dashed line, symbols representing means ± SEM) and
model-predicted (line) relative spike counts during the first 5 (left) and the first
13 (right) ms of the auditory nerve fiber response to the probe stimulus normal-
ized to the response to the masker stimulus. The error bars represent SEM.
(D) Schematic representation of tentative active zone structure at wild-type,
ribbon-occupied Bsn mutant, and ribbonless Bsn mutant afferent IHC
synapses, respectively. (Top) Sections; (bottom) view as seen from the
synaptic cleft. The illustration summarizes the findings of smaller CaV1.3
channel/membrane-proximal vesicle
a synaptic ribbon, altered substructure of CaV1.3 clusters in Bsn mutants.
complements in the absence of
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Please cite this article in press as: Frank et al., Bassoon and the Synaptic Ribbon Organize Ca2+Channels and Vesicles to Add Release Sites and
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Page 12

In summary, using the model as a quantification of the in vivo
results allowed us to draw conclusions about presynaptic quan-
tities from postsynaptic measurements. Generally, the model
validated our structural and functional findings, made indepen-
dently in vitro. Additionally, it advanced our mechanistic under-
standing by permitting the discrimination between a reduction
in the number of (1) generally available release sites and the
reduction in the (2) occupancy of those release sites: the param-
eters suggest that the reduced response amplitude in BsnDEx4/5
fibersisprimarilyduetoareductioninthetotalnumberofrelease
sites (35%) and to a lesser degree caused by a reduction in their
occupancy. These two effects combine such that in the model
the number of release sites occupied at rest is reduced to 50%
of wild-type, which is in agreement with the 55% reduction in
number of membrane-proximal vesicles observed in electron
micrographs of BsnDEx4/5AZs (Figure 1J).
DISCUSSION
In this study, we examined effects of genetic Bassoon disruption
at several structural and functional levels. EM tomography
revealed a spectrum of synapse morphologies from wild-type-
like to loosely anchored ribbons to ribbonless. Intriguingly, we
found that Bsn mutant synapses with a partially anchored ribbon
(ribbon occupied) exhibited an intermediate phenotype between
wild-type AZs and mutant ribbonless AZs. While fewest synaptic
Ca2+channels were found at ribbonless AZs in BsnDEx4/5IHCs,
the ribbon-occupied BsnDEx4/5AZs harbored more, but still
fewer, Ca2+channels than wild-type AZs—similar to the quanti-
fication of membrane-proximal vesicle number. Fast and sus-
tained exocytosis was reduced in proportion to the overall
reduction in membrane-proximal vesicle and Ca2+channel
number, while the Ca2+sensitivity of exocytosis remained
normal. Moreover, vesicle replenishment was impaired. A mech-
anistic computational model of synaptic transfer, used to fit the
in vivo data, independently supported morphological and func-
tional in vitro findings. We conclude that Bassoon disruption
and the associated ribbon loss reduces the number of functional
release sites, impairs their refilling, and consequently lowers
the RRP.
Structural Consequences of Bassoon Disruption
and Ribbon Loss
The most prominent phenotype of Bassoon disruption is the loss
of synaptic ribbons from a majority of AZs (Khimich et al., 2005;
tom Dieck et al., 2005). In contrast to retinal photoreceptors
(Specht et al., 2007), mature hair cells of Bsn mutants exhibited
some (although few) ribbon-occupied synapses at typical loca-
tions (Figures 1 and 2). Together with observations at Bsnwt
synapses without a ribbon, study of these ribbon-occupied
mutant synapses helped to test the role of the synaptic ribbon.
Bothsemiquantitative immunofluorescence
(Figures 2B and 2C) and confocal imaging of synaptic Ca2+influx
(Figure 4I1) revealed that ribbon presence was associated with
an increase in the number of Ca2+channels at BsnwtAZs. Using
STED microscopy, we furthermore observed a stripe-like
arrangement of the Ca2+channel cluster(s) at ribbon-occupied
BsnwtAZs. These structures were reminiscent of the electron-
microscopy
dense material seen in electron tomograms of AZs in mouse
IHCs (Figures 1E2–1H2) and frog saccular hair cells (Lenzi
et al., 2002), and the row-like arrays of intramembrane particles
observed in freeze-fracture electron micrographs (Roberts et al.,
1990; Saito and Hama, 1984). At all BsnDEx4/5synapses, this
CaV1.3 cluster geometry was dissolved into a pattern of small
spots (Figure 2F), similar to alterations of presynaptic densities
seen in electron tomography (Figures 1E2–1G2). This coinci-
dence supports the view that the CaV1.3 clusters are an integral
part of the presynaptic density (Lenzi et al., 2002). While this
difference in cluster geometry could, in principle, reflect a direct
effect of Bassoon loss, we did not find evidence for direct
interactions between Bassoon and CaV1.3 (heterologous ex-
pression; Figure S4).The observation of spots ratherthanstripes
at ribbon-occupied BsnDEx4/5synapses might also reflect a
decreased organizational impact of the ribbon when its
anchorage is loosened. It is interesting to note that the CAST/
ELKS1 homolog Bruchpilot has been implicated in clustering of
presynaptic Ca2+channels at the Drosophila neuromuscular
junction (Kittel et al., 2006). Bruchpilot is an integral component
of presynaptic electron-dense projections (T-bars, which were
absent in Bruchpilot mutants) and physically interacts with
presynaptic Ca2+channels, at least in vitro (Fouquet etal., 2009).
Our finding of abnormal clustering of synaptic Ca2+channels
uponBassoondisruptionissupportedbyacomparisonbetween
theassociatedreductioninwhole-cell(?20%)andsynapticCa2+
influx (?50%; both in the presence of BayK8644). The stronger
decrease in synaptic Ca2+influx indicates an increased fraction
of extrasynaptic Ca2+channels in mutant IHCs. Similar to Ca2+
channels, the number of membrane-proximal vesicles appears
to be greater when the ribbon is present. At least by trend, rib-
bonless mutant AZs showed the fewest vesicles, whereas the
presence of a ribbon increased this figure, but not to wild-type
levels.
Functional Consequences of Bassoon Disruption
and Ribbon Loss
How do these findings relate to synaptic function? Specifically,
how are the number of release sites—formed by vesicle docking
sites and closely colocalized Ca2+channels—and synaptic
exocytosis affected by Bassoon disruption? First, when probing
fast and sustained exocytosis in BsnDEx4/5IHCs, we observed
a decrease in amplitude that was roughly comparable to the
observedreductioninvesiclenumberandsynapticCa2+channel
number. Second, both intrinsic and apparent Ca2+cooperativity
of exocytosis was normal in BsnDEx4/5IHCs. Together, these
observations suggest that the coupling between Ca2+influx
through the remaining Ca2+channels and the fusion of the
remaining vesicles was unaffected. Yet one faces the caveat
that a static technique such as EM cannot distinguish between
fewer physical docking sites or their lower occupancy due to
impaired replenishment. Thus, distinguishing between these
two scenarios is aided by probing vesicle resupply, which was
slightly impaired in BsnDEx4/5mice (Figure 6). This finding stands
in agreement with the study of Hallermann and colleagues (Hal-
lermann et al., 2010), which shows that vesicle reloading at
a central synapse is impaired in Bsn mutants, evident by
enhanced synaptic depression during sustained high-frequency
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Page 13

trains. However, our study of a ribbon synapse revealed addi-
tional defects; fitting a mechanistic computational model to our
in vivo data (Figure 7 and Figure S6) indicated that synaptic
transmission at the IHC synapse of BsnDEx4/5was impaired
primarily due to a reduced number of functional release sites
(in accordance with our morphological and functional in vitro
data) and, to a lesser extent, their slower refilling under the given
stimulus protocol (Figure 7).
Toward Disentangling the Interplay between Bassoon
and the Synaptic Ribbon
Bassoon, via interaction with RIBEYE (tom Dieck et al., 2005),
contributes to ribbon anchorage. In hair cells, some residual
and partial ribbon anchorage is observed, probably involving
additional anchoring proteins. Those ribbon-occupied’’Bsn
mutant synapses were inferior to their wild-type counterparts
with regard to both Ca2+channel clustering and membrane teth-
ering of vesicles. These observations could either be explained
by (1) a direct effect of functional Bassoon loss, or (2) by a limited
capacity of ‘‘sick ribbons’’ ribbons to perform their task(s).
Several recent studies at ‘‘conventional’’ synapses show that
Bassoon is not required for synaptic transmission per se but is
involved in clustering (Mukherjee et al., 2010) and replenishment
of synaptic vesicles (Hallermann et al., 2010). Our results are
generally consistent with these findings; however, ribbon
synapses do require Bassoon also for basic synaptic transmis-
sion. It is likely that the more severe synaptic phenotype found
in IHCs reflects the perturbation of ribbon-supported functions.
For example, in contrast to Hallermann et al. (2010), we do find
a substantial reduction in the number of release sites in Bsn
mutant IHCs. Additionally, the trend toward fewer membrane-
tethered vesicles in general but more vesicles at ribbon-occu-
pied than ribbonless Bsn mutant synapses could, for example,
be explained by a combinatorial effect of primary Bassoon loss
and secondary Piccolo loss (Mukherjee et al., 2010) in the case
of ribbonless synapses. Interestingly, no evidence for a reduced
quantal content was found in BSN mutant cerebellar synapses
byHallermann etal.(2010)However,thecomplexnatureofinter-
actions between the numerous members of the cytomatrix
of the active zone (Schoch and Gundelfinger, 2006) demand
careful evaluation of ‘‘one-protein, one-function’’ hypotheses.
The absence of detectable direct effects of Bassoon disruption
onbasalsynaptictransmissionatconventionalsynapses(Haller-
mann et al., 2010; Mukherjee et al., 2010) and the intermediate
phenotypes seen in ribbon-occupied mutant synapses might
favor a hypothesis of ‘‘sick ribbons’’ over direct Bassoon effects
underlying the majority of observed synaptic and auditory
phenotypes in Bsn mutants. Future studies, including silencing
of ribbon components such asPiccolo and RIBEYE, arerequired
to further our understanding of the roles of the synaptic ribbon
and Bassoon for active zone structure and function as well as
their dynamic regulation.
EXPERIMENTAL PROCEDURES
A more detailed version of the Experimental Procedures is published in
Supplemental Information. Unless stated otherwise, all chemicals were
obtained from Sigma-Aldrich.
Animals
Mice with deletion of exons 4 and 5 of the Bassoon gene (BsnDEx4/5; Altrock
et al., 2003) or carrying a gene-trapped allele (Bsngt, Lexicon Pharmaceuticals,
Inc.), and wild-type littermates were used. All experiments were approved by
the University of Go ¨ttingen Board for Animal Welfare and the Animal Welfare
Office of the State of Lower Saxony.
Immunohistochemistry
Apical cochlearturnswere fixedinmethanolfor 20min at?20?C and prepared
aspreviouslydescribed (Khimich etal.,2005; Meyeretal.,2009).Thefollowing
antibodies were used: mouse anti-CtBP2 (BD Biosciences), rabbit anti-GluR2/
3 (Chemicon), rabbit anti-CaV1.3 (Alomone Labs), mouse anti-GluR2 (Chemi-
con), mouse anti-Sap7f407 to Bassoon (Abcam), and rabbit anti-BSN1.6 to
Bassoon (provided by E.D. Gundelfinger).
Confocal and STED Microscopy
Confocal image stacks were acquired with a Leica SP5 microscope and 1003
oil immersion objective. For STED-imaging, two different microscopes were
used: theLeica TCS STED (Figures 2D and 2E)and acustomapparatus(Harke
et al., 2008) with a resolution of around 80 nm. For size and shape analysis of
Ca2+channel clusters, XY scans were acquired after finding the fluorescence
maximum with a XZ-scan.
Electron Microscopy and Tomography
Cochleae were processed for electron microscopy as described (Meyer et al.,
2009 and Pangrsic et al., 2010). Thin sections were examined with a Philips
CM120 BioTwin transmission electron microscope (Philips Inc.) with a
TemCam F224A camera (TVIPS) at 20,0003 magnification. Images were
subsequently analyzed with iTEM software (Olympus). Tilt series from
250 nm sections were recorded at 27,5003 magnification in the range of
129?, then calculated with Etomo (http://bio3d.colorado.edu/).
Patch-Clamp and Confocal Ca2+Imaging of IHCs
IHCs from apical coils of freshly dissected organs of Corti (P20 through P31)
were patch-clamped as described (Moser and Beutner, 2000) and fluctuation
analysis (FA) was performed similarly as previously described (Meyer et al.,
2009). Currents were low-pass filtered at 8.5 kHz or 5 kHz and sampled at
100kHz(FA)or40kHz(Ca2+currents,DCmmeasurements),respectively.Cells
with holding current > ?50 pA were discarded. Ca2+currents were further iso-
lated with a P/n protocol. In FA and Ca2+current activation recordings, series
resistance was compensated online (20%–50%; t = 10 ms). Residual series
resistance averaged 4.4 ± 0.4 MU (Bsnwt; n = 35 ensembles) and 4.3 ±
0.3 MU (BsnDEx4/5; n = 33 ensembles) in FA experiments. Flash photolysis
was performed essentially as described in Beutner et al. (2001). Confocal
Ca2+imaging was performed as described (Frank et al., 2009).
Single-Unit Recordings
Single-unit recordings from auditory nerve fibers of 6- to 10–week-old Bsnwt
and BsnDEx4/5mice (n = 7 each) were performed as described by Taberner
and Liberman (2005) and Buran et al. (2010).
Data Analysis
Data analysis was performed with Matlab (Mathworks), Igor Pro (Wavemet-
rics), and ImageJ software and is described in more detail in Supplemental
Information. Two-tailed t tests or the Mann-Whitney-Wilcoxon test were
used for statistical comparisons between two samples (*p < 0.05, **p < 0.01,
***p < 0.001).
SUPPLEMENTAL INFORMATION
Supplemental Information includes Supplemental Experimental Procedures,
six figures,twotables, and twomovies and canbefound withthisarticleonline
at doi:10.1016/j.neuron.2010.10.027.
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Page 14

ACKNOWLEDGMENTS
We thank S. Blume, N. Dankenbrink-Werder, A. Gonzalez, M. Ko ¨ppler, and
B. Kracht for expert technical assistance. This work was supported by grants
of the Max Planck Society (Tandem-Project grant to Nils Brose and T.M.), the
German Research Foundation Fellowship to N.S., Center for Molecular Phys-
iology of the Brain Grant FZT-103 to T.M. and A.E., the German Federal
Ministry of Education and Research (01GQ0810, Bernstein Focus for Neuro-
technology)to T.M. and A.E., the State of Saxony-Anhalt/European Structural
Funds (EFRE-IfN C2/1) to E.D.G., and by the National Institutes of Health
(Grants DC0188 to M.C.L., DC009433 and HL087120 to A.L., and T32 AI
07260 to K.E.B.) and T32 AI 07260 (to K.E.B). M.A.R. and T.P. were supported
by fellowships of the Alexander von Humboldt Foundation. A.N. is a Fellow of
the Bernstein Center for Computational Neuroscience Go ¨ttingen. The study
was designed by T.M., T.F., N.S., A.N., and D.R. The experimental work was
performed by T.F. (Ca2+imaging, Ca2+current, and Cmrecordings), M.A.R.
(confocal and STED microscopy), N.S. (single-unit recordings under supervi-
sion of M.C.L.), T.P. (flash photolysis), D.K. (confocal microscopy), and D.R.
(electron microscopy). A.E. and B.H. contributed to STED microscopy, and
A.E. contributed to image analysis. A.N. performed modeling and contributed
todataanalysis.A.F.andE.D.G.providedthemice(genetrapmutantincollab-
oration with Lexicon Pharmaceuticals) and discussion. K.E.B and A.L. per-
formed protein-protein interaction experiments. T.M., T.F., N.S., A.N., and
M.A.R. prepared the manuscript.
Accepted: October 8, 2010
Published: November 17, 2010
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Please cite this article in press as: Frank et al., Bassoon and the Synaptic Ribbon Organize Ca2+Channels and Vesicles to Add Release Sites and
Promote Refilling, Neuron (2010), doi:10.1016/j.neuron.2010.10.027
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