The Rockefeller University Press $30.00
J. Cell Biol. Vol. 190 No. 6 1067–1077
Correspondence to Jens Rettig: firstname.lastname@example.org
Abbreviations used in this paper: CAPS, calcium-dependent activator protein
for secretion; DKO, double KO; KO, knockout; LDCV, large dense-core vesicle;
MHD, Munc13 homology domain; RRP, readily releasable pool; SRP, slowly
releasable pool; SV, synaptic vesicle; UPP, unprimed pool.
In neuroendocrine cells and synaptic terminals, only a fraction of
the secretory vesicles that are docked at the plasma membrane
can be released upon stimulation, indicating that a maturation or
priming step, which renders secretory vesicles fusion competent,
must take place after docking (Parsons et al., 1995; Plattner et al.,
1997). Interestingly, the pool of primed and fusion-competent
vesicles is heterogeneous in many secretory systems (Bittner and
Holz, 1992). In chromaffin cells, for example, high time resolu-
tion experiments revealed two populations of vesicles with dif-
ferent release rates, a readily releasable pool (RRP) and a slowly
releasable pool (SRP), which produce two phases of release
(Voets et al., 1999).
Several recent experiments showed that calcium-
dependent activator protein for secretion (CAPS) proteins play
a key role in the priming of large dense-core vesicles (LDCVs)
and synaptic vesicles (SVs; Stevens and Rettig, 2009). CAPS1
was discovered as a cytosolic factor that is required for
regulated fusion of LDCVs in PC12 cells (Walent et al., 1992).
Subsequently, it was shown that CAPS1 is a homologue of the
Caenorhabditis elegans protein UNC-31, whose mutation
leads to an uncoordinated phenotype with motor deficits (Ann
et al., 1997) and depresses release of SVs (Jockusch et al.,
2007) and LDCVs (Elhamdani et al., 1999). Deletion of CAPS
proteins causes a strong reduction in the size of the releasable
pool of vesicles in several organisms (Renden et al., 2001;
Speidel et al., 2005; Jockusch et al., 2007; Speese et al., 2007;
Liu et al., 2008).
In mouse and human, there are two CAPS genes encoding
isoforms that both contain a Munc13 homology domain (MHD;
Speidel et al., 2003). This domain is part of the minimal struc-
ture required for the function of Munc13-1 (Basu et al., 2005;
Madison et al., 2005; Stevens et al., 2005), a vesicle-priming
protein in neurons (Tokumaru and Augustine, 1999) and neuro-
endocrine cells (Ashery et al., 2000). The sequence required for
priming activity by Munc13-1 consists of a stretch of 672 amino
acids, including both MHDs (Basu et al., 2005; Madison et al.,
2005; Stevens et al., 2005), that has also been termed the MUN
soluble N-ethyl-maleimide sensitive fusion protein attach-
ment protein (SNAP) receptor complex consisting of syn-
taxin, SNAP-25, and synaptobrevin. Using mice lacking
both isoforms of the calcium-dependent activator protein
for secretion (CAPS), we show that LDCV priming in ad-
renal chromaffin cells entails two distinct steps. CAPS is re-
quired for priming of the readily releasable LDCV pool and
riming of large dense-core vesicles (LDCVs) is
a Ca2+-dependent step by which LDCVs enter a
release-ready pool, involving the formation of the
sustained secretion in the continued presence of high Ca2+
concentrations. Either CAPS1 or CAPS2 can rescue se-
cretion in cells lacking both CAPS isoforms. Furthermore,
the deficit in the readily releasable LDCV pool resulting
from CAPS deletion is reversed by a constitutively open
form of syntaxin but not by Munc13-1, a priming protein
that facilitates the conversion of syntaxin to the open con-
formation. Our data indicate that CAPS functions down-
stream of Munc13s but also interacts functionally with
Munc13s in the LDCV-priming process.
Two distinct secretory vesicle–priming steps in
adrenal chromaffin cells
Yuanyuan Liu,1 Claudia Schirra,1 Ludwig Edelmann,2 Ulf Matti,1 JeongSeop Rhee,4 Detlef Hof,1 Dieter Bruns,1
Nils Brose,4 Heiko Rieger,3 David R. Stevens,1 and Jens Rettig1
1Institut für Physiologie and 2Institut für Molekulare Zellbiologie, Universität des Saarlandes, 66421 Homburg, Germany
3Institut für Theoretische Physik, Universität des Saarlandes, 66123 Saarbrücken, Germany
4Max-Planck-Institut für Experimentelle Medizin, Abteilung Molekulare Neurobiologie, 37075 Göttingen, Germany
© 2010 Liu et al. This article is distributed under the terms of an Attribution–Noncommercial–
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License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at
T H E J O U R N A L O F C E L L B I O L O G Y
JCB • VOLUME 190 • NUMBER 6 • 2010 1068
loss of CAPSs, indicating that CAPS function may indeed be
similar to that of Munc13s. However, expression of Munc13-1
does not enhance priming in the absence of CAPS1, although its
expression in the presence of CAPS1 leads to the expected en-
hancement of secretion. These results indicate that LDCV priming
in mouse chromaffin cells involves the opening of the protein
syntaxin and that opening of syntaxin is facilitated by CAPS.
Both CAPS isoforms enhance priming activity and preferen-
tially prime LDCVs of the RRP.
CAPS2 restores secretion to wild-type
levels in CAPS DKO chromaffin cells
Deletion of both CAPS genes in mouse chromaffin cells (CAPS
DKO) causes an 50% decrease in exocytosis as a result of a
strong (>50%) decrease in the size of the RRP and almost com-
plete block of sustained release (Liu et al., 2008). We stimulated
catecholamine secretion from mouse chromaffin cells using flash
photolysis of caged calcium to examine the ability of virally ex-
pressed CAPS2 protein to restore secretion in mouse chromaffin
cells lacking both CAPS1 and CAPS2. After UV flash illumina-
tion, secretion (as measured by a membrane capacitance change)
domain (Basu et al., 2005). Interestingly, an MUN domain–like
structure has also been identified in CAPS (Koch et al., 2000;
Hammarlund et al., 2008). In light of the role of the MUN do-
main in priming by Munc13s and the accumulating evidence
that CAPSs also promote priming (Stevens and Rettig, 2009), an
attractive hypothesis is that CAPSs carry out their priming func-
tion in a fashion similar to that of Munc-13s. Munc13-1 functions
by binding to syntaxin (Betz et al., 1997; Richmond et al., 2001),
promoting a conformational change in syntaxin that allows it to
engage in SNARE complex formation (Dulubova et al., 1999).
In mouse chromaffin cells, deletion of CAPSs (double
knockout [KO; DKO]) causes a reduction of the releasable LDCV
pool and of sustained release (Liu et al., 2008), the latter of which
occurs in the continued presence of elevated Ca2+ because vesi-
cles that are primed are immediately released and, thus, is an in-
dicator of vesicle priming. In addition, the loss of CAPS1 results
in reduced transmitter loading into chromaffin granules (Speidel
et al., 2005). Expression of CAPS1 in the KO background re-
stores normal transmitter loading (Speidel et al., 2005) and secre-
tion, increases the readily releasable LDCV pool, and enhances
sustained release (Liu et al., 2008).
In this study, we show that both CAPS1 and CAPS2 facili-
tate LDCV priming and that open syntaxin can overcome the
Figure 1. CAPS2 restores secretion to wild-type levels in CAPS DKO cells. (A) Responses to flash photolysis of caged calcium in CAPS DKO cells (n = 19)
and CAPS DKO cells expressing CAPS2 protein (n = 24). Upon elevation of intracellular calcium (top), the resulting change in membrane capacitance
(middle) shows a clear enhancement in those cells expressing CAPS2. Carbon fiber amperometry verifies that the observed increase in capacitance is
caused by an increase in (bottom) catecholamine release. (B) Analyses of the kinetics of the capacitance traces yield estimates of the releasable pools and
the sustained release rate. The RRP was strongly enhanced by CAPS2 expression (***, P < 0.001), whereas the SRP was unaffected. The rate of sustained
release in the period in which calcium remained elevated was also enhanced after CAPS2 expression (P < 0.001). (C) CAPS1 enhances secretion in wild-
type (WT) cells. Overexpression of CAPS1 in wild-type chromaffin cells (n = 26) results in a modest enhancement of secretion as compared with untreated
cells (n = 24). (D) A modest strengthening of the RRP of vesicles did not reach statistical significance. (E) CAPS2 expression enhances the exocytotic burst
in wild-type mouse chromaffin cells. Responses to a calcium stimulus induced by flash photolysis of caged calcium (top) in CAPS2-expressing cells (n = 31)
and wild-type chromaffin cells (n = 33) indicate that secretion is enhanced (middle), and this increase is mirrored by an increase in catecholamine release
as indicated by amperometric detection (bottom). (F) There was a significant increase in the RRP (**, P < 0.01) as compared with that of untreated wild-type
cells, with no difference in the SRP and a reduction in the sustained rate of release (P < 0.01). Error bars indicate mean ± SEM.
1069CAPS primes to the readily releasable pool • Liu et al.
60.8 fF and 60.7 fF in CAPS1-expressing wild-type cells and
control cells, respectively. The sustained release rate was slightly
greater in CAPS1-expressing cells (26.1 ± 5.2 fF) than in con-
trol cells (19.1 ± 2.4 fF; Fig. 1 D).
We also tested whether overexpression of CAPS2 in wild-
type cells would have a similar effect as CAPS1 on secretion.
As shown in Fig. 1 E, introduction of CAPS2 in wild-type chromaf-
fin cells resulted in a comparable, selective increase in the RRP.
Although the RRP was enhanced (CAPS2 expression, 147.3 ±
18.4 fF [n = 31]; vs. control cells, 85.1 ± 8.1fF [n = 33]), the
SRP was unchanged (76.2 ± 8.3 vs. 76.4 ± 10.0 fF), and the sus-
tained release slightly decreased (27.1 ± 3.0 vs. 16.5 ± 2.0 fF;
Fig.1, E and F). These results indicate that both CAPS1 and
CAPS2 overexpression promote priming into the RRP in wild-
type chromaffin cells.
Open syntaxin restores RRP in CAPS DKO
cells to wild-type levels
In subsequent experiments, we tested whether expression of an
open form of syntaxin (syntaxin1A L165A/E166A; Dulubova
et al., 1999) can reverse the secretion deficit in CAPS DKO chro-
maffin cells (Fig. 2 A). Expression of open syntaxin in chro-
maffin cells from CAPS DKO mice led to strongly enhanced
secretion (threefold increase) as compared with that of DKO
cells. This enhancement was accounted for by an approximately
threefold increase in the RRP size (open syntaxin–expressing
DKO cells, 129.2 ± 21.8 fF [n = 24]; vs. CAPS DKO cells, 43.5 ±
8.9 fF [n = 23]; Fig. 2 B). There was also a modest increase in the
SRP size in open syntaxin–expressing DKO cells (79.5 ± 11.3 fF)
relative to untreated CAPS DKO cells (DKO, 52.3 ± 8.9 fF),
but there was virtually no detectable sustained component in
either group of cells. Application of a second flash after a 2-min
occurred in an exocytotic burst consisting of an RRP and an
SRP followed by a sustained release phase (Fig. 1). Expression of
CAPS2 in chromaffin cells from CAPS DKO mice resulted in a
strong enhancement of the exocytotic burst and of sustained re-
lease as compared with DKO cells (Fig. 1 A). By fitting the exo-
cytotic burst as the sum of two exponentials and sustained release
as a linear phase, we estimated the size of the RRP and SRP,
their time constants, and the rate of sustained release. There was
a significant increase in the RRP of DKO cells expressing CAPS2
(125.3 ± 25.8 fF; n = 24) compared with that of untreated DKO
cells (42.8 ± 18.6 fF; n = 19; P < 0.001; Mann-Whitney U test)
but no change in the release time constant of the RRP (CAPS2,
23.5 ± 5.2 ms; vs. DKO, 23.2 ± 1.4 ms). The amplitude of the
SRP was not altered in CAPS2-expressing DKO cells (51.7 ±
12.2 fF; n = 24) when compared with DKO cells (51.8 ± 7.5 fF;
n = 19). Sustained release, which was not measurable in DKO
cells, was 19.2 ± 3.7 fF/s in the CAPS2-expressing DKO cells
(Fig. 1 B). Thus, as is the case for CAPS1, expression of CAPS2
in CAPS DKO cells restores secretion with a selective effect on
the RRP and the sustained release phase.
CAPS1 and CAPS2 overexpression
increase the RRP size in wild-type
We next tested how overexpression of CAPS1 in wild-type cells
affects secretion. The results of this experiment are shown in
Fig. 1 C. CAPS1 overexpression in wild-type chromaffin cells
led to a modest enhancement of the exocytotic burst and sus-
tained release after photolysis of caged Ca2+, neither of which
was statistically significant. The RRP after CAPS1 overexpres-
sion in wild-type cells (n = 24) was 105 ± 16.1 fF as compared
with 79 ± 13.8 fF in control cells (n = 26). The SRP sizes were
Figure 2. Open syntaxin restores secretion
in CAPS DKO cells to wild-type levels. (A) Re-
sponses to flash photolysis of caged calcium
in CAPS DKO cells (n = 23) and CAPS DKO
cells expressing open syntaxin (n = 24) show
that open syntaxin restores secretion. The burst
component of the capacitance response is
much greater in the open syntaxin–expressing
cells (middle), and the catecholamine release
in amperometric recordings is also strongly
enhanced (bottom). (B) Estimates of the releas-
able pools and the sustained rate indicate that
open syntaxin strongly enhances the RRP (***,
P < 0.001), although the SRP is also enhanced
(this difference was not significant). Note that
there is little sustained release after the burst.
(C) Examination of a second flash stimulation
to the same cells after a 2-min recovery period
shows that open syntaxin–treated CAPS DKO
cells recover poorly after flash stimulation.
(D) Pool analysis shows that open syntaxin–
expressing CAPS DKO cells recover more
poorly and do not exhibit a greater RRP after a
second flash. **, P < 0.01. Error bars indicate
mean ± SEM.
JCB • VOLUME 190 • NUMBER 6 • 2010 1070
We tested the residual secretory capacity that remains
after the ramp stimulus by applying a flash 3 s after the end of
the ramp stimulus. The DKO cells secreted slightly more in re-
sponse to the postramp flash (131.0 ± 31.7 fF; n = 22) than did
the open syntaxin–expressing DKO cells (98.3 ± 24.9 fF; n = 23;
Fig. 3 B). The secretion induced by the flash was equivalent to
65% of the total secretion in the DKO cells and to 25% of
the total secretion in the open syntaxin–expressing DKO cells.
Thus, in the open syntaxin–expressing DKO cells, 75% of the
secretory capacity was released during the ramp stimulation,
whereas only 35% of the secretory capacity was released by the
ramp stimulation in the DKO cells. The kinetics of release after
the flash stimulation were slow and similar in both groups, indi-
cating that residual release originated from an SRP of vesicles.
Total secretion was much greater in the open syntaxin–expressing
DKO cells in these ramp stimulation experiments, as was the
case in the original flash experiments. Thus, open syntaxin by-
passes the requirement for CAPS and generates a large pool of
Chromaffin cells from mutant mice that only express open
syntaxin are characterized by a large reduction in the number of
morphologically docked vesicles (Gerber et al., 2008). The lack
of sustained release and the apparent fast exhaustion of release
that we observed in chromaffin cells expressing open syntaxin
might therefore be because of a reduction in vesicle docking
rather than to effects on priming. To test this, we expressed open
syntaxin in wild-type cells (Fig. 4 A). Open syntaxin expression
led to an approximately twofold increase in the RRP size (209.3 ±
26.0 fF; n = 24) relative to untreated wild-type cells (95.8 ± 9.4 fF;
n = 22) and enhanced the SRP size, although this effect was not
statistically significant (open syntaxin–expressing wild-type
cells,146.6 ± 25.3 fF; vs. untreated wild-type cells, 101.7 ± 14 fF)
but reduced the sustained phase (open syntaxin–expressing wild-
type cells, 4.6 ± 2.1 fF/s; vs. untreated wild-type cells, 22.5 ±
3.8 fF; Fig. 4 B).
As in CAPS DKO cells, wild-type cells expressing open
syntaxin recovered from stimulation poorly, as illustrated by re-
ductions of all phases of release in the responses to a second flash
stimulation (Fig. 4 C). In the responses to a second flash, the RRP
size was 104.6 ± 18.9 fF in open syntaxin–expressing wild-type
cells (n = 20) versus 136.8 ± 21.5 fF in untreated wild-type cells
(n = 18). The SRP was 56.3 ± 10.5 fF in open syntaxin–expressing
wild-type cells versus 99.8 ± 13.0 fF in untreated wild-type cells.
Sustained release was 1.6 ± 1.0 fF/s in open syntaxin–expressing
cells and 11.4 ± 2.5 fF/s in untreated cells (Fig. 4 D). The reduc-
tion of sustained release is thus independent of CAPS function.
These experiments indicate that open syntaxin promotes priming
into the RRP but simultaneously reduces sustained release and
pool recovery, possibly because of a reduction in the number of
Open syntaxin expression leads to reduced
We next analyzed the distribution of LDCVs in chromaffin cells
to determine whether open syntaxin causes a docking defect
such as the one reported for mutant mice expressing only open
syntaxin (Gerber et al., 2008). We compared the distributions of
recovery period resulted in a reduced open syntaxin response
with a small RRP and no sustained component, whereas in the
CAPS DKO cells, the response to the second flash was equiva-
lent to the first response (Fig. 2, C and D). These findings are
compatible with the view that the supply of primable vesicles is
limited in the open syntaxin–expressing cells.
Open syntaxin expression in CAPS DKO
cells leads to rapid exhaustion of release
Stimulation of mouse chromaffin cells with a slowly rising cal-
cium concentration (calcium ramp) leads to biphasic secretion,
the late phase of which is likely the result of priming during
the stimulus, i.e., analogous to the sustained release during
flash photolysis (Sørensen et al., 2002). In CAPS DKO cells,
the late phase of secretion is either very small or absent (Liu
et al., 2008). We examined the effects of ramp stimulation in
DKO cells in which open syntaxin was expressed. In agree-
ment with the data obtained by flash stimulation (Fig. 2), open
syntaxin–expressing DKO cells showed strongly enhanced se-
cretion (295.5 ± 38.9 fF; n = 29) as compared with untreated
DKO cells (72.2 ± 13.2 fF; n = 27; Fig. 3 A). Secretion during
ramp stimulation started slowly, so we used the second deriva-
tive of the capacitance trace to more accurately determine the
increase in slope at the beginning of the secretory phase.
The Ca2+ value at the time of a peak in the second derivative of the
smoothed capacitance trace (Schonn et al., 2008), taken as the
threshold Ca2+ concentration required for secretion, was 850 nM
in both DKO and open syntaxin–expressing DKO cells. Both
the response of DKO cells and that of the open syntaxin–
expressing DKO cells were sigmoid. In spite of the greater se-
cretion in the open syntaxin–expressing DKO cells (or perhaps
as a result of this), secretion of the open syntaxin–expressing
cells reached a plateau before the end of the stimulation. This
was not the case in the DKO cells, which secreted throughout
Figure 3. The releasable pools in CAPS DKO cells expressing open syn-
taxin are rapidly exhausted. (A) The free calcium concentration (top)
and the capacitance change (bottom) are shown. Those cells expressing
open syntaxin exhibit very strong secretion (n = 29) compared with CAPS
DKO cells not expressing open syntaxin (n = 27). (B) To determine the
amount of secretion remaining after the calcium ramp stimulation, a flash
was applied 3 s after the ramp ended to increase calcium to high levels.
The residual secretion in the CAPS DKO cells was larger than that of the
DKO cells expressing open syntaxin and accounted for 65% of the total
secretion, whereas the flash response in the open syntaxin–expressing
CAPS DKO cells accounted for 25% of the total secretion. Error bars
indicate mean ± SEM.
1071 CAPS primes to the readily releasable pool • Liu et al.
expression of a full-length Munc13-1–GFP construct in CAPS
DKO cells failed to restore secretion (Fig. 6 A). Unexpectedly, all
phases of flash-induced secretion in DKO cells expressing
Munc13-1 (RRP, 31.1 ± 8.5 fF; SRP, 24.7 ± 5.6 fF; sustained,
1.9 ± 1.0 fF/s; n = 23) were reduced compared with untreated
DKO cells (RRP, 48.4 ± 12.8 fF; SRP, 58.1 ± 10.0 fF; sustained,
2.9 ± 1.6 fF/s; n = 22; Fig. 6 B). We also tested whether expres-
sion of a truncated construct of Munc13-1 containing the mini-
mal priming domain (Stevens et al., 2005) can rescue secretion in
CAPS DKO cells, but expression of this construct also failed to
restore secretion in CAPS DKO cells, although it enhanced secre-
tion in wild-type mouse chromaffin cells (unpublished data).
Munc13-1 enhances secretion only in the
presence of CAPS1
Munc13-1 expression strongly enhances secretion in wild-type
mouse chromaffin cells (Stevens et al., 2005) but reduces secre-
tion in CAPS DKO cells (Fig. 6, A and B). In view of this dis-
crepancy, we tested whether the lack of a positive Munc13-1
effect on secretion seen in CAPS DKO cells might be directly
related to the lack of CAPS. We first examined the ability
of Munc13-1 to enhance secretion in cells from CAPS1+/
CAPS2/ mice (Fig. 6 C). In these cells, Munc13-1 expression
(n = 25) enhanced all phases of secretion significantly when
compared with untreated cells (n = 23). For CAPS1+/ CAPS2/
cells expressing Munc13-1, the RRP was 281.5 ± 39.9 fF, the
SRP was 278.3 ± 44.0 fF, and the sustained release was 50.7 ±
7.2 fF/s. In untreated CAPS1+/ CAPS2/ cells, the RRP was
145.8 ± 19 fF, the SRP was 82.5 ± 10.0 fF, and the sustained re-
lease was 20.0 ± 3.3 fF/s (Fig. 6 D).
LDCVs in untreated chromaffin cells from wild-type (n = 21)
and CAPS DKO cells (n = 16) to those in wild-type cells after
expression of open syntaxin (n = 10) and in CAPS DKO after
expression of open syntaxin (n = 7). We determined the shortest
distance from the plasma membrane of all identifiable LDCVs
in chromaffin cells derived from wild-type or CAPS DKO em-
bryonic day (E) 18/postnatal day (P) 0 mice either with or with-
out expression of open syntaxin. Representative micrographs
are shown in Fig. 5 (A–D). The distributions of measured dis-
tances (Fig. 5 E) showed a clear reduction in the fraction of
LDCVs in close apposition to the membrane in open syntaxin–
overexpressing CAPS DKO (62% reduction) and wild-type cells
(77% reduction), as compared with untreated cells of CAPS
DKO and wild-type mice. We conclude from these data that the
reduction in sustained release by expression of open syntaxin is
the result of a reduction in the transport of LDCVs to the plasma
membrane, i.e., docking.
Munc13-1 does not restore secretion
in CAPS DKO cells
The results of expressing open syntaxin indicate that LDCV prim-
ing is facilitated by opening of syntaxin, thus enabling syntaxin
to engage in SNARE complex formation. The MHD domain of
CAPS may be involved in the conformational change of syntaxin
required for priming, as is believed to be the case for the MHD
domains of Munc13s (Basu et al., 2005; Madison et al., 2005;
Stevens et al., 2005). If this were indeed the case, secretion in
chromaffin cells from CAPS DKO mice should also be restored
by expression of Munc13-1. Surprisingly, but in agreement with
data in cultured hippocampal neurons (Jockusch et al., 2007),
Figure 4. Expression of open syntaxin in wild-
type cells enhances the RRP selectively. (A) Flash
photolysis of caged calcium produces a
larger burst of secretion in open syntaxin–
expressing wild-type (WT) cells (n = 24) when
compared with cells not expressing open syn-
taxin (n = 22). This is mirrored in an increase
in catecholamine release. (B) Analysis of the
kinetics of the releasable pools and the rate of
sustained release show that the RRP is strongly
enhanced (**, P < 0.01), whereas the SRP is
unaltered. The sustained release is reduced
(***, P < 0.001). (C) Examination of the
second flash response after a 2-min recovery
period shows a deficit in refilling pools emp-
tied by a flash stimulation in open syntaxin–
expressing wild-type cells. (D) Analysis of pools
in the responses to the second stimulation.
Error bars indicate mean ± SEM.
JCB • VOLUME 190 • NUMBER 6 • 2010 1072
(n = 12; Fig. 7 C). Thus, it appears that Munc13-1 requires
CAPS1 to promote secretion in chromaffin cells, indicating that
the two proteins interact functionally in the process of chro-
maffin granule priming.
The CAPS effect on RRP does not explain
the lack of sustained release
CAPS deletion reduces total secretion from chromaffin cells,
with a strong reduction in the RRP of LDCVs. However, the
effect on sustained release is much greater (Fig. 1; Speidel et al.,
2005; Liu et al., 2008). The current model for LDCV matura-
tion and release in chromaffin cells involves a linear path from
a depot pool to a docked pool (unprimed pool [UPP]) followed
by priming into the SRP and further maturation into the RRP
Strikingly, when we examined the effect of Munc13-1 ex-
pression in cells from CAPS1/ CAPS2+/ mice, we found that
Munc13-1 expression (n = 23) not only failed to increase secretion,
but even decreased it (Fig. 7 A) relative to untreated cells (n = 22),
as was the case in the DKO cells (Fig. 6, A and B). The RRP of
CAPS1/ CAPS2+/ cells expressing Munc13-1 was 72.8 ±
16.6 fF, whereas in untreated CAPS1/ CAPS2+/ cells, the RRP
was 98.8 ± 12.4 fF. The SRP of Munc13-1–expressing CAPS1/
CAPS2+/ cells was 33.8 ± 5.7 fF and that of untreated cells
was 50.3 ± 7.5 fF, whereas the sustained release in Munc13-1–
expressing CAPS1/ CAPS2+/ cells was 6.3 ± 2.2 fF/s and that
in untreated CAPS1/ CAPS2+/ cells was 9.2 ± 2.4 fF/s
(Fig. 7 B). In contrast, expression of Munc13-1 in wild-type
(n = 15) cells resulted in the expected robust enhancement of
all phases of secretion as compared with wild-type controls
Figure 5. Open syntaxin expression reduces
the numbers of docked vesicles in chromaffin
cells. (A and B) Representative electron micro-
graphs of chromaffin cells from wild-type (WT)
and CAPS DKO mice are shown. N, nucleus;
M, mitochondria. (C and D) Representa-
tive electron micrographs of chromaffin cells
from wild-type and CAPS DKO mice 6 h after
tion are shown. (A–D) Bars, 2 µm. (E) Relative
frequency distribution of the granule distances
from the plasma membrane in chromaffin cells
from CAPS DKO (n = 16) and wild-type mice
(n = 21) and CAPS DKO with open syntaxin
expression (n = 7) and wild-type with open
syntaxin expression (n = 10). Although there
was no difference in docked vesicles in CAPS
DKO cells compared with wild-type cells, ex-
pression of open syntaxin led to a strong re-
duction in LDCVs adjacent to the membrane in
both populations. Bin width was 60 nm. Error
bars indicate mean ± SEM. (F) High magnifi-
cation inset taken from the boxed region in A.
Dashed line shows the distance of 60 nm from
the plasma membrane that was taken to define
docked vesicles. Bar, 200 nm.
1073 CAPS primes to the readily releasable pool • Liu et al.
k-2 or increasing k2 increases the RRP and the total number
of primed vesicles, although leaving SRP virtually unchanged
(Fig. 8 A, dashed line). This will not alter the sustained release
rate, which depends only on UPP size and the rate constants
k1 and k-1 (Fig. 8 B). A decrease in RRP priming would only de-
crease sustained release in a parallel scheme in which total prim-
ing is the sum of SRP and RRP priming. Such a scheme does not
fit the results by Voets et al. (1999), which showed that refilling
of the RRP is accompanied by a simultaneous SRP decrease of
The effects on pool size of CAPS deletion and of CAPS
rescue in the DKO background are readily simulated by a re-
spective decrease or increase of the rate constant of priming
(transfer) into the RRP (k2). The effects of open syntaxin ex-
pression on the releasable pools in both wild-type and DKO
cells can be modeled in the same way. However, as discussed in
the previous paragraph, this does not explain the observed lack
of sustained release in CAPS DKO cells and cells overexpress-
ing open syntaxin.
Because open syntaxin causes a docking deficit, we ex-
amined to what extent this might explain the lack of sustained
release in our open syntaxin experiments (docking rate re-
duced to 45%). The ratio k1/k-1 combined with the UPP size
controls overall priming rate, explaining why a depletion of
the UPP will lead to lower sustained rates and smaller pool
sizes (Fig. 8 C). The effect on pool size can be overcome by an
increase in conversion from the SRP to the RRP (k2 is en-
hanced or k-2 is decreased; k2/k-2 > 1), which increases burst
and RRP size.
Adding an increase in the rate constant for priming the
RRP (k2) to the reduced forward docking rate (k0) produced a
Secretion from the SRP and RRP requires the appropri-
ate Ca2+ stimulus. Manipulation of the rate constants k2 and k-2
will alter the balance between SRP and RRP and thus alter
RRP size. Altering k1 or k-1 will change the numbers of primed
vesicles at steady state without affecting the SRP/RRP ratio.
We have performed numerical simulations of the vesicle pools
and their release using the following scheme (see Materials
Using the constants suggested by Sørensen (2004), we
solved the equations for steady-state pool size for UPP, SRP, and
RRP and then solved the differential equations describing the
changes in pool sizes, release rate, and release of vesicles (after
a stepwise increase in [Ca2+]i to 10 µM) over time using a fourth-
order Runge-Kutte integration.
Because priming is considered the step that delivers releas-
able vesicles, the k1 step would be equivalent to priming in a
linear model. This step is calcium dependent, with a Kd for Ca2+
near 2.5 µM (Voets, 2000). This results in an increase in priming
when basal [Ca2+]i changes at subthreshold concentrations (100
to 900 nM) and a strong increase in priming after flash pho-
tolysis, which causes the strong sustained release observed under
these conditions. Increasing or decreasing k1 leads to an increase
or decrease, respectively, in the size of the primed vesicle pool
with no change in the SRP/RRP ratio (Ca2+ = 300 nM, k1 0.003;
Ca2+ = 600 nM, k1 0.006; Fig. 8 A).
The slowly releasable vesicles then undergo an additional
maturation step (k2), making them readily releasable. Decreasing
Figure 6. Expression of Munc13-1 does not
enhance secretion in CAPS DKO chromaffin
cells. (A) The free calcium (top), capacitance
changes (middle), and amperometric record-
ings (bottom) in CAPS DKO cells (n = 22) and
CAPS DKO cells expressing Munc13-1 (n = 23)
are shown. Munc 13–1 expression did not re-
store release in CAPS DKO cells. (B) Estimates
of the kinetic parameters show that all phases
of release were suppressed in cells express-
ing Munc13-1. (C) Munc13-1 overexpression
in cells heterozygous for CAPS1 (CAPS1+/
CAPS2/) has the expected enhancing effect
on secretion. Secretion was enhanced in
CAPS1 heterozygotes expressing Munc13-1
(n = 25) when compared with CAPS1 hetero-
zygotes not expressing Munc13-1 (n = 23).
(D) Analysis of the kinetics of capacitance
responses shows that Munc13-1 enhances the
RRP (**, P < 0.01), SRP (***, P < 0.001), and
sustained release (P < 0.001) in the response
in CAPS1-expressing chromaffin cells. Error
bars indicate mean ± SEM.
JCB • VOLUME 190 • NUMBER 6 • 2010 1074
Our results show that CAPS2 promotes LDCV priming to the
RRP in chromaffin cells (Fig. 1), as does CAPS1 (Liu et al.,
2008), although there may be subtle functional differences be-
tween the two CAPS isoforms. The selective effect of CAPS
deletion on the RRP is not compatible with a docking defect.
This conclusion is supported by our modeling data (Fig. 8) and
the present (Fig. 5) and previous analyses of LDCV distribu-
tions in CAPS KO chromaffin cells (Speidel et al., 2005; Liu
et al., 2008).
The fact that the CAPS DKO phenotype can be rescued by
expression of open syntaxin is consistent with the notion that
response in which the burst was larger, the RRP was enhanced
relative to the SRP, the UPP was depleted, and the sustained rate
was decreased (Fig. 8 C). Changes in the components are sum-
marized in Fig. 8 D. However, if the UPP (docking) is normal,
as in the case of the CAPS DKO, a loss of sustained release will
require a stronger reduction in priming. This can be achieved by
removing the enhancement of priming because of high calcium
after the flash (Fig. 8 C, dashed line [the calcium-dependent in-
crease in k1 was removed]) to achieve the observed deficit in
sustained release. This change approaches the effect of CAPS
deletion on sustained secretion we observed. If this loss of prim-
ing is preferentially affecting the RRP, the results will model the
effects of CAPS loss.
Figure 8. Numerical simulations of prim-
ing and release indicate that CAPS alters
the Ca2+-dependent enhancement of priming.
(A) Simulated flash responses under conditions
of moderate priming (300 nM free calcium;
black) and high priming (600 nM free calcium;
blue). Changing basal free calcium changes
the basal priming rate, altering pool size with
no effect on the relative sizes of the SRP and
RRP or on sustained release. Enhancing the
priming rate into the RRP at a moderate prim-
ing rate (basal calcium 300 nM; dashed line)
enhances the RRP with no effect on sustained
release. (B) The bar graph shows the RRP, the
SRP, and the sustained rate under these condi-
tions. (C) Decreasing the UPP (red) results in
decreased pool size and sustained release.
This decrease in pool size can be compen-
sated by increasing the priming rate into the
RRP (k2/k-2 ratio > 1), which increases the
burst size but not the sustained rate. Note that
sustained release is reduced proportionally to
the reduction in docking. Sustained release is
dependent on the calcium level after the flash.
Removing the calcium-dependent enhancement
of secretion reduces the sustained component
(dashed line) and approximates the effect of
CAPS deletion on sustained release. (D) The
bar graphs illustrate the pool sizes and sus-
tained rates under these conditions.
Figure 7. The priming effect of Munc13-1
requires the presence of CAPS1 but not of
CAPS2. (A) Responses to flash photolysis in
CAPS1 deletion cells having one functional
CAPS2 allele. Munc13-1–expressing cells
(n = 23), in spite of slightly higher resting calcium
concentrations (top), secreted less than did cells
from CAPS1/ CAPS2+/ littermates (n = 22;
middle). The catecholamine release results are
consistent with the capacitance data (bottom).
(B) The kinetic analysis of flash responses dem-
onstrates that Munc13-1 did not enhance any
component of the responses. (C) Summary of
the results of expressing Munc13-1 in chro-
maffin cells from various CAPS deletion mice.
In CAPS DKO and CAPS1/ CAPS2+/ cells,
Munc13-1 expression failed to enhance re-
sponses, whereas in CAPS1+/ CAPS2/
and wild-type (WT) cells (C57Black6, not
littermates), expression of Munc13-1 (n = 15)
produced a robust enhancement of secretion
versus untreated controls (n = 12). Error bars
indicate mean ± SEM.
CAPS primes to the readily releasable pool • Liu et al.
stabilized open syntaxin could bypass the need for Munc13 and
CAPS. An interaction of CAPS with syntaxin has recently been
reported (James et al., 2009) in a study showing that the fuso-
genic effect of CAPS in a SNARE complex–dependent liposome
fusion assay is inhibited by a soluble syntaxin fragment and that
CAPS binds to syntaxin-containing SNAREs and syntaxin alone.
This function of CAPS appears to result in enhanced SNARE
complex formation, which would be congruent with a model of
combined Munc13/CAPS function.
Although numerous data have firmly established Munc13s
as absolutely essential for priming of SVs in neurons (Augustin
et al., 1999; Richmond et al., 1999), the inability of Munc13-1
to promote priming in the absence of CAPS1 (Figs. 6 and 7) in
conjunction with the lack of a measurable secretion defect in
Munc13-1 KO chromaffin cells (unpublished data) might indicate
that Munc13-1 is not required for LDCV priming in chromaffin
cells. The lack of rescue by the truncated Munc13-1 construct
containing the minimal priming domain would then indicate that
this conclusion is valid for all Munc13 isoforms expressed in
adrenal chromaffin cells. Consequently, priming of LDCVs into
the RRP might proceed constitutively but might be accelerated by
Munc13s and other factors. In this scenario, CAPS would either
promote priming into the RRP directly in a calcium-dependent
manner or function in an alternative calcium-dependent priming
pathway via the SRP to the RRP. A selective function at the RRP
cannot explain CAPS effects on sustained release, as discussed in
Results. Thus, an alternative CAPS-dependent priming pathway
seems more likely. This priming pathway would be calcium depen-
dent because a loss of the calcium-dependent enhancement of
priming is the only manipulation that practically abolishes sustained
release (Fig. 8) and must preferentially prime to the RRP. This
would result in a CAPS-dependent RRP and CAPS-dependent
sustained release, explaining the reduced RRP and sustained re-
lease in the CAPS DKO.
In conclusion, our data demonstrate that CAPS plays a facili-
tatory role in the second step of LDCV priming into the RRP in
adrenal chromaffin cells. Thus, priming in adrenal chromaffin cells
consists of a sequential two-step process.
Materials and methods
CAPS1 and CAPS2 KO mice were described previously (Speidel et al.,
2005; Jockusch et al., 2007). CAPS DKO mice were generated by breed-
ing the CAPS1 mutation into the CAPS2 mutant background. Genotypes
were confirmed by PCR using primers as described previously (Speidel
et al., 2005; Jockusch et al., 2007).
Chromaffin cell preparation and infection
All electrophysiological experiments were performed on mouse chromaffin
cells in primary culture. The cells were prepared as previously described
(Sørensen et al., 2002). In brief, the mice were prepared after hysterec-
tomy on E18/E19. The adrenal glands were rapidly removed and placed
in cold Locke’s solution. The glands were incubated for 20 min in a DME
solution containing 20 U/ml papain (Roche). After the removal of the pa-
pain solution, the glands were washed in an inactivating solution (DME
plus 10% BSA). After 4 min in the inactivating solution, the glands were
triturated and cells were plated on glass coverslips. 30 min later, 3 ml DME
was added to the wells containing the coverslips, and the cells were incu-
bated for up to 4 d at 37°C in DME gassed with 8% CO2. For the CAPS
rescue experiments, isolated chromaffin cells were infected with 50 µl
activated pSFV1-CAPS1-IRES-GFP or pSFV1-CAPS2-IRES-GFP after a previ-
ously described protocol (Ashery et al., 1999, 2000). Open syntaxin was
CAPS and Munc13 proteins operate in the same molecular
priming pathway (Richmond et al., 2001). Similarly, expression
of open syntaxin rescues the locomotion and vesicle docking
defects in C. elegans UNC-31 (CAPS) mutants (Zhou et al.,
2007; Hammarlund et al., 2008), although rescue of the trans-
mitter release defects caused by UNC-31 mutations was not
tested in these studies. The effects of open syntaxin, although
stronger in CAPS DKO cells, were also seen in wild-type cells
(Fig. 4). The blockade of sustained release caused by the pres-
ence of open syntaxin is, to a large degree, likely because of the
observed docking deficit (Fig. 5) that has been previously de-
scribed in open syntaxin mutant mice (Gerber et al., 2008) and
unrelated to CAPS function or dysfunction, as it occurs in wild-
type and CAPS DKO chromaffin cells alike. Whether wild-type
syntaxin overexpression could rescue the CAPS DKO pheno-
type in a similar fashion as open syntaxin could not be tested
because expression of wild-type syntaxin requires the presence
of about equal amounts of Munc18. Without the Munc18-
induced conversion into a form that can be transported, as Rowe
et al. (1999) stated, “syntaxin remains stuck in the Golgi-TGN
area….leading to severe structural and membrane traffic alter-
ations.” Because open syntaxin does not interact with Munc18,
apparently this sorting problem does not exist for this mutant.
As open syntaxin rescues the UNC-13 and UNC-31 mutant
phenotypes in C. elegans (Zhou et al., 2007; Hammarlund et al.,
2008) and the effects on RRP size of CAPS DKO in chromaffin
cells (Fig. 2), we reasoned that CAPSs and Munc13s are function-
ally related and tested whether overexpression of Munc13-1 can
also rescue the priming deficits in the CAPS DKO chromaffin cells.
This was not the case, although overexpression of Munc13-1
strongly enhanced secretion in cells with one functional CAPS1
allele remaining (Fig. 6). At present, we can only speculate about
the molecular basis of the observed isoform specificity. Although
the existence of two splice variants has been reported for CAPS1
(Ann et al., 1997), at least six splice variants exist for CAPS2
(Sadakata et al., 2007). Interestingly, three of these splice variants
have alternative exons in the MHD domain, whereas the remain-
ing three splice variants do not contain the MHD at all. It seems
conceivable that only the isoforms without the MHD are ex-
pressed at the time point of our measurements (i.e., E18/P0).
These splice variants are probably unable to bind syntaxin and,
thus, cannot serve as a platform for Munc13 to act on. Further ex-
periments with specific antibodies for the splice variants will be
necessary to prove or refute this hypothesis.
The observation that exogenous Munc13-1 cannot boost
priming in chromaffin cells in the absence of CAPS1 may indi-
cate a functional interaction between CAPSs and Munc13s. Such
an interaction has been suggested to occur in mouse hippo-
campal neurons (Jockusch et al., 2007), where overexpression of
Munc13-1 failed to reverse deficits in synaptic transmission in
CAPS DKO neurons. Interestingly, in C. elegans neurons, dense-
core vesicle secretion, which is strongly CAPS dependent, is en-
hanced by phorbol esters, but only when CAPS is present (Zhou
et al., 2007). Because Munc13-1 overexpression in chromaffin
cells enhances the sizes of both SRP and RRP (Ashery et al.,
2000), Munc13-1 may act upstream of CAPS by opening
syntaxin, followed by CAPS binding to open syntaxin. Thus, a
JCB • VOLUME 190 • NUMBER 6 • 2010 1076
High pressure freezing
Sapphire discs with cultured cells were dipped into DME with 30% FCS,
transferred into flat specimen carriers, and frozen in a high pressure freezer
(EM PACT2; Leica). Wild-type and CAPS DKO cells were frozen 6 h after
infection. Untreated control cells were frozen at the same time.
Freeze substitution and embedding
Frozen mouse chromaffin cells were fixed in an automatic freeze substitution
apparatus (AFS2; Leica) as described previously (Edelmann et al., 2007). In
brief, cryosubstitution was performed with 2% osmium tetroxide in anhydrous
acetone and 2% H2O. The temperature was increased linearly from 90°C to
70°C over 20 h, from 70°C to 50°C over 20 h, and from 50°C to
10°C over 4 h. After washing with acetone, the cells were embedded in
epon-812 (30% epon/acetone for 10 min at 10°C, 70% epon/acetone for
1 h at 10°C, and pure epon for 1 h at 20°C; Electron Microscopy Sciences).
The temperature was increased linearly from 20 to 60°C over 4 h, and epon
was polymerized at 60°C for 1 d. After polymerization, the carrier and the sap-
phire disc were removed from the epon block. The embedded cells were local-
ized at the surface of the block and could be inspected with a light microscope.
Comparing light micrographs of the infected living DKO and wild-type chro-
maffin cells with the cell patterns sketched from the corresponding block, we
were able to identify the virus-infected cells in the resin block. During trimming of
the block, care was taken so that some of the surrounding cells were left for orien-
tation purposes when viewed at higher resolution in the electron microscope.
Ultrathin (70 nm) sections were cut parallel to the cell monolayer
and collected on pioloform-coated copper grids using an electron micro-
scope (EM UC7; Leica) stained with uranyl acetate and lead citrate and
analyzed with an electron microscope (Tecnai 12 Biotwin; Philips).
Only cells with a visible nucleus and preserved plasma membrane
were analyzed. LDCVs were recognized by their round, dense core. An
outline of both the plasma membrane and the nucleus was generated man-
ually, and vesicles were marked manually and outlined with a circle. The
radius of the vesicle and the shortest distance from its edge to the plasma
membrane were calculated using software written in house (by D. Hof).
We use the model for the chromaffin cell exocytotic pools (Sørensen, 2004)
quantifying the dynamical evolution of the number of vesicles u, s, and r in
the UPP, SRP, and RRP, respectively, by a set of ordinary differential equa-
tions ( u, s, and r denote the time derivative of u, s, and r, respectively):
k0 (k-0) is the forward (backward) docking rate, k1 (k-1) is the forward (back-
ward) priming rate, and k2 (k-2) is the forward (backward) rate for the
transfer SRP↔RRP. fs and fr are the release rates from SRP and RRP, respec-
tively, and dsize is the depot size, which is set to 2,000 vesicles. All rates
are constant for constant Ca2+ stimulus; the standard model assumes that
only the forward priming rate k1 and the release rates fs and fr are Ca2+ de-
pendent. The total number of released vesicles, denoted F, is given by
Ff sf r
. For our base scenario, we use the rates given in Table I and
indicate when we use values deviating from these rates in the simulation.
For low Ca2+ (100 nM), we calculate the stationary depot sizes by
solving the linear equations for u, s, and r which one obtains by setting the
time derivatives to zero ( u = s = r = 0). This yields the depot size shown
in Fig. 8 (B and D). The stationary solution for low Ca2+ is then used as initial
values for u, s, and r, and the time evolution of the depot sizes and released
vesicles (F) at high Ca2+ is simulated by integrating the aforementioned dif-
ferential equations numerically with a fourth-order Runge-Kutta method imple-
mented in a C-code on a standard PC. The sustained rate is then extracted
from the time course of the number of released vesicles F(t) obtained from the
simulation by applying the same fit procedure as used to extract the experi-
mentally determined sustained rate from data for the time course of the mem-
brane capacitance during the first 5 s of flash photolysis of caged Ca.
delivered via a pSFV1-syntaxinL165A/E166A-IRES-GFP. For Munc13-1
overexpression experiments, we used pSFV1–Munc13-1–GFP or pSFV1–
Munc13-1–644-1735-GFP, which have been previously described (Stevens
et al., 2005).
Patch clamp analysis and amperometry
Conventional whole cell recordings were performed with 4–6-MV pipettes
and an EPC-9 patch clamp amplifier together with PULSE software (HEKA).
For measurements from isolated chromaffin cells, the extracellular solution
contained 145 mM NaCl, 2.4 mM KCl, 10 mM Hepes, 4.0 mM MgCl2,
1.0 mM CaCl2, and 10 mM glucose, pH 7.4. The intracellular solution for
isolated cells contained 100 mM Cs-glutamate, 2 mM Mg-ATP, 0.3 mM
Na2-GTP, 40 mM Cs-Hepes, 5 mM nitrophenyl-EGTA (NP-EGTA), 4 mM
CaCl2, 0.4 mM furaptra, and 0.4 mM Fura-4F, pH 7.2. Capacitance mea-
surements were performed using the Lindau-Neher technique implemented
as the ‘sine+dc’ mode of the ‘software lock-in’ extension of PULSE software.
A 1-kHz, 70-mV peak to peak sinusoid stimulus was applied about a DC
holding potential of 70 mV. All experiments were performed at room
temperature. Data are shown as mean ± SEM. We used the Mann-Whitney
U test for comparison of differences between groups. Curve fits were per-
formed using IGOR Pro (Wavemetrics).
Measurements of [Ca2+]i and photolysis of caged Ca2+
[Ca2+]i was measured using a mixture of two indicator dyes, Fura-4F and
furaptra. The dyes were excited with light alternated between 350 and
380 nm using a monochromator-based system, and the fluorescent signal
was measured using a photomultiplier (T.I.L.L. Photonics). To convert the
ratio R of the fluorescent signals at both wavelengths into [Ca2+]i, an in vivo
calibration curve was used (Voets, 2000). To obtain stepwise increases in
[Ca2+]I, short flashes of ultraviolet light from an arc flash lamp (Xenon;
Rapp OptoElectronics) were applied to the whole cell. The monochromator
light was not only used to measure [Ca2+]i but also to maintain calcium
levels for 5 s after the flash (Figs. 1–3, 5, and 6) and allowed us to adjust
[Ca2+]i after a flash or to achieve calcium ramps by photolysing smaller
amounts of NP-EGTA. Trains of light at 350 and 380 nm for ratio measure-
ment of calcium were generated via the monochromator.
Amperometric recordings of catecholamines
Amperometry recordings on isolated chromaffin cells were performed as
previously described (Bruns et al., 2000). Carbon fiber electrodes used for
amperometry were produced as follows. Carbon fibers (5-µm diameter)
were glued to copper cannulae using a conducting carbon paste (Electro-
dag 5513; Bavaria Elektronik) and glued inside a glass pipette. The pi-
pettes were pulled with a conventional puller. The carbon fiber extending
beyond the pulled pipette tip was coated with a cathodal paint by elec-
trolysis (BASF). The assembly was baked for 20 min at 50°C. The junction
between the fiber and glass was sealed with Sylgard (Dow Corning) and
baked again at 50°C. Before use, the carbon fibers were broken off to ex-
pose the tip for recording.
The electrode was connected to the head stage of an EPC7 patch
clamp amplifier (HEKA), and a holding potential of 800 mV was applied
in the voltage clamp mode. After the whole cell configuration was achieved,
the carbon fiber was positioned so that it lightly touched the cell that was
being recorded. Catecholamines contacting the carbon fiber were immedi-
ately oxidized, producing a current on the pipette that was countered by
the patch clamp and allowing recording of catecholamine release as a
measure of the amperometric current.
Preparation of cells for EM
Acutely dissociated chromaffin cells from wild-type and CAPS DKO mice
(E18/P0) were plated on collagen-coated sapphire discs (EM PACT2;
Leica) in 4-well plates. After 2 d in culture, some wells were infected with
50 µl activated pSFV1-syntaxinL165A/E166A-IRES-GFP for 5 h. The in-
fected cells were visualized using a standard laser-scanning confocal
microscope (LSM 710; Carl Zeiss, Inc.) with an excitation wavelength of
488 nm. Low resolution overview images of the whole sapphire discs were
acquired with a 10× objective.
Table I. Base scenario rates used in this study
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We thank Carolin Bick, Katrin Sandmeier, Manuela Schneider, and Reiko
Trautmann (Department of Physiology) and Birgit Leis and Norbert Pütz (Depart-
ment of Anatomy and Cell Biology, University of the Saarland) for expert techni-
This work was supported by the Deutsche Forschungsgemeinschaft
(grants SFB 530, GRK 845, and GRK 1326 to D. Bruns and J. Rettig) and an
intramural funding program (HOMFOR) of the University of the Saarland Medi-
cal School at Homburg.
Submitted: 29 January 2010
Accepted: 17 August 2010
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