Massive Ca-induced Membrane Fusion and Phospholipid Changes Triggered by Reverse Na/Ca Exchange in BHK Fibroblasts

Abstract

Baby hamster kidney (BHK) fibroblasts increase their cell capacitance by 25-100% within 5 s upon activating maximal Ca influx via constitutively expressed cardiac Na/Ca exchangers (NCX1). Free Ca, measured with fluo-5N, transiently exceeds 0.2 mM with total Ca influx amounting to approximately 5 mmol/liter cell volume. Capacitance responses are half-maximal when NCX1 promotes a free cytoplasmic Ca of 0.12 mM (Hill coefficient approximately 2). Capacitance can return to baseline in 1-3 min, and responses can be repeated several times. The membrane tracer, FM 4-64, is taken up during recovery and can be released at a subsequent Ca influx episode. Given recent interest in signaling lipids in membrane fusion, we used green fluorescent protein (GFP) fusions with phosphatidylinositol 4,5-bisphosphate (PI(4,5)P(2)) and diacylglycerol (DAG) binding domains to analyze phospholipid changes in relation to these responses. PI(4,5)P(2) is rapidly cleaved upon activating Ca influx and recovers within 2 min. However, PI(4,5)P(2) depletion by activation of overexpressed hM1 muscarinic receptors causes only little membrane fusion, and subsequent fusion in response to Ca influx remains massive. Two results suggest that DAG may be generated from sources other than PI(4,5)P in these protocols. First, acylglycerols are generated in response to elevated Ca, even when PI(4,5)P(2) is metabolically depleted. Second, DAG-binding C1A-GFP domains, which are brought to the cell surface by exogenous ligands, translocate rapidly back to the cytoplasm in response to Ca influx. Nevertheless, inhibitors of PLCs and cPLA2, PI(4,5)P(2)-binding peptides, and PLD modification by butanol do not block membrane fusion. The cationic agents, FM 4-64 and heptalysine, bind profusely to the extracellular cell surface during membrane fusion. While this binding might reflect phosphatidylserine (PS) "scrambling" between monolayers, it is unaffected by a PS-binding protein, lactadherin, and by polylysine from the cytoplasmic side. Furthermore, the PS indicator, annexin-V, binds only slowly after fusion. Therefore, we suggest that the luminal surfaces of membrane vesicles that fuse to the plasmalemma may be rather anionic. In summary, our results provide no support for any regulatory or modulatory role of phospholipids in Ca-induced membrane fusion in fibroblasts.

Full text

The Journal of General Physiology
ARTICLE
© 2008 Yaradanakul et al.
The Rockefeller University Press $30.00
J. Gen. Physiol. Vol. 132 No. 1 29–50
www.jgp.org/cgi/doi/10.1085/jgp.200709865
29
Correspondence to Donald Hilgemann:
d o n a l d . h i l g e m a n n @ u t s o u t h w e s t e r n . e d u
INTRODUCTION
Eukaryotic cells use precisely orchestrated membrane
fusion and  ssion events to perform multiple cell func-
tions ( Mayer, 2002 ; Bankaitis and Morris, 2003 ; Howell
et al., 2006 ). Fusion events in the secretory pathway are
under the control of protein – protein interactions of
SNAREs and associated proteins that are the subject of
intense ongoing investigation ( Koumandou et al., 2007 ).
Among fusion events, the release of neurotransmitters
from neurons attracts the most attention owing to its fast
triggering by Ca binding to synaptotagmins ( Geppert
et al., 1994 ) and to its fundamental role in neuronal signal-
ing ( Katz, 2003 ). It is less well appreciated that other cell
types use Ca-activated fusion processes for other func-
tions ( Breitbart and Spungin, 1997 ; Gundersen et al.,
2002 ; Steinhardt, 2005 ; Czibener et al., 2006 ), and that
those mechanisms allow analysis of the membrane speci-
 city of fusion processes, the physical basis of membrane
mixing during fusion, and the nature of sensors that ini-
Abbreviations used in this paper: AMP-PNP, adenosine 5  -(  ,  -imido)
triphosphoate; BHK, baby hamster kidney; DAG, diacylglycerol; PI(4,5)P
2
,
phosphatidylinositol 4,5-bisphosphate; PS, phosphatidylserine.
tiate fusion. Even in neurons, asynchronous neurotrans-
mitter release appears to be controlled by Ca sensors that
are different from those used in fusion events closely
coupled to Ca in ux ( Maximov and Sudhof, 2005 ; Sun
et al., 2007 ). Of most interest for this article, many eu-
karyotic cells employ Ca-triggered membrane fusion as
part of a membrane repair reaction initiated by Ca in ux
through cell surface wounds ( Togo et al., 2000 ; Reddy
et al., 2001; Steinhardt, 2005 ). Standard cell culture  bro-
blasts, such as CHO cells, can rapidly expand their sur-
face membranes via membrane fusion in response to a
large increase of cytoplasmic Ca ( Coorssen et al., 1996 ).
The cytoplasmic Ca concentrations needed to trigger
the cell wound response are evidently higher than those
needed for neurotransmitter release ( Schneggenburger
and Neher, 2005 ). In  broblasts, fusion appears to be
initiated in the range of 10 – 30 μ M free Ca ( Coorssen
Massive Ca-induced Membrane Fusion and Phospholipid Changes
Triggered by Reverse Na/Ca Exchange in BHK Fibroblasts
Alp Yaradanakul ,
1
Tzu-Ming Wang ,
1
Vincenzo Lariccia ,
1
Mei-Jung Lin ,
1
Chengcheng Shen ,
1
Xinran Liu ,
2
and Donald W. Hilgemann
1
1
Department of Physiology and
2
Department of Neuroscience, University of Texas Southwestern Medical Center at Dallas,
Dallas, TX 75390
Baby hamster kidney (BHK)  broblasts increase their cell capacitance by 25 – 100% within 5 s upon activating maxi-
mal Ca in ux via constitutively expressed cardiac Na/Ca exchangers (NCX1). Free Ca, measured with  uo-5N,
transiently exceeds 0.2 mM with total Ca in ux amounting to ⵑ 5 mmol/liter cell volume. Capacitance responses
are half-maximal when NCX1 promotes a free cytoplasmic Ca of 0.12 mM (Hill coef cient ≈ 2). Capacitance can
return to baseline in 1–3 min, and responses can be repeated several times. The membrane tracer, FM 4-64, is
taken up during recovery and can be released at a subsequent Ca in ux episode. Given recent interest in signaling
lipids in membrane fusion, we used green  uorescent protein (GFP) fusions with phosphatidylinositol 4,5-bisphos-
phate (PI(4,5)P
2
) and diacylglycerol (DAG) binding domains to analyze phospholipid changes in relation to these
responses. PI(4,5)P
2
is rapidly cleaved upon activating Ca in ux and recovers within 2 min. However, PI(4,5)P
2
de-
pletion by activation of overexpressed hM1 muscarinic receptors causes only little membrane fusion, and subsequent
fusion in response to Ca in ux remains massive. Two results suggest that DAG may be generated from sources
other than PI(4,5)P in these protocols. First, acylglycerols are generated in response to elevated Ca, even when
PI(4,5)P
2
is metabolically depleted. Second, DAG-binding C1A-GFP domains, which are brought to the cell surface
by exogenous ligands, translocate rapidly back to the cytoplasm in response to Ca in ux. Nevertheless, inhibitors
of PLCs and cPLA2, PI(4,5)P
2
-binding peptides, and PLD modi cation by butanol do not block membrane fusion.
The cationic agents, FM 4-64 and heptalysine, bind profusely to the extracellular cell surface during membrane fu-
sion. While this binding might re ect phosphatidylserine (PS) “ scrambling ” between monolayers, it is unaffected
by a PS-binding protein, lactadherin, and by polylysine from the cytoplasmic side. Furthermore, the PS indicator,
annexin-V, binds only slowly after fusion. Therefore, we suggest that the luminal surfaces of membrane vesicles that
fuse to the plasmalemma may be rather anionic. In summary, our results provide no support for any regulatory or
modulatory role of phospholipids in Ca-induced membrane fusion in  broblasts.

30 Membrane Fusion and Phospholipids
DAG ( Gundersen et al., 2002 ). Both DAGs and phorbol
esters can mimic Ca in activating fusion in oocytes, and
activation of the Ca-independent PKC-  is implicated to
be important ( Gundersen et al., 2002 ). Thus, it is pos-
sible that in the oocyte a PLC is the actual Ca sensor
for this type of fusion response. In yeast, a novel PLC
activity has been suggested to play an important role in
homotypic vacuole fusion with DAG serving potentially
as a second messenger that facilitates fusion ( Mayer
et al., 2000 ; Jun et al., 2004 ). Finally, it is noteworthy
that phospholipases appear to have a fusion-promoting
(fusogenic) in uence in multiple membrane fusion pro-
cesses ( Harrison and Roldan, 1990 ; Haslam and Coorssen,
1993 ; Goni and Alonso, 2000 ; Choi et al., 2002 ; Brown
et al., 2003 ; Choi et al., 2006 ). As described in this ar-
ticle, Ca in ux indeed strongly activates PLCs in BHK
cells in the same Ca range and on the same time scale in
which membrane fusion occurs. In addition, we identify
another pronounced membrane change. Extracellular
binding of multiple cationic agents suggests that the ex-
tracellular cell surface rapidly becomes anionic during
and after membrane fusion. We describe a wide range
of experiments to address the possible roles of these
changes in membrane fusion, and the results suggest
that membrane fusion and phospholipid changes are
independent processes that are triggered independently
by large cytoplasmic Ca transients.
MATERIALS AND METHODS
Cell Lines and Transfections
BHK cells expressing NCX1 ( Linck et al., 1998 ) with and without
expression of hM1 receptors were maintained as previously
described ( Yaradanakul et al., 2007 ). BHK cells were grown in
Dulbecco ’ s modi ed minimum essential medium (DMEM) sup-
plemented with 10% FBS with periodic selection by 2.5 mg/ml
methotrexate for NCX1 and continuous selection with 200  g
hygromycin for hM1 receptors. Lipofectamine 2000 was used for
DNA transfections following the manufacturer ’ s protocols. Cells
were harvested with 0.25% trypsin and kept at 35 ° C for 20 min in
culture medium before experimentation.
Patch Clamp
Patch clamp and capacitance recording were performed as de-
scribed previously ( Yaradanakul et al., 2007 ). In brief, giga seals
were established with 4 – 6 μ m inner diameter pipette tips that
were coated with dental wax before cutting and polishing to re-
duce capacitance. The majority of capacitance recordings were
performed with our own software, described in an accompanying
article (Wang and Hilgemann, 2008). Sinusoidal voltage oscilla-
tion (20 mV, 0.5 – 2 kHz) was usually employed when input resis-
tances were < 2 M Ω , and square wave oscillation (20 mV, 0.5 kHz)
was usually employed when input resistances were 1 M Ω or higher.
In the former case, the optimal phase angle was selected digitally
via small changes of the optimal capacitance compensation pa-
rameters. In the latter case, single exponentials were  t to the
falling phase of current transients, and input resistance, cell con-
ductance, and cell capacitance were calculated online. As de-
scribed in the accompanying article, the routines were validated
by simulating typical cell electrical properties with the perturbation
et al., 1996 ). In sea urchin eggs, the threshold Ca con-
centration is ⵑ 3 mM ( Terasaki et al., 1997 ), just a few
fold less than the Ca concentration of sea water. This
Ca sensitivity is low enough so that Ca binding by an-
ionic phospholipids, such as phosphatidylserine (PS)
( Wilschut et al., 1981; Papahadjopoulos et al., 1990 ) and
PIP(4,5)P
2
( Toner et al., 1988 ), might in principle play a
triggering role, as described for pure phospholipid vesi-
cles ( Fraley et al., 1980 ; Wilschut et al., 1981 ). As stressed
by others, however, Mg does not substitute for Ca in the
cell wound response ( Steinhardt et al., 1994 ; Steinhardt,
2005 ), whereas Mg binds nearly as well to anionic mem-
branes as Ca until the vesicles are brought into close
proximity ( Feigenson, 1986, 1989 ). Work in diverse cell
types suggests, overall, that the cell wound response in-
volves both SNAREs and synaptotagmins that are related
to those underlying neurotransmitter release, albeit with
a complication that the wound response can probably
involve multiple membrane types, including endosomes
and lysosomes ( Krause et al., 1994 ; Steinhardt et al.,
1994 ; Bement et al., 1999 ; Detrait et al., 2000 ; Rao et al.,
2004 ; Andrews, 2005 ; Andrews and Chakrabarti, 2005 ).
From the functional effects of synaptotagmin fragments
( Rao et al., 2004 ; Andrews, 2005 ; Andrews and Chakrabarti,
2005 ) and from knockdown studies ( Jaiswal et al., 2004 ),
synaptotagmin VII has been proposed to be the Ca sensor
in the cell wound response. Dysferlins are another group
of Ca-binding proteins with C2 domains that are being
considered for a role in cell membrane repair responses,
in particular in muscle ( Bansal and Campbell, 2004 ;
Washington and Ward, 2006 ).
In this article, we extend the analysis of Ca-induced
membrane fusion in  broblasts with baby hamster kid-
ney (BHK) cells, using cardiac Na/Ca exchangers to ac-
tivate a large Ca in ux and to manipulate cytoplasmic
Ca. For multiple reasons, it appeared important to us to
characterize the function of Ca-dependent phospholi-
pases, especially PLCs in the protocols that evoke mem-
brane fusion in  broblasts. First, phosphatidylinositol
4,5-bisphosphate (PI(4,5)P
2
), the substrate of PLCs,
can bind to and modulate the function of synaptotag-
mins ( Tucker et al., 2003 ; Bai et al., 2004 ) and an as-
sociated protein, CAPS ( Grishanin et al., 2004 ), which
control membrane fusion in the active zones of neu-
rons ( Geppert et al., 1994 ). In addition, there appear
to be requirements for PI(4,5)P
2
in one or more pro-
cesses leading up to membrane fusion ( Milosevic et al.,
2005 ). Second, diacylglycerol (DAG), the lipid product
of PLC activity, can modulate proteins involved in mem-
brane fusion, in particular the munc proteins ( Madison
et al., 2005 ; Speight and Silverman, 2005; Latham and
Meunier, 2006 ), and a DAG-dependent PKC can evi-
dently facilitate the cell wound response ( Togo et al.,
2003 ; Steinhardt, 2005 ). In Xenopus oocytes, the Ca-de-
pendent exocytosis of cortical granules upon fertiliza-
tion appears to be initiated by the activation of PKCs by

Yaradanakul et al. 31
dried by the same procedures used for other mass determina-
tions. They were then dissolved in 50 μ l of a solution containing
0.3 mM tetradecylsulfate and 0.6% Triton X-100. An equal amount
of DAG kinase reaction buffer (2 mM ATP, 2 mM DTT, 100 mM
imidazole, 100 mM NaCl, 25 mM MgCl
2
, 2 mM EGTA, pH 6.6)
was then added to separate samples with and without 10 μ g/ml
DAG kinase (Calbiochem, #266726). The aliquots were reacted to
completion in 30 min at 30 ° C, phospholipids were again extracted
and analyzed as previously described ( Nasuhoglu et al., 2002a ),
and the total mono- and diacylglycerol was determined from the
increase of glycerol phosphate in HPLC analysis of deacylated
phospholipids treated versus not treated with DAG kinase.
Electron Microscopy
The BHK cell preparation for electron microscopy was performed
largely as described for other cell types ( Sugita et al., 2001 ). Cells
were removed from dishes, as described above, and they were
spun down to form loose pellets before  xing in 2% glutaralde-
hyde with 1% sucrose in 0.1 M sodium cacodylate buffer (pH 7.4)
at 37 ° C for 2 h. The pellets were then carefully separated and
rinsed twice in buffer and post- xed in 0.5% OsO
4
, 0.8% K-ferri-
cyanide in the same buffer for 30 min at room temperature. After
rinsing with distilled water, specimen were stained in bloc with
2% aqueous uranyl acetate for 15 min, dehydrated in ethanol and
embedded in Poly/bed 812 for 24 h. Thin sections (65 nm) were
post-stained with uranyl acetate and lead citrate, and they were
viewed with an FEI Tecnai G2 Spirit Biotwin transmission elec-
tron microscope.
RESULTS
BHK Cell Response to Maximal NCX1-mediated Ca Infl ux
Fig. 1 presents a typical electrophysiological recording
from an NCX1-expressing BHK cell during repeated brief
activation of the maximal outward exchange current
when the cytoplasm was weakly Ca buffered. Whole-cell
current, capacitance, and conductance are monitored
as described in Materials and methods with a 20-mV
sinusoidal voltage oscillation at 500 Hz. The cytoplas-
mic solution contains 2 mM ATP, 0.2 mM GTP, 40 mM
Na, and 0.5 mM EGTA with 0.25 mM Ca (0.4 μ M free
Ca), and the extracellular solution is Na free. The out-
ward current is activated by switching from an extracel-
lular solution with 0.5 mM EGTA and no Ca to one with
2 mM Ca (i.e., 1.5 mM free Ca; see Materials and meth-
ods for further details). Panel A shows the entire ex-
perimental records over 6 min in which the exchange
current was activated and deactivated four times. Panel B
shows the records during each activation cycle at higher
time resolution, together with the calculated capacitance
derivative ( dCap/dt ).
The peak outward current is 1.23 nA in the  rst re-
sponse, and the average current for the initial 1-s activa-
tion time is 0.85 nA. We estimated cell volume to be 10
pL, the cell being nearly round under these conditions
with a maximal diameter of 28 μ m. Thus, the initial rate
of Ca in ux, assuming 3Na/1Ca exchange, corresponds
to 1.2 mmol per liter cell volume per second, and the
total Ca in ux is estimated to be > 1 mmol per liter cell
volume during the 1-s application of extracellular Ca.
patterns employed, whereby cell parameters and parameter
changes could be retrieved with an accuracy of about 99.9%. A
few recordings employed phase-lock ampli ers, as described pre-
viously ( Yaradanakul et al., 2007 ), and we detected no qualitative
or quantitative differences between capactiance recordings per-
formed with the two methods.
For experiments without imaging, the voltage-clamped cells
were moved rapidly between four parallel solution streams main-
tained at 35 ° C. For experiments with simultaneous confocal
imaging, a commercial recording chamber (RC-26; Warner In-
struments) was employed in conjunction with a Nikon TE2000-U
microscope, and for extracellular solution changes four solution
lines with on/off switches were merged to a single temperature-
controlled outlet.
Solutions, Chemicals, and Constructs
The standard extracellular solution for maximal outward Na/Ca
exchange current contained (in mM) 110 NMG, 20 TEA-OH,
15 HEPES, 3 MgCl
2
, and 0.5 EGTA, set to pH 7.0 with aspartate,
with activation of current by substituting 2 MgCl
2
for 2 CaCl
2
. The
standard cytoplasmic solution contained (in mM) 80 NMG, 40
NaOH, 20 TEA-OH, 15 HEPES, 0.5 EGTA, 0.25 CaCl
2
, 0.5 MgCl
2
,
1.6 MgATP, 0.4 TRIS-ATP, and 0.2 MgGTP, set to pH 7.0 with as-
partate. For Figs. 3 – 5 , 80 mM NMG was replaced in both solu-
tions with 80 mM LiOH. In Fig. 3 , 40 mM LiOH was replaced with
40 NaOH in the extracellular solution used to activate exchange
current. In the experiments for Figs. 6 and 7 , the cytoplasmic
solution contained (in mM) 60 NaOH, 30 KOH, 30 TEA-OH, 0.5
MgCl
2
, 0.5 EGTA, 0.25 CaCl
2
, 30 HEPES and set to pH 6.9 with
aspartate. For experiments with inward exchange current in Fig.
13 C , the extracellular solution contained (in mM) 120 NaOH
(or LiOH), 1.5 EGTA, 26 TEA-OH, 20 HEPES, and 1 MgCl
2
, set
to pH 7.0 with aspartic acid. The cytoplasmic solution contained
(in mM) 120 LiOH, 20 TEA-OH, 1 MgCl
2
, 3.0 EGTA, CaCl
2
to
achieve the desired free Ca, 15 HEPES, 2 ATP, and 0.2 GTP, set
to pH 6.9 with aspartic acid. Rhodamine-conjugated hepta-lysine
(rhodamine-KKKKKKK- amide; K7-Rhod) was prepared by Mul-
tiple Peptide Systems (NeoMPS, Inc.). Annexin-V Alexa Fluor
488 (#A13201) conjugate was from Invitrogen and was employed
at 1:100 dilution. All other chemicals were from Sigma-Aldrich
and were the highest purity available. The PLC  1PH-GFP fusion
protein construct was a gift of Tobias Meyer (Stanford Univer-
sity, Stanford, CA), and the construct for C1A-GFP fusion pro-
tein ( Oancea et al., 1998 ) was provided by Mark Shapiro (UT
San Antonio).
Imaging
Confocal imaging was as described ( Yaradanakul et al., 2007 ) with
a Nikon TE2000-U microscope and a Nikon 60 × 1.45 NA oil im-
mersion objective. A 40 mW Spectra Physics 163-CO2 laser was
used for 488/514 nm excitation (i.e., for GFP constructs and FM
dye); a 1.5 mW Melles Griot cylindrical HeNe Laser was used for
543.5 nm excitation (i.e., for rhodamine constructs). The time
lapse images were recorded either at 160 × 160 or 256 × 256 reso-
lution with an ⵑ 400-ms total exposure time. The exposure inter-
val was 3 s in all records presented. Lasers were operated at 1%
power with a detector pinhole setting of 60 μ m and an average
linear gain of 7.00. Fluroescence intensity is quanti ed in most
 gures as percent of maximal  uorescence occurring during the
recording (%).
Lipid Analysis
Anionic phospholipid mass measurements were performed as
previously described ( Nasuhoglu et al., 2002a ). The total mono-
and diacylglycerols were determined via a bacterial DAG kinase
that phosphorylates both mono and diglycerides ( Preiss et al.,
1987 ). To do so, phospholipids were  rst extracted, washed, and

32 Membrane Fusion and Phospholipids
exchange current has been completely activated. Sec-
ond, the outward NCX1 current under the conditions
of these experiments (i.e., with 120 mM extracellular
NMG and no other extracellular monovalent cations) is
nearly voltage independent ( Matsuoka and Hilgemann,
1992 ; Matsuoka and Hilgemann, 1994 ) and therefore is
not expected to contribute substantial conductance
changes. Third, the slow rise of conductance correlates
roughly in time with the rate of rise of capacitance (i.e.,
dCap/dt ), and the peaks of these signals occur at approx-
imately the same time, as denoted by a second dotted
vertical line in each response.
The peak exchange current typically decreased by at
least 40% from one Ca pulse to the next with Ca pulse
durations of 1 – 5 s. Also evident in these records, the rise
of capacitance typically occurred with a longer delay at the
second and later responses when the exchange current
was decreased. Nevertheless, the second increase of capac-
itance was typically larger and occurred with a higher max-
imal rate than during the  rst response. This is consistent
with reports that the cell wound response shows facili-
tation when induced multiple times ( Togo et al., 2003 ).
Membrane capacitance starts to rise and achieves a
maximal rate of rise at roughly the time point of com-
plete exchange current deactivation. The rise of capaci-
tance occurred typically with a delay of at least 0.1 s.
The rise was often preceded by a decline, amounting to
a few percent of the  nal rise (see negative derivative
signal). The initial cell capacitance was 30.6 pF in this
experiment, the maximal rate of rise of capacitance was
7.5 pF/s, and the total increase of capacitance at the
 rst Ca pulse amounted to 4 pF (i.e., 13% of the initial
cell capacitance). The cell resistance was 0.3 G Ω (3 nS)
before activating exchange current.
During Ca in ux, the cell conductance increases by
20 nS within 700 ms and then decreases partially toward
baseline. The irreversible component of this increase
probably re ects a decrease of seal resistance. The tran-
sient part of the response may in part re ect exchange
current activity, but several arguments suggest that it
re ects mostly conductive properties of fusion pores
(Lindau and Alvarez de Toledo, 2003; Neef et al., 2007 ).
First, the transient conductance changes do not track
exchange current. Conductance rather rises slowly after
Figure 1. Typical records of membrane capaci-
tance, conductance, and current from BHK cells
consitutively expressing the cardiac Na/Ca ex-
changer (NCX1). Cell parameters are monitored
via 20-mV sinusoidal membrane voltage pertur-
bation at 0.5 kHz. Input resistance ≈ 1 M Ω . Cy-
toplasmic solution contains 40 mM Na and 0.5
mM EGTA with 0.25 mM Ca (free Ca, 0.4 μ M).
Exchange current was activated four times for
increasing durations of 1 – 6 s by applying and
removing 2 mM extracellular Ca, as indicated
below the records. (A) Complete signal records.
(B) Expanded signal records for the time pe-
riods during and immediately after activation of
exchange current. dCap/dt is the  rst derivative
of capacitance. Two vertical dotted lines in each
activation cycle mark the time when current was
activated and the time at which the maximal rate
of rise of capacitance occurred. Note that the rise
of capacitance is preceded by a small negative ca-
pacitance phase. The time from current activation
to peak rate of rise of capacitance increases as the
peak exchange current decreases in this sequence.

Yaradanakul et al. 33
Current and Ca Dependencies of Capacitance Responses
Using the protocol described in Fig. 1 , we proceeded to
analyze the exchange current and Ca dependencies of
the capacitance responses. As evident in Fig. 1 B , capaci-
tance increases smoothly during the same time period in
which exchange current is deactivated by removing ex-
tracellular Ca. This observation indicates that Na/Ca ex-
changers do not generate a local Ca signal that is critical
for this type of membrane fusion. To determine the mean
free Ca concentrations occurring in cells, we employed
the low af nity Ca indicator dye  uo-5N (Invitrogen) at
a concentration of 3 μ M in the pipette solution. Fig. 2 A
shows a typical  uorescence record obtained during
NCX1 activation under the same conditions as Fig. 1
(0.5 mM EGTA with 0.25 mM total Ca, 0.4 μ M free Ca).
After recording background  uorescence for 1.5 min,
the cell was exposed to 2 mM extracellular Ca for 3 s, as
is usual with substitution for 2 mM Mg,  uorescence sig-
nals were allowed to dissipate for 30 s. Then, the same
cytoplasmic solution was perfused into the cell with the
dye saturated by 1 mM free Ca (i.e., 1.5 mM total Ca with
0.5 mM EGTA) to determine the maximal Ca response of
the dye in the cell. Peak free Ca induced by Ca in ux was
then estimated assuming a K
d
for Ca of 90 μ M with free
Ca calculated as K
d
/( F
max
/ F  1), where F
max
is the cell  uo-
rescence with Ca-saturated  uorophore and F is the peak
 uorescence obtained in response to activating Ca in ux.
Fig. 2 B shows our calculation of total Ca in ux (i.e., the
integral of the exchange current related to the estimated
As in most experiments, the same magnitude of capaci-
tance increase was achieved at the third and fourth Ca
pulses when the Ca pulse period was increased to com-
pensate for the decreased exchange current.
We stress that both the magnitudes of capacitance in-
crease and the extent to which capacitance signals re-
versed to baseline were somewhat variable and showed
statistically highly signi cant dependence on cell batch
and growth conditions. The average increase of capaci-
tance was found to vary by up to  vefold between cell
batches with similar basal capacitance and exchange
current density. For example, we veri ed in two datasets
that removal of serum from cells for 24 h caused a
highly signi cant decrease of the capacitance response
by > 50%, while the exchange current density was un-
changed. In ⵑ 50% of cell batches, cell capacitance did
not reverse completely, as occurs in Fig. 1 , and in ⵑ 25%
of cases the cell capacitance decreased to values lower
than the initial cell capacitance before activating Ca in-
 ux. Rarely, reversal amounted to only a small fraction
of the initial capacitance increase, and capacitance in-
crements were then very small at the second to fourth
Ca pulses. Finally, we mention that capacitance re-
sponses in CHO and HEK293 cells were typically smaller
than those of BHK cells when equivalent NCX1 ex-
pression was achieved by transient or stable expression
methods. Responses in CHO cells seldom exceeded
40% of basal cell capacitance, and responses in HEK293
cells were usually close to 15%.
Figure 2. Measurement of cytoplasmic free Ca changes during activation of outward Na/Ca exchange currents in BHK cells. The pi-
pette solution contains 0.5 mM EGTA and 0.25 mM Ca (i.e., 0.4 μ M free Ca) with 3 μ M Fluo 5N (K
d
, 90 μ M). Here, and in all  gures,
Ca concentrations given indicate the free Ca concentration. (A) Typical  uorescence and current records. When 2 mM Ca is applied
for 4 s, current rises rapidly to a peak of 175 pA and decays to baseline within < 1 s when Ca is removed. Peak  uorescence occurs at 3 s
and begins to decay rapidly before current is deactivated. Upon perfusion of the pipette with 1 mM additional Ca,  uorescence rises to
a steady “ maximal ” level with a time constant of ⵑ 20 s. (B) Free and total Ca are calculated from the  uorescence and current records
as described in the text. Peak free Ca is estimated to be 0.20 mM at a time point when total Ca in ux amounts to 3.0 mmol/liter cell
volume. (C) Peak free and total Ca in ux values obtained 14 control cells and 5 cells perfused with 2 μ M thapsigargin and 2 mM AMP-
PNP to block Ca pumping. Results are not signi cantly different.

34 Membrane Fusion and Phospholipids
From experiments in which we employed BHK cell
lines with different NCX1 current densities, our impres-
sion was that the rate and extent of capacitance increase
were nearly proportional to the current density. As de-
scribed in Fig. 3 , we analyzed quantitatively the relation-
ships between peak current density and capacitance
changes by varying extracellular Ca. By performing these
experiments in the presence of 40 mM Na on both
membrane sides, at 0 mV, the maximal free cytoplasmic
Ca is thermodynamically limited to not exceed the free
Ca applied to the extracellular side. Fig. 3 (A and B)
shows representative results for applying 100 and 10 μ M
free Ca together with 40 mM extracellular Na. We quan-
ti ed the rate of rise of capacitance as the fractional in-
crease of cell capacitance per second ( dCap / dt / Cap , i.e.,
dCap / dt divided by the basal cell capacitance). Fig. 3 C
presents the dependence of capacitance change on
the extracellular Ca concentrations employed (i.e., the
maximal possible cytoplasmic free Ca), and Fig. 3 D
presents the dependence on the peak current density.
The maximal fractional rate of rise of capacitance is
ⵑ 0.3 per second (i.e., 30% per second). In both plots,
the rate of rise of capacitance shows a sigmoidal depen-
dence on current with a Hill coef cient of ⵑ 2. Half-sat-
uration occurs at 140 μ M free extracellular Ca and 8
pA/pF, respectively. From these results and knowledge
of the peak Ca/peak current density relationship from
Fig. 2 , we conclude that a free Ca concentration > 10 μ M
is required to achieve 10% of the maximal fusion rate,
and that a concentration > 100 μ M is required to achieve
the half-maximal fusion rate. We mention that the ca-
pacitance delay, de ned as the time from activation
of current to the time at which capacitance crossed
the zero line (see Fig. 3 B), decreased with increasing
cell volume) and the free Ca from the calibration just
described. In this experiment, peak free Ca is 170 μ M
and total Ca in ux is estimated to amount to 6 mmol/
liter cell volume. At the peak of the Ca transient, the ra-
tio of total to free Ca is estimated to be 25. As shown in
Fig. 2 C , the average peak free Ca for 14 cells was 206
μ M, and the average total Ca in ux amounted to 5,183
 mol/liter cell volume. A major concern about this
dye calibration method is whether the maximal  uo-
rescence really re ects  uorescence of the saturated dye
or whether free Ca might still be signi cantly controlled
by cellular Ca pumps. Therefore, we performed a series
of experiments in which the cytoplasmic solution con-
tained thapsigargin (2 μ M), as well as adenosine 5  -(  ,  -
imido) triphosphoate (AMP-PNP) (2 mM), to ensure that
Ca pumps were inactive. As evidence of ef cacious pump
inhibition, the time from peak to 50% decay of the Ca
transients was increased by 2.4-fold in the treated cells
(unpublished data). As shown in Fig. 2 C , inhibition of
Ca pumps did not affect exchange currents (i.e., the
calculated total Ca in ux is unchanged), and the calcu-
lated peak free Ca was unchanged. Accordingly, we con-
clude that the free Ca is accurately determined by the
protocol of Fig. 2 A .
Figure 3. Dependence of BHK cell capacitance responses on the
extracellular Ca concentration and exchange current density (0
mV, 35 ° C). (A) Typical current, capacitance, and conductance
records during application of 0.1 mM free Ca together with 40
mM Na, thereby limiting the maximal free cytoplasmic Ca con-
centration to 0.1 mM. (B) Typical current, capacitance, and con-
ductance records during application of 10 μ M free Ca together
with 40 mM Na, thereby limiting the maximal free cytoplasmic
Ca concentration to 10 μ M. (C) Maximal rate of capacitance in-
crease as fractional increase of total cell capacitance per second
(dCap/dt/Cap) in dependence on the applied extracellular free
Ca. (D) Maximal fractional rate of capacitance rise in depen-
dence on peak exchange current density.
Figure 4. Capacitance and current record of a BHK cell express-
ing NCX1 in perforated patch whole cell con guration. Same
solutions as previous  gures with 40 μ M  -escin in the pipette
solution and capacitance recording via square wave perturbation.
Error bars represent the results of 13 similar experiments.

Yaradanakul et al. 35
posite results for 13 experiments. In brief, capacitance
changes were similar in magnitude to results presented
above, the average increase being 28% in this series.
The rates of capacitance increase and subsequent rever-
sal are both somewhat lower than averages for several
batches of BHK cells with normal whole-cell patch
clamp. Peak exchange currents decreased from one Ca
in ux episode to the next roughly to the same extent as
in the routine whole-cell results.
Metabolic Dependence of Membrane Fusion
We next address the question of whether metabolic pro-
cesses signi cantly control and/or modulate this type of
membrane fusion, ATP-dependent processes in gen-
eral, and protein phosphorylation in particular. In the
experiments described in Fig. 5 , ATP and GTP were
both replaced in the cytoplasmic solution with a nonhy-
drolyzable ATP analogue, AMP-PNP (2 mM). Fig. 5 A
shows a typical fusion response after cytoplasmic perfu-
sion with AMP-PNP for 4 min. The capacitance response
is still large, and the maximal rate of capacitance change,
9 pF/s (14% per second), occurs as usual ⵑ 0.5 s after
starting Ca in ux. The  nal increase of capacitance is
just 40% of the initial cell capacitance. Thus, long-term
ATP removal clearly does not block cell capacitance re-
sponses. Nevertheless, analysis of multiple responses
reveals that the extent of the capacitance responses is
signi cantly blunted by the ATP analogue. As shown in
the  rst two data bars of Fig. 5 B , the response magni-
tudes were reduced by 55%. To test whether this de-
crease might require a protein phosphatase activity, we
perfused cells from the same cell batch with 5 μ M cyclo-
sporin to test for a role of calcineurin and with 4 mM
 uoride and 9 mM pyrophosphate to inhibit phospha-
tases nonselectively. As shown further in Fig. 5 B , the ca-
pacitance responses were not signi cantly restored in
AMP-PNP solutions by these interventions. The peak
NCX1 current was signi cantly increased by  uoride
and pyrophosphate. We stress however that Mg will be
current density in an almost linear fashion (unpub-
lished data).
Other Experimental Factors
Next, we tested how the magnitude and speed of re-
sponses were in uenced by the speed of Ca transients
and several other potentially important experimental
factors. Ca in ux rates achieved with the Ca ionophore,
A23187, were in general not adequate to cause large ca-
pacitance responses. To achieve the highest possible Ca
in ux rates with A23187, cells were incubated with high
concentrations of ionophore (10 μ M) in the absence of
extracellular Ca, and then 1 mM extracellular Ca was
applied as in experiments with NCX1. Capacitance re-
sponses remained rather small and occurred with much
larger time constants (20 – 50 s) than for activation of
NCX1 outward current. Similarly, capacitance responses
to pipette perfusion of cytoplasmic solutions with high
Ca (e.g., 1.5 mM Ca with 0.5 mM EGTA, as in Fig. 2 A )
were much slower than for exchange current activation,
and responses were often blunted or negative if pipette
perfusion was not rapid. Another factor that raised con-
cern in early experiments was our use of large tip diam-
eters to facilitate voltage clamp and pipette perfusion of
the cytoplasm. In this regard, we can report that results
were very similar when conventional patch pipettes
were employed. As shown in Fig. 4 , we also checked
whether results might be dependent on dialysis of the
cytoplasm. To do so, we performed similar experiments
with a perforated patch clamp technique, speci cally
the  -escin method ( Fan and Palade, 1998 ) that allows
fast cell dialysis with a molecular weight cutoff of 5 – 10
kD. Using a concentration of 40 μ M detergent, perfused
into the pipette tip with the standard cytoplasmic solu-
tion, we monitored the establishment of voltage clamp
using square wave voltage oscillation. In most cells, ac-
cess resistances decreased to < 5 M Ω , and square wave
voltage oscillation allowed accurate capacitance analy-
sis. Fig. 4 shows a typical experiment together with com-
Figure 5. Effects of nonhydrolyzable
ATP analogue and protein phospha-
tase inhibitors on the BHK cell capaci-
tance response and exchange current
rundown. (A) Capacitance, conduc-
tance, and current records from a BHK
cell perfused with 2 mM AMP-PNP for
4 min before activating exchange cur-
rent. (B) Capacitance changes and
peak exchange current densities for
control cells, cells perfused with 2 mM
AMP-PNP, cells perfused with 2 mM
AMP-PNP, and with 5 μ M cyclosporin
for 4 min before activating exchange
current, and cells perfused with AMP-
PNP with 9 mM pyrophosphate and 4
mM  uoride.

36 Membrane Fusion and Phospholipids
panel I, shows typical  uorescence changes of BHK cells
expressing PH domains in response to maximal M1 re-
ceptor activation with 0.3 mM carbachol (Fig. 6 A), in re-
sponse to maximal outward exchange current (B), and
then sequentially to Ca and carbachol in the same cell
(C). The  uorescence curves give the average cytoplas-
mic  uorescence of the voltage-clamped BHK cells, per-
fused internally with 0.5 mM EGTA and 60 mM Na at
35 ° C with corresponding cell micrographs given above
the records. Note that cells are nearly round under the
conditions of the experiments, and that some cells form
large blebs after opening with large-diameter pipette tips,
even before activating receptors or Ca in ux.
In response to carbachol ( Fig. 6 A ), cytoplasmic  uo-
rescence rises by fourfold within 20 s as PLC  1PH-GFP
domains are lost from the cell surface. Upon removal
of carbachol, the amount of cytoplasmic  uroescence
begins to decrease within 5 s and returns to baseline in
a nearly exponential fashion with a best- t time con-
stant of 32 s. In response to activation of outward NCX1
current for 12 s (Fig. 6 B), PLC  1PH-GFP domain re-
sponses of similar magnitude and kinetics are recorded.
As shown in Fig. 6 C , the magnitudes of responses to
chelated in this protocol. From our own experience as
well as the literature ( DiPolo et al., 2000 ; Wei et al.,
2002 ), cytoplasmic Mg chelation signi cantly stimulates
the reverse exchange operation.
Ca-activated PI(4,5)P
2
Cleavage
As outlined in the Introduction, recent studies suggest
that phosphoinositide metabolism can modulate some
membrane fusion processes. From previous studies
( Rhee, 2001 ; Nasuhoglu et al., 2002b ) the free Ca con-
centrations occurring in our experiments can be ex-
pected to cause activation of PLCs and PI(4,5)P
2
depletion. Thus, it seemed important to analyze in de-
tail PI(4,5)P
2
breakdown during these protocols in BHK
cells in relation to the capacitance responses. Figs. 6 – 9
present the major electrophysiological, optical, and bio-
chemical results from these experiments.
First, we compared PI(4,5)P
2
breakdown in BHK cells
in response to activation of a G protein – coupled receptor
(GPCR) that activated PLCs and a rise of cytoplasmic Ca
via NCX1. To do so, we expressed PLC  1PH-GFP fusion
protein together with M1 receptors ( Selyanko et al.,
2000 ) in BHK cells expressing NCX1 constitutively. Fig. 6,
Figure 6. PLC activation
monitored via PLC  1PH-GFP
(panel I) and C1-GFP (panel
II) protein fusions in BHK
cells voltage clamped to 0 mV
via large-diameter pipette
tips. The continuous rec-
ords give the average  uo-
rescence in a central region
of the cytoplasm. As evident
in micrographs, shown above
the cells, the large-diameter
tips promote bleb formation.
(Panel I, A) The PLC  1PH-
GFP domain, initially local-
ized mostly to the surface
membrane, rapidly translo-
cates to the cytoplasm during
carbachol (0.3 mM) applica-
tion for 20 s. When agonist
is removed, PH domains re-
turn to the cell membrane
with a time constant of ⵑ 32
s. (B) Similar PH domain re-
sponses recorded when cells
are loaded with high Na (40
mM) and Ca in ux via NCX1
is activated for 12 s by ap-
plying and removing 2 mM
extracellular Ca. (C) Com-
parison of PH domain signals
in a cell activated  rst by Ca
in ux, then by carbachol, and  nally again by Ca. (Panel II, D) The C1 domain is initially cytoplasmic and then rapidly translocates to
the membrane during carbachol (0.3 mM) application for 20 s. (E) Slower and smaller C1 domain response, typically obtained during
activation of outward NCX1 current. (F) Typical C1 domain cytosolic  uorescence changes when carbachol (0.3 mM) is applied continu-
ously and Ca (2 mM) is then applied for 15 s together with carbachol, followed by washout. C1-GFP domains dissociate rapidly from the
plasma membrane during Ca in ux and accumulate diffusely in the cytoplasm.

Yaradanakul et al. 37
inward exchange currents had no evident effect on the
PH domain distribution in cells (unpublished data).
Thus, we conclude that free Ca in the range of 10 μ M is
needed to signi cantly activate PLCs in the absence of
GPCR activation.
Given that high free Ca can strongly activate PI(4,5)P
2
breakdown in BHK cells, it may be expected that PLC
activity will contribute to NCX1 current rundown in the
protocols with massive Ca in ux. However, as just
described, the PH domains redistribute reversibly in
response to both carbachol and Ca changes on a rather
fast time scale. By comparison, the suppression of out-
ward exchange current in response to Ca in ux epi-
sode is very long-lived and often does not reverse at all
( Figs. 1 and 4 ). Thus, PI(4,5)P
2
depletion cannot be
the sole or even the major mechanism of long-term ex-
change current inactivation and rundown in the experi-
ments described.
Dual C1A Domain Responses to a Rise of Cytoplasmic Ca
In panel II of Fig. 6 , and in Fig. 7 , we describe responses
of the DAG-binding C1A-GFP fusion proteins ( Oancea
et al., 1998 ). Fig. 6 (D and E) shows the usual signals
observed for carbachol and NCX1 activation, respectively
activation of NCX1 were typically at least as large as the
responses to carbachol in the same cell, and responses
could be repeated multiple times with only little loss of
signal magnitude.
From these results, it is clear that Ca in ux can acti-
vate large PLC responses in BHK cells. To determine
more accurately the Ca range over which PLCs become
activated, we analyzed PH domain distributions in de-
pendence on the inward exchange current in the pres-
ence of different free Ca concentrations in the pipette
solution. From previous experiments, it is clear that free
cytoplasmic Ca decreases signi cantly during activation
of inward current by extracellular Na, even in the pres-
ence of several millimolar EGTA buffering ( Yaradanakul
et al., 2007 ). Since the free cytoplasmic Ca cannot ex-
ceed the free Ca of the patch pipette solution, protocols
with inward current can give a clear indication of the
minimum Ca required to activate PLCs in the absence
of receptor activation. To summarize brie y results from
> 20 experiments, removal of extracellular Na to deacti-
vate inward current caused clear PLC activation when
cells were perfused with solutions containing free Ca
of 20 μ M or higher. When cells were opened with free
Ca of 8 μ M or less, the activation and deactivation of
Figure 7. Translocation of C1-GFP domains to the surface membrane of BHK cells by short-chain DAG8 (A), phorbol ester (B), and
carbachol (C), followed by redistribution to the cytoplasm in all three cases by activation of Ca in ux (i.e., outward NCX1 current). (A)
Application of 20 μ M C8 diacylglycerol brings the majority of C1 domain to the surface membrane with a time constant of ⵑ 45 s. Upon
activating reverse exchange current, domains return quantitatively to the cytoplasm with a time constant of ⵑ 35 s. (B) Application of
0.2 μ M phorbol ester (PMA) brings C1 domains to the cell surface with a time constant of ⵑ 30 s. During activation of exchange current,
C1 domains reach a substantially higher concentration in the cytoplasm than in the basal state with a time constant of ⵑ 30 s, and C1 do-
mains begin to move back to membrane immediately upon terminating Ca in ux. (C) C1-GFP domain records together with cell capaci-
tance and membrane current in a BHK cell expressing NCX1 and hM1 receptors. With application of carbachol (0.3 mM) cytoplasmic
 uorescence decreases by 60% with a time constant of ⵑ 20 s, and cell capacitance increases modestly. Upon activation of Ca in ux, cell
capacitance increases by ⵑ 8 pF (30%) in less than 10 s, and C1 domains accumulate again in the cytoplasm with time constant of ⵑ 10 s.
Upon deactivating exchange current, C1-GFP domains move partially back to the surface membrane.

38 Membrane Fusion and Phospholipids
uniform distribution develops with a time constant of
just 15 s.
As summarized in Fig. 7 , we next tested how Ca in ux
affects the C1 domain distribution when domains are
brought to the surface membrane by applying exoge-
nous C1 domain-binding reagents, speci cally short
chain diocylglycerol (DAG8, 20 μ M) and phorbol ester
(PMA, 0.2 μ M). As shown in Fig. 7 (A and B) , these
agents cause a pronounced loss of C1-GFP domain
from the cytoplasm and accumulation at the cell sur-
face (a to b). Subsequent activation of NCX1 outward
current by a 20 – 30-s application of extracellular Ca re-
sults in complete translocation of C1 domains back to
the cytoplasm. In the example shown for PMA (Fig. 7 B)
it appears possible that DAG-generating mechanisms
are signi cantly active in the absence of agonist, as there
is a substantial overshoot of C1 domain in the cytoplasm
upon activating Ca in ux. The cytoplasmic redistribu-
tion of C1 domains reverses over 2 min when the Ca in-
 ux via NCX1 is stopped by removal of extracellular Ca.
Fig. 7 C presents in more detail C1 domain responses
(i.e., cytosolic C1 domain  uorescence) together with
membrane capacitance and current during continuous
activation of M1 receptors with a short activation of out-
ward exchange current by extracellular Ca. Carbachol
is applied for 100 s before activation of NCX1. During
this time, ⵑ 60% of C1 domains move out of the cyto-
plasm with a time constant of ⵑ 20 s, and cell capaci-
tance increases by ⵑ 1 pF. Activation of NCX1 for 20 s
causes an 8-pF increase of capacitance, corresponding
to a 30% increase of cell capacitance. Within this same
time frame, 80% of C1 domains return to the cytoplasm
and exchange current decreases by 75%. As exchange
current decreases and is deactivated by removing extra-
cellular Ca, C1 domains are again lost from the cyto-
plasm partially. However, the C1 domain population in
the cytoplasm remains elevated substantially from the
nadir achieved during the initial application of carba-
chol. We conclude from these experiments that the dis-
tribution of C1 domains between the cell membrane
and cytoplasm is subject to both rapid, short-term effects
of raising Ca and longer-term in uences.
Phospholipid Changes in Response to Changes of
Cytoplasmic Ca
We can suggest four mechanisms by which high cyto-
plasmic Ca may cause translocation of C1 domains from
the cell surface to the cytoplasm, and we attempted to
eliminate these one by one in appropriate experiments.
First, DAG metabolism may be Ca dependent, a major
mechanism being that some DAG kinases are activated
by Ca ( Jiang et al., 2000a ; Luo et al., 2004 ; Topham,
2006 ). Second, C1 domains interact not only with DAG
but also with negatively charged phospholipids, espe-
cially PS ( Bittova et al., 2001 ; Ho et al., 2001 ), which can
bind Ca ( Wilschut et al., 1981 ; Papahadjopoulos et al.,
( > 20 observations), and F illustrates a major complexity
encountered. As shown in D and E, the C1 domain
appears almost uniformly distributed throughout the
cytoplasm in unstimulated, voltage-clamped BHK cells.
When M1 receptors are overexpressed, as in all three
cells depicted, application of carbachol (0.3 mM) re-
sults in the association of these domains with the mem-
brane within 20 s. Upon removal of carbachol, the
domains reequilibrate into the cytoplasm with some-
what larger time constants (90 s) than those just de-
scribed for PH domains. Thus, metabolism of DAG by
lipases and kinases may be somewhat slower than resyn-
thesis of PI(4,5)P
2
(i.e., phosphorylation of PI and PIP).
As apparent in Fig. 6 E , activation of Ca in ux by NCX1
causes a qualitatively similar response to that of carba-
chol, as anticipated, but the extent of the responses and
the rate of rise of the responses were consistently less
than for carbachol. As shown in Fig. 6 F , the blunted re-
sponse of C1A domains to Ca elevation re ects a sec-
ond, strong opposing in uence of Ca on the distribution
of C1 domains. This opposing in uence is vividly re-
vealed when the C1 domains are  rst brought to the
membrane by carbachol, and then extracellular Ca is
applied to induce a rise of cytoplasmic Ca in the contin-
ued presence of carbachol. As shown in Fig. 6 F, Ca
in ux in the presence of carbachol results in a rapid re-
turn of C1 domain to the cytoplasm whereby a nearly
Figure 8. Phosphoinositide and phosphatidate changes in BHK
cells in response to Ca in ux via NCX1. The bar graphs represent
the relative phospholipid concentrations, as percent of total an-
ionic phospholipid, for cells under control conditions (control),
cells treated with 25 μ M nystatin in the absence of Ca for 10 min
(nystatin  Ca), and nystatin-treated cells that were exposed to 4
mM extracellular Ca for 1 min (nystatin + Ca).

Yaradanakul et al. 39
these agents ( Jiang et al., 2000b ), and we report in this
same connection that a substantial rise of phosphatidic
acid that occurs in parallel with loss of phosphatidylino-
sitol during M1 receptor activation ( Li et al., 2005 ) was
unaffected by the DAG kinase inhibitor, R59022, at a
concentration of 20 μ M. Third, we performed the pro-
tocols of Fig. 7 with 2 mM AMP-PNP and no ATP in the
pipette solution, and the C1 domain still translocated
rapidly to the cytoplasm when Ca in ux was activated
(two observations). On this basis, the activation of DAG
kinase activity at the cytoplasmic cell surface by a rise of
cytoplasmic Ca seems eliminated as a cause of the C1
domain translocation.
As described in Figs. 8 and 9 , we measured phospho-
lipid changes in response to increasing cytoplasmic Ca in
two different approaches using BHK cells. In the  rst ap-
proach, we used NCX1 to mediate a large Ca in ux. To
do so, the ionophore, nystatin (25 μ M), was applied to
load cells with Na in Ca-free PBS (130 mM NaCl, 20 mM
HEPES, 15 mM glucose, pH 7.4) for 10 min. Thereafter,
4 mM Ca was added to the extracellular medium for
1 min without addition of Na (130 mM TEA-Cl). Anionic
phospholipids were then determined as described, and
the relative mass of the four phospholipids of most inter-
est (PIP, PI(4,5)P
2
, PI, and PA) are shown in Fig. 8 . Cali-
brations are given as percent of total anionic phospholipid,
whereby the majority of anionic phospholipid is PS
and cardiolipin. The nystatin treatment resulted in a
30% reduction of PI(4,5)P
2
in the absence of Ca, and
there was little or no change of other phospholipids. The
Ca addition caused PIP to fall by 70% and PI(4,5)P
2
to
fall by 50%, as expected for PLC activation. During the
1990 ) and thereby might cause release of C1 domains
from the membrane. Third, DAG might be generated by
PLCs on internal membranes by multiple Ca-dependent
mechanisms, thereby favoring translocation of C1 do-
mains to the cytoplasm. And fourth, DAG might trans-
locate to the outside of the cell during membrane
fusion, assuming that phospholipid mixing between bi-
layers occurs during fusion, thereby decreasing DAG on
the inner membrane lea ette and releasing C1 domain
back to the cytoplasm. In this light, we performed both
 uorescence and biochemical studies to determine how
a rise of cytoplasmic Ca affects phosphoinositides, DAG,
and phosphatidic acid, and how those effects depend
on cytoplasmic ATP.
Three types of experiments address whether Ca-
activated DAG kinase activity might play a role in the C1
domains translocation to the cytoplasm in response to
Ca in ux. The  rst type of evidence is represented by
the PMA experiment in Fig. 7 . PMA is not phosphory-
lated by DAG kinases, and yet activation of Ca in ux
causes C1 domains to move to the cytoplasm after being
brought to the cell surface by PMA. In this response,
DAG kinase activity at the cell membrane cannot be the
cause of the C1 domain movement. Second, we tested
for inhibition of C1 translocation by the two available
DAG kinase inhibitors at concentrations recommended
for strong inhibition (DAG kinase inhibitors I and II
[Calbiochem]; R59022 and R59949, each at 10 μ M). We
found no inhibition of the C1 translocation by either of
these agents (unpublished data; two observations each)
upon activating Ca in ux. These results alone are not
conclusive because not all DAG kinases are inhibited by
Figure 9. Phosphoinositide, phospha-
tidate, and total mono/diacylglycerol
(M/DAG) levels of permeabilized BHK
cells in dependence on ATP, AMP-PNP,
and Ca. (A) PIP and PI(4,5)P
2
levels of
BHK cells permeabilized with 40 μ M  -
escin. All results are the average of two
to four measurements. Left bars give
levels for control cells and permeabi-
lized cells with 2 mM ATP and no Ca
(0.5 mM EGTA). The set of three bars
in the middle give PIP and PI(4,5)P
2
levels with ATP, without ATP for 5 min,
and with 2 mM AMP-PNP for 5 min. The
right two bars give phospholipid levels
of control cells and cells exposed to 0.4
mM free Ca for 2 min. (B) From left
to right, the bar graphs give mono and
diacylglycerol (M/DAG) and phospha-
tidate (PA) levels without and with 0.4
mM free Ca for 1 min (left four bars),
without and with 0.3 mM carbachol for
3 min, without and with 0.4 mM free Ca
together with 0.3 mM carbachol, and
without and with 0.4 mM free Ca for 1
min in the presence of AMP-PNP and
no ATP (right four bars).

40 Membrane Fusion and Phospholipids
are consistent with diacylglycerol kinases being activated
by Ca in these cells. Using cells with M1 receptors, we
did not obtain signi cant effects of carbachol (0.3 mM)
on acylglycerol levels, while phosphatidate increased
by 15%. In response to the combined application of
0.4 mM free Ca and carbachol (0.3 mM), acylglycerol
levels increased by 40% and phosphatidate levels in-
creased by 24%. Finally, as shown in the right four bars
of Fig. 9 B , incubation of cells with AMP-PNP (2 mM)
instead of ATP caused a decrease of acylglycerol. In the
presence of AMP-PNP, application of Ca (0.4 mM) for 1
min caused an increase of acylglycerol by 12% with no
change of phosphatidate. While the increase is small, as
a relative change, the absolute change is in fact larger
than the entire PI(4,5)P
2
mass of the cells, and we point
out that the increase of acylglycerol was accompanied by
a signi cant decrease of phosphatidylinositol (unpub-
lished data). Since PIP and PI(4,5)P
2
are depleted and
cannot be synthesized in this condition, phospholipases
evidently cleave signi cant amounts of phosphatidyl-
inositol. In summary, these biochemical data demon-
strate  rst that high cytoplasmic Ca indeed promotes
generation of phosphatidic acid by activating DAG ki-
nases, although this activity cannot explain the transloca-
tion of C1 domains into the cytoplasm described in Figs.
6 and 7 . Second, they demonstrate that Ca might indeed
support the generation of DAG on internal membranes
from sources other than PI(4,5)P
2
in the circumstances
of Figs. 6 and 7 .
Further Evidence against Roles for PLC, PLD, and cPLA2
Activities, as well as other Ca-dependent Processes, in
Membrane Fusion
Using the BHK cell line with high NCX1 expression,
we tested a wide range of agents expected to change
phosphatidylinositide metabolism, bind phosphati-
dylinositides, and/or inhibit phospholipases. All neg-
ative results reported here without  gures re ect at
least three and usually six or more control and treat-
ment observations. We found no signi cant inhibition
of membrane fusion or the rate of fusion by any of the
following agents when included in the pipette solution
and when fusion was quanti ed as a percent change
of capacitance and related to the peak exchange cur-
rent density: U73122 (10 μ M), edelfosine (10 μ M), nitro-
coumarin (20 μ M), neomycin (30 μ M), heptalysine
(40 μ M), and 2000 MW polylysine (20 μ M) to inhibit
PLCs and/or bind PI(4,5)P
2
and other anionic phospho-
lipids ( Ben-Tal et al., 1996 ; Bucki et al., 2000 ), IP
3
(0.1
mM) to release PLC-  from the membrane ( Cifuentes
et al., 1994 ), wortmannin (4 μ M) to inhibit PI3- and
type III PI4-kinases ( Nakanishi et al., 1995 ), adenosine
(1 mM) to inhibit type II PI4-kinases ( Barylko et al.,
2001 ), phosphatidylinositol transfer protein from yeast
(0.1 mg/ml) to remove phosphatidylinositol from cell
membranes ( Routt and Bankaitis, 2004 ), recombinant
same 1 min, PI fell from 46.3 to 44.3% of total anionic
phospholipid, while PA increased from 12.8 to 17.4% of
total phospholipid. As with receptor activation, the fact
that PI mass decreases and PA mass increases by similar
amounts, more than threefold greater than changes of
PIP and PI(4,5)P
2
, suggests that there is a substantial  ux
of phospholipid from PI to PIP to PI(4,5)P
2
to DAG and
 nally to PA in this protocol. In summary, these results
for cytoplasmic Ca elevation are qualitatively very similar
to, but quantitatively less drastic, than published results
for M1 receptor activation in cells with constitutive re-
ceptor expression ( Li et al., 2005 ).
In another approach, we established a BHK model
with the surface membrane permeabilized by  -escin
(40 μ M). As already noted, this concentration of  -escin
generates pores in most cells with a permeation cutoff
of 5 – 10 kD ( Fan and Palade, 1998 ). Therefore, we used
this model to analyze the dependencies of anionic phos-
pholipids and the total di- and monoacylglycerol on
Ca, both in the presence and absence of ATP. An extra-
cellular solution containing isotonic KCl (140 mM) with
2 mM ATP, 0.2 μ M free Ca (1 mM EGTA with 0.2 mM
Ca), and 2 mM MgCl
2
was employed. As demonstrated
in Fig. 9 A , phosphoinositide levels become highly de-
pendent on the presence of nucleotides in the extracel-
lular medium in this model, thereby demonstrating that
the surface membrane is highly permeable to solutes.
PIP and PI(4,5)P
2
levels of permeabilized cells were
30 and 60% lower than control cells, respectively ( Fig.
9 A , left dataset). Removal of ATP from the extracellu-
lar solution for 5 min caused PIP and PI(4,5)P
2
to fall
by ⵑ 80%, and the fall was still more pronounced for
PI(4,5)P
2
when AMP-PNP was included in the extracel-
lular solution. Addition of 2 mM Ca to the extracellular
solution for 2 min resulted in almost no change of PIP
levels but complete depletion of PI(4,5)P
2
( Fig. 9 A ,
right dataset).
Fig. 9 B shows our measurements of acylglycerols (M/
DAG) and phosphatidic acid, using diacylglycerol ki-
nase to generate phosphatidate and lysophosphatidate
from acylglycerols ( Preiss et al., 1987 ). In preliminary
experiments, we established that the bacterial DAG
kinase employed in this assay converted both 1 -O -palm-
itylglycerol and dipalmitylglycerol, in amounts > 3-fold
greater than determined for cell lysates to phosphory-
lated metabolites that were recovered quantitatively as
glycerol phosphate in our HPLC assay of anionic phos-
pholipid metabolites. As shown in Fig. 9 B , the total
di- and monocylglycerol content of BHK cells is similar
to that of phosphatidate, namely 6 – 8% of total anionic
phospholipid. For orientation, this is six to eight times
more than the content of PI(4,5)P
2
. As described in the
left four bars, the addition of Ca to generate a free con-
centration of 0.4 mM in the extracellular medium, contain-
ing 2 mM MgATP, caused a 30% decrease of acylglycerol
and a 15% increase of phosphatidate. These changes

Yaradanakul et al. 41
cause multiple free radical scavengers (TEMPO, 5 mM;
sucrose, 40 mM; dithiothreitol, 2 mM; acetylcysteine, 2
mM; and ascorbate, 4 mM) were without signi cant ef-
fect in these protocols at concentrations suggested in
the literature to suppress free radical signaling.
Failure to Correlate Capacitance Changes with Transporter
or Channel Activity Changes
The source of the membrane that traf cs in these ex-
periments is not well de ned. In this connection, we
tested whether Na/Ca exchange activities or Na/K
pump activities might increase with membrane fusion
or decrease with the subsequent fall of capacitance. To
do so, we activated the transport currents very brie y by
application of extracellular Ca or K, respectively, to de-
termine if maximal activity increases with membrane
fusion and/or decreases during the period of mem-
brane retrieval. We found no evidence that these trans-
porter activities followed the changes of membrane
area inferred from capacitance changes. In addition, we
C1A domains (20 μ M) to bind DAG ( Colon-Gonzalez
and Kazanietz, 2006 ), 0.6% butanol to stop DAG gen-
eration from phosphatidate (Choi et al., 2002), and a
putatively speci c phenylacrylamide cPLA2 inhibitor
(Calbiochem #525143, 2 μ M; Seno et al., 2000 ). The
fact that exchange currents are not inhibited by agents
that bind anionic phospholipids may appear surprising.
The likely reason is that in these protocols there are no
extracellular NCX1 ligands present until Ca is applied.
Therefore, exchangers orient with ion transport sites
open to the outside, and this con guration does not
support the Na-dependent inactivation ( Matsuoka and
Hilgemann, 1994 ). We also tested for possible roles of
other Ca-dependent processes. A possible role of Ca-ac-
tivated micro lament depolymerization seems unlikely
because phalloidin (5 μ M) had no signi cant effect
when perfused into cells for 5 min before activating
Ca in ux. A role of Ca-dependent proteolysis by cal-
pain ( Demarchi and Schneider, 2007 ) seems unlikely
because high concentrations of a peptidic calpain in-
hibitor IV (20 μ M, Calbiochem) had no signi cant ef-
fect when added to the cytoplasmic solution. A role of
Ca-activated free radical generation seems unlikely be-
Figure 10. Electron microscopic analysis of BHK cell vesicles in
close proximity to the surface membrane. (A) Electron micro-
graph of submembrane vesicles in a BHK cell. (B) Analysis of
noncoated and coated vesicle numbers identi ed at the given dis-
tances from the surface membrane of > 20 cells.
Figure 11. Typical staining and destaining in response to Ca in-
 ux in a BHK cell expressing NCX1. Background  uorescence
without dye is negligible. After incubating the cell with 10 μ M FM
4-64 for 50 s, Ca in ux was activated for 10 s. Thereafter,  uo-
rescence at the cell surface is greatly increased. Upon washing out
the FM dye after 2.2 min,  uorescence declines by 60%. When Ca
is introduced brie y a second time,  uroescence decreases rap-
idly by ⵑ 55%. This decrease corresponds to 28% of the total cell
 uorescence before dye washout. Fluorescence micrographs and
line scans at the time points indicated are provided above and
below the  uorescence graph, respectively.

42 Membrane Fusion and Phospholipids
average number of vesicles per square micron of mem-
brane. With an average diameter of 80 nm, fusion of at
least 50 vesicles per square micron would be required to
double membrane area. Our estimate of vesicular den-
sities was only ⵑ 10% of this value, indicating an impor-
tant discrepancy between ultrastructural analysis and
functional results. Some possible explanations are that
(1) vesicles are lost during the procedures employed,
(2) vesicles fuse during the  xation protocols employed,
and that (3) the vesicle population that fuses is gener-
ated during procedures to establish whole-cell voltage
clamp. We mention that pipette perfusion of the  xa-
tive solutions into cells did not cause membrane fusion,
based on capacitance recording.
It is beyond the scope of this article to determine the
source of the discrepancy just outlined. Rather, we used
optical methods to examine membrane cycling per se
in these protocols with a hydrophobic  uorescence
dye used in neuronal membrane cycling studies (e.g.,
Klingauf et al., 1998 ; Pyle et al., 1999 ). Fig. 11 shows typical
results for the dye, FM 4 – 64 ( Cochilla et al., 1999 ),
which binds and dissociates rapidly from membranes
and  uoresces only when bound. In preliminary stud-
ies, we labeled BHK cells with FM 4-64 for periods of
minutes to several hours with the expectation that de-
staining might be observed during the activation of
membrane fusion. However, in our experience com-
partments close to the surface membrane were not well
labeled with FM dye, and there was little destaining
upon activating Ca in ux in voltage-clamped cells. From
these results, we conclude that the membrane that cycles
performed similar experiments in cells expressing ROMK-
type potassium channels ( Zeng et al., 2003 ), and results
were similarly negative.
Evidence for Membrane Fusion as the Cause of Ca-
induced Capacitance Increase
The fact that phospholipase inhibitors had little or no
effect on membrane fusion tends to eliminate the possi-
bility that large biochemical changes could be causing
dielectric changes in these experiments. Nevertheless,
the large magnitudes of capacitance responses, in some
cases doubling cell capacitance, do raise questions about
the nature of the membrane fusing. Therefore, we per-
formed an ultrastructural study to analyze subplasma-
lemmal vesicles and membrane structures in BHK cells
that might participate in fusion events within 2 – 5 s of
activating Ca in ux. Cells were trypsinized and removed
from plates, as in preparation for electrophysiological
experiments. Cells were then  xed under conditions es-
tablished previously for ultrastructural studies of sub-
membrane compartments in neurons (see Materials and
methods). As shown in Fig. 10 , electron micrographs
indeed show the presence of numerous vesicles within
short distances of the surface membrane, both coated
and uncoated vesicles with diameters ranging from 45
to 90 nm. In > 20 cells examined, the distributions of
noncoated and coated vesicles were found to be simi-
lar in dependence on distance from the membrane.
Assuming that sections are 65 nm thick, we counted the
numbers of vesicles per linear micron of membrane
within 500 nm of the membrane, and we projected an
Figure 12. Extracellular cell surface accumulation
of K7-Rhod (3 μ M) upon activating maximal Ca
in ux in an NCX1-expressing BHK cell. (A) Cell
membrane  uorescence and whole cell capacitance
changes during and after application of 2 mM Ca
for 5 s. Both the control and the Ca-containing
solutions contain K7-Rhod. Fluorescence increases
with nearly the same time course as membrane ca-
pacitance, and the K7-Rhod probe washes off the
cell rapidly 45 s after activating Ca in ux. Note that
the background  uorescence of the light intensities
given in the graph ( ⵑ 20%) represents the homo-
geneous  uorescence of K7-Rhod throughout the
solution. (B) Average peak exchange currents and
membrane capacitance changes in response to 5 s
application of Ca for six control cells and six cells
perfused with 20 μ M 2000 MW polylysine. Results
are not signi cantly different. (C) Fluorescence and
membrane capacitance changes, determined as in
A, for 7 control cells, 8 cells perfused with 20 μ M
2000 MW polylysine, and 11 cells perfused with 0.2
μ M lactadherin.

Yaradanakul et al. 43
membrane lea ette. Given that this dye is a divalent cat-
ion, the increased dye binding in response to membrane
fusion may indicate that the extracellular membrane
surface becomes anionic during membrane fusion. This
explanation is supported by further experimentation
described in Figs. 12 and 13 . As apparent in Fig. 11 ,
ⵑ 30% of the total dye  uorescence typically did not
wash out in these experiments after inducing mem-
brane fusion and continuing dye application for 2 or
more minutes. The remaining  uorescence after wash-
out (e), which is roughly equivalent to the initial  uor-
escence on applying dye (b), is decreased by > 50% upon
applying a second pulse of Ca. This result suggests that
the membrane that is internalized after the  rst Ca pulse
can be fused again to the cell surface in response to a
second pulse of Ca, as expected from the reversibility of
capacitance changes in these experiments.
On the Nature of Cell Surface Changes during
Membrane Fusion
The extracellular cell surface might become anionic
for three reasons during protocols with massive Ca in-
 ux. First, high cytoplasmic Ca activates phospholipid
in voltage clamp experiments does not normally cycle
in these cells, but rather is available as a membrane re-
serve in response to massive Ca in ux, as expected with
cell wounding.
Next, therefore, we performed experiments to label
and destain BHK cells during voltage-clamp experi-
ments, and Fig. 11 shows typical  uorescence records.
Images and line scans used to determine the cell sur-
face  uorescence are shown below the  gure. Back-
ground  uorescence at the onset of the experiment was
negligible, and FM 4-64 (10 μ M) was applied and mem-
brane  uorescence increased to a steady state with a
time constant of a few seconds. After applying Ca to in-
duce membrane fusion there was a large further in-
crease of  uorescence with a time constant of about
a minute. As described subsequently, this increase re-
 ects, at least in part, internalization of membrane in
parallel with presentation of new membrane for stain-
ing. However, after 2.2 min, the majority of the increase
in  uorescence in response to Ca is rapidly reversed
upon removing dye. Thus, a large part of the signal does
not re ect membrane fusion and internalization, but
rather a changed af nity of the dye to the extracellular
Figure 13. Extracellular cell surface accumulation of annexin-V Alexa Fluor 488 conjugate in response to increasing cytoplasmic Ca
and activation of membrane fusion. In A and B, pipette perfusion of 40 mM Na was used to activate outward exchange current with 2
mM extracellular Ca and 1:100 annexin-V dilution. The pipette solution initially contains 40 mM Li. After recording background for
2 min, lithium is substituted for 40 mM Na via pipette perfusion through a 40- μ m inner diameter quartz capillary. Panel A shows the
surface membrane  uorescence, and B shows line scans of the cell before (solid line) and 5 min after activating exchange current.
(C) Inward exchange current and capacitance responses induced in giant excised BHK patches by applying cytoplasmic solutions with
4 and 200 μ M free Ca. The pipette contains 140 mM Na, and the pipette tip is switched between three solutions, as indicated. Capacitance
responses are negligible on applying 4 μ M free Ca. Two applications of 200 μ M free Ca result in a total 300 fF increase of patch capacitance,
estimated to be ⵑ 10% of the initial patch membrane capacitance. (D) Extracellular binding of annexin-V Alexa Fluor 488 to the extra-
cellular surface of a giant excised BHK patch upon applying 200 μ M free Ca to the cytoplasmic surface. The pipette solution contains 2
mM extracellular Ca with 1:100 annexin-V dilution.

44 Membrane Fusion and Phospholipids
concentration of 0.2 μ M, well above its K
d
for PS (Shi
et al., 2004), and cells were perfused with this solution for
3 min before activating exchange current. As shown in
Fig. 12 C , the increase of heptalysine binding on the
extracellular side was unaffected by the presence of
lactadherin on the cytoplasmic side.
To address speci cally whether PS increases in the
outer monolayer of BHK cells, we monitored the binding
of  uorescently labeled annexin-V (annexin-V Alexa
Fluor 488; 530BP) to cells, bearing in mind two limita-
tions. First, annexins require Ca to bind PS ( Maffey
et al., 2001 ) so that experiments must be performed en-
tirely in the presence of extracellular Ca. Second, while
several annexins bind PS selectively ( Ravanat et al.,
1992 ; Maffey et al., 2001 ; Kastl et al., 2002 ), the associa-
tion rates appear to be rather low so that binding occurs
with time constants on the order of 1 min ( Blackwood
and Ernst, 1990 ). Fig. 13 describes results from both
whole-cell recording and from giant excised membrane
patches. Fig. 13 (A and B) show the time course (A) and
distribution of annexin binding to voltage-clamped
BHK cells when outward exchange current is activated
by perfusing Na (40 mM) into cells via pipette perfu-
sion in substitution for Li (2 mM extracellular Ca with
annexin-V Alexa Fluor 488 at 1:100 dilution). Annexin-
V begins to bind to the outer cell surface abruptly and
continues almost linearly for the duration of the record-
ing. In six experiments, the cell surface  uorescence
increased by > 10-fold during Na perfusion. For the sta-
tistical analysis in Fig. 13 A , the cell membrane  uo-
rescence after 2 min annexin-V incubation without Na
was normalized to the  uorescence after perfusion of
Na for 5 min. As evident in the micrographs in Fig.
13 B , staining was typically not uniform. Patches of
annexin-V  uorescence are evident before inducing Ca
in ux. During the Ca in ux period, staining becomes
concentrated in areas that probably represent cell mem-
brane blebs.
Although annexin binding takes many seconds in sim-
ple assays, it is striking that binding in these experiments
shows no sign of saturation over more than 4 min after
activating Ca in ux. Thus, the extracellular appearance
of PS might be slower than the binding of cationic
agents, as described in the previous  gures. To allow a
more adequate kinetic analysis with well controlled Ca
concentrations on the cytoplasmic side, we performed
similar experiments using giant excised membrane
patches. As described in an accompanying article (Wang
and Hilgemann, 2008), large “ releasable ” pools of mem-
brane vesicles can be maintained in excised giant patches.
Fig. 13 C shows a typical recording from a BHK cell
patch with statistics for four patches in which inward ex-
change currents and capacitance changes were moni-
tored in response to rapidly applying cytoplasmic Ca.
In brief, we  nd that it is possible to maintain a fusable
membrane pool in BHK patches in which membrane
transport proteins ( ipases) that cause phospholipid
randomization between monolayers with externaliza-
tion of PS ( Bevers et al., 1999 ; Balasubramanian and
Schroit, 2003 ). In this same connection, most models of
membrane fusion require that, in order to achieve fu-
sion of opposing membranes, transitional hemifused
membrane structures must form in which nonbilayer
structures exist ( Siegel, 1999 ; Kozlovsky and Kozlov,
2002 ; Markin and Albanesi, 2002 ). Such structures may
facilitate phospholipid translocation across monolayers
( Homan and Pownall, 1988 ; Fattal et al., 1994 ). Second,
it is possible that the lumen of vesicles that fuse contain
large quantities of anionic phospholipids and/or an-
ionic sugars that become part of the extracellular mem-
brane monolayer during fusion. And third, membrane
proteins in the vesicles that fuse might contain unusu-
ally high numbers of anionic residues that can bind
and/or interact with FM dye. With this background, we
tested whether another  uorescent polyvalent cation
might bind to the extracellular surface of BHK cells
during the Ca response, similar to the FM dye. Fig. 12 A
shows cell-associated  uorescence changes with 3 μ
M
rhodamine-labeled heptalysine (K7-Rhod) in the extra-
cellular solutions. Fluorescence of the K7-Rhod solu-
tions was signi cant, and  uorescence at the edges of
voltage-clamped cells incubated with K7-Rhod was
nearly indistinguishable from this background (see “ a ”
in Fig. 12 A). Upon initiating Ca in ux by application of
2 mM extracellular Ca,  uorescence rose in 5 s to a high
level that is about  ve times greater than the solution
background. The time course of the  uorescence rise
nearly mimicked the time course of membrane fusion.
As illustrated further in Fig. 12 A ,  uorescence washed
off nearly completely within 10 s, indicating that the in-
crease of  uorescence is caused by increased binding at
the extracellular cell surface.
To address the different possible mechanisms just
outlined, we reasoned that the presence of agents that
bind PS on the cytoplasmic side would decrease and/or
slow the appearance of anionic groups on the outside if
PS randomization were a signi cant factor. Therefore,
we perfused cells with two agents that would be ex-
pected to bind PS signi cantly. To bind PS nonselec-
tively, 2000 MW polylysine (20 μ M) was added to the
perfusion solution. As shown for an initial set of experi-
ments in Fig. 12 B, the presence of cytoplasmic polyly-
sine did not signi cantly decrease the peak exchange
current or the percent increase of cell capacitance upon
activating exchange current. As shown in Fig. 12 C for
another set of experiments with optical recording, the
increase of heptalysine binding on the extracellular
side was unaffected by the presence of polylysine on the
cytoplasmic side. Lactadherin (Shi et al., 2004) is a milk
protein that binds PS with high af nity in a Ca-indepen-
dent manner. To bind PS selectively, therefore, puri ed
lactadherin was added to the cytoplasmic solution at a

Yaradanakul et al. 45
sion of a large population of vesicles, as predicted from
most membrane fusion models ( Siegel, 1999 ; Kozlovsky
and Kozlov, 2002 ; Markin and Albanesi, 2002 ), whereby
transmembrane voltage might transiently become di-
vided across two membranes in series. For reasons outlined
with the Results, transient conductance components in
our records ( Fig. 1 ) are likely to re ect the generation
of “ fusion pores ” during the fusion process ( Lindau
and Alverez de Toledo, 2003 ), although we cannot
entirely eliminate some contribution from changes of
transporter function that do not mirror the exchange
current per se.
Using a low af nity Ca indicator, we  nd that free cy-
toplasmic Ca peaks at ⵑ 0.2 mM with peak outward ex-
change currents of 0.1 – 0.2 nA activated for 2 – 4 s in cells
of 15 – 30 pF ( Fig. 2 ). Since the rate of rise of capaci-
tance increases over the entire range of exchange cur-
rents that can be activated ( Fig. 3 ), it seems certain that
the af nity of the underlying Ca sensor is rather low. In
experiments in which the maximal free cytoplasmic Ca
was thermodynamically capped by using equal cytoplas-
mic and extracellular Na concentrations ( Fig. 3, C and
D ), the half-maximal free Ca to activate the fusion re-
sponse is estimated to be 140 μ M, and this approximate
K
d
is consistent with results from excised patches de-
scribed in Fig. 13 C and in an accompanying article
(Wang and Hilgemann, 2008). As demonstrated in Fig.
3 B , clear increases of cell capacitance can nevertheless
be detected when free cytoplasmic Ca is limited to not
exceed 10 μ M (i.e., with 40 mM Na on both membrane
sides). This is readily explained if the Ca dependence of
the fusion process has only weak cooperativity, as deter-
mined here and in the accompanying article (Wang and
Hilgemann, 2008), in comparison to that observed for
neurotransmitter release in neurons ( Schneggenburger
and Neher, 2005 ). In fact, the Hill coef cients given in
Fig. 3 (2.2 and 2.1) may be an overestimate because Ca
buffering by EGTA (0.5 mM) can dampen the rise of
free Ca when exchange currents are small. Given the
low Ca cooperativity, it is plausible that the fusion mech-
anism analyzed here can be weakly activated by physio-
logical Ca transients, in addition to Ca in ux via cell
wounds. Both the apparent Ca af nity (K
d
ⵑ 120 μ M)
and the low cooperativity of responses (Hill coef cients
of ⵑ 2) are similar to those determined for asynchro-
nous neurotransmitter release in neurons ( Sun et al.,
2007 ). Finally, it seems notable that the maximal fusion
rate (i.e., dCap / dt ) decreases from one Ca in ux episode
to the next, while the maximal capacitance increase
does not decrease. Our explanation is that the rundown
of fusion simply re ects the rundown of exchange cur-
rents used to activate the response. Thus, the fusion mech-
anism itself seems to be highly resistant to washout of
soluble cell proteins, although it is modestly suppressed
over several minutes by substitution of ATP for a nonhy-
drolyzable ATP analogue ( Fig. 5 ).
rises quite high in the pipette tip and by taking precau-
tions to excise patches gently. As indicated below the
records, inward exchange currents were  rst activated
multiple times by 4 μ M free Ca, whereby the patch
capacitance remains stable. Thereafter, solution was
switched three times from the one with 4 μ M free Ca to
one with 200 μ M. This Ca concentration activates the
maximal exchange current, and patch capacitance in-
creases during the  rst two high Ca exposures.
As shown in Fig. 13 D , annexin-V binding from within
the patch pipette tip indeed increases rather rapidly in
this protocol. To monitor annexin-V binding, excised
patches were positioned nearly horizontal to the stage
to allow imaging of the pipette tip. Before applying
Ca to the cytoplasmic membrane face,  uorescence
of the annexin-V conjugate is negligible in the pipette
tip. Upon cytoplasmic application of 200 μ M free Ca,
membrane in the patch pipette becomes well de ned
by annexin  uorescence with a time constant of ⵑ 50 s
(three similar observations), substantially faster than in
whole cell experiments. Thus, we conclude that PS ap-
pearance in the extracellular cell surface may indeed
occur rapidly with membrane fusion, and that the ex-
periments with pipette perfusion of Na ( Fig. 13 A ) are
compromised by the use of pipette perfusion to activate
Ca in ux.
DISCUSSION
We have characterized a Ca-activated membrane fu-
sion mechanism in BHK  broblasts that can increase
membrane surface area by 25 – 100% in just a few seconds.
In parallel, PLCs cleave the bulk of cellular PI(4,5)P
2
and possibly substantial amounts of PI, and the extracel-
lular cell surface evidently becomes markedly anionic.
The molecular mechanisms and relationships of these
responses, which are discussed here, are of substan-
tial interest from multiple biological and biophysi-
cal perspectives.
Massive Membrane Fusion via Low Affi nity Ca Sensing in
BHK Cells
This study extends previous work in which “ caged Ca ”
was used to induce large Ca transients in CHO cells
( Coorssen et al., 1996 ). Clearly, the magnitudes of Ca-
induced capacitance changes in  broblasts can match
or exceed those occurring in cells with profuse neu-
rotransmitter release ( Kilic, 2002 ; Bauer et al., 2007 ;
Zeniou-Meyer et al., 2007 ). In this study, the responses
begin with a longer delay (100 – 500 ms) than described
for  ash photolysis, as expected to allow cytosolic Ca
accumulation by Ca transport. Differently from the
 ash photolysis studies, we routinely observe an initial
decrease of capacitance during which the conductance
signal component increases. One possible explanation
is that the negative capacitance phase re ects hemifu-

46 Membrane Fusion and Phospholipids
only very low rates ( Fig. 3 ). Overall, our work appears to
negate any role of PLCs in this type of membrane fusion
when Ca in ux is massive. The fusion responses were
unaffected by depletion of PI(4,5)P
2
(e.g., by receptor
activation in Fig. 7 ) and the parallel generation of DAG.
Activation of M1 receptors by carbachol can initiate
small increases of capacitance ( Fig. 7 C ), and as de-
scribed previously ( Yaradanakul et al., 2007 ), but subse-
quent Ca in ux still activates massive membrane fusion
( Fig. 7 ). And  nally, responses to maximal Ca in ux
were unaffected by multiple agents known to inhibit
PLCs (U73122, edelfosine, neomycin, or heptalysine),
as well as by pipette perfusion of DAG-binding C1 do-
mains, agents that circumvent DAG generation via PLD
pathways, namely 0.6% butanol (Choi et al., 2002),
and agents that are expected to bind PI(4,5)P
2
as well as
other anionic phospholipids ( Fig. 12 ). At this time, we
cannot entirely discount roles for phosphoinositides
and/or DAG-dependent mechanisms in long-term reg-
ulation of this type of fusion, or a role in setting the Ca
dependence of fusion. Clearly, ATP-dependent pro-
cesses do affect the maximal fusion amplitude ( Fig. 5 ),
and these are addressed further in an accompanying ar-
ticle (Wang and Hilgemann, 2008).
Our imaging results for PH and C1 fusion proteins al-
low comparisons of responses to receptor activation and
Ca elevation ( Figs. 6 and 7 ). PH domains respond to a
large increase of cytoplasmic Ca just as rapidly as they
respond to receptor activation, and in experiments at
35 ° C the domains reequilibrate after brief PLC activa-
tion with a time constant of ⵑ 1 min. These time courses
are likely to re ect the turnover rate of PI(4,5)P
2
in
these cells. As pointed out in Results, the C1 domains
reequilibrate somewhat more slowly after brief activa-
tion of PLCs. Of most interest, Ca in ux clearly has dual
in uences on the distribution of C1 domains. When C1
domains are brought to the surface membrane by mul-
tiple means ( Figs. 6 and 7 ), Ca in ux causes rapid trans-
location of the C1 domains back to the cytoplasm with a
time constant of < 1 min, and multiple results argue
against metabolism of DAG being the cause of C1 trans-
location. DAG kinase inhibitors were without effect, re-
sults were similar with PMA ( Fig. 7 ), which cannot be
metabolized, and results were similar when cells were
perfused with nonhydrolyzable ATP analogues.
Anionic Phospholipids and Membrane Fusion
The present results are relevant to several open ques-
tions about anionic phospholipids in membrane fusion.
Since the cytoplasmic Ca requirements for membrane
fusion in BHK cells are quite high, the binding of Ca by
anionic phospholipids can be considered as a possible
trigger for fusion. In fact, the fusion of anionic vesicles
composed of puri ed phospholipids can show a signi -
cant degree of selectivity for Ca versus Mg in the con-
centration range occurring in the present experiments
Several additional observations seem fundamental.
First, the capacitance responses can reverse completely
and can be repeated several times in the same cell ( Fig.
1 ). In the context of this article, it must suf ce to sum-
marize that the reversal is somewhat variable. Reversal
was often only partial (e.g., Fig. 4 ), but it could also ex-
ceed the initial membrane fusion response when al-
lowed to progress unabated after a short episode of Ca
in ux. Second, we analyzed functionally whether any
conductances readily analyzed, including Na/K pumps,
Na/Ca exchange, and some K channels, traf c with the
membrane that cycles during these responses, and our
outcomes were negative in all of these cases. A long-
term increase of cell conductance that often occurs with
each fusion episode (e.g., Fig. 1 ) probably re ects a de-
crease of the seal resistance. It would seem that mem-
brane proteins that enter the surface membrane during
fusion can be subsequently internalized without mixing
with the bulk of membrane proteins in the cell surface,
even during multiple fusion-retrieval episodes. This in-
direct conclusion is similar to conclusions for mem-
brane cycling during neurotransmitter release ( Willig
et al., 2006 ) that are presently under debate ( Wienisch
and Klingauf, 2006 ), and we stress that this conclusion
may not apply at all to membrane phospholipids. Third,
it is revealing that the membrane compartment that cy-
cles could not be labeled by preincubating cells with FM
dyes. This observation suggests that the membrane
compartment(s) that fuse to the surface do not traf c
under standard cell culture conditions. Fourth, we have
demonstrated that substantial numbers of vesicles are
in close proximity to the surface membrane of rounded
BHK cells, removed from dishes ( Fig. 10 ), but the quan-
tities of vesicles are not adequate to account for the fu-
sion responses. Further work with different approaches
will clearly be essential to account for the membrane
that cycles in response to Ca in ux. Fifth, although we
did not provide detailed analysis, it is our routine obser-
vation that the second fusion response is larger and/or
faster than the  rst response (e.g., Fig. 1 ). This “ facilita-
tion ” of fusion may re ect the same facilitation observed
for wound healing per se in other cell types, whereby
the rate of healing is approximately doubled at a sec-
ond response ( Togo et al., 2003 ; Steinhardt, 2005 ).
Ca-dependent PLC Activation in BHK Cells Appears to be
Unrelated to the Initiation of Membrane Fusion
As outlined in the Introduction, both DAG and PI(4,5)P
2
may modulate membrane fusion events by multiple
mechanisms. As noted in connection with Fig. 6 , the cy-
toplasmic Ca concentrations needed to activate fusion
at half-maximal rates are probably greater than those
needed to activate PLCs in BHK cells without receptor
activation. As noted further with those Results, we could
clearly detect PLC activities with 10 μ M free cytoplasmic
Ca, a concentration that causes membrane fusion at

Yaradanakul et al. 47
studies of annexin binding to PS-containing membranes
indeed show rather slow kinetics ( Blackwood and Ernst,
1990 ; Kastl et al., 2002 ). Thus, the results may be kineti-
cally limited by the rate of annexin-V binding. In fact,
annexin-V binds substantially faster to the extracellular
surface of giant excised patches, when Ca is rapidly ap-
plied to the cytoplasmic side ( Fig. 13 D ), than occurs in
the whole cell experiments. Together, the results of Figs.
12 and 13 would suggest that PS does not translocate
from the cytoplasmic lea et but rather may appear in
the extracellular surface immediately as a result of
membrane fusion, as expected if the inner lea et of the
membranes of vesicles that fuse is anionic. Clearly, this
suggestion requires more experimentation in which
means are found to inhibit membrane fusion selectively
without inhibiting Ca transients. We point out that extra-
cellular PS exposure has been described previously in
mast cells in response to degranulation ( Demo et al.,
1999 ; Martin et al., 2000 ) and in auditory hair cells in as-
sociation with membrane cycling ( Shi et al., 2007 ). Simi-
lar to the present study, it is an open question in both of
those studies whether PS “ translocates ” from the cyto-
plasmic lea et of the surface membrane during fusion
or whether it enters the extracellular cell surface mono-
layer from the luminal monolayer of vesicles. A parallel
problem in red blood cells is that phospholipid scram-
bling can be associated with membrane shedding. To ac-
count for membrane shedding, a membrane “ division ”
process must occur in which transiently nonbilayer mem-
brane structures are generated ( Comfurius et al., 1990 ).
In summary, we have analyzed in more detail than
heretofore massive membrane fusion at the cell surface
of a standard immortalized  broblast in response to a
rapid increase of cytoplasmic Ca. The activation of PLCs
that occurs in parallel appears to be unrelated to the fu-
sion process and does not modulate it to any signi cant
degree in our experiments with massive Ca in ux. Our
results to date are consistent with membrane fusion
causing the appearance of anionic phospholipids at the
extracellular surface by a mechanism that does not in-
volve PS “ scrambling ” between monolayers. Our results
provide no support for any speci c or nonspeci c role
of “ signaling ” phospholipids in this type of membrane
fusion or its triggering by Ca.
We thank Marc Llaguno (University of Texas Southwestern Medi-
cal Center at Dallas) for critical discussions and advice, Kenneth
D. Philipson (University of California, Los Angeles, CA) for the
BHK cell line, NCX1 reagents and discussions, Mark Shapiro
(University of Texas at San Antonio, TX) for hM1 constructs em-
ployed, and Gary Gilbert (Harvard Medical School, Boston, MA)
for lactadherin protein.
This work was supported by HL0679420 and HL051323 to D.
W. Hilgemann.
Lawrence G. Palmer served as editor.
Submitted: 6 August 2007
Accepted: 25 April 2008
( Newton et al., 1978 ; Duzgunes et al., 1981 ; Summers
et al., 1996 ), and furthermore, Ca binding to PS-con-
taining bilayers brought into close apposition can occur
with both high selectivity and af nity ( Feigenson, 1986 ;
Feigenson, 1989 ). Our results contradict this possibility
because membrane fusion is unaffected by agents that
bind PS (polylysine and lactadherin in Fig. 12 ), as well
as other anionic phospholipids, and would be expected
to sequester them away from other binding partners. It
would be interesting to know if C1 domains, which bind
PS as well as DAG ( Johnson et al., 2000 ; Bittova et al.,
2001 ), might be released from cell membranes by these
PS binding agents. As evident in Fig. 7 (B and C) , the
shift of C1 domains to the cytoplasm in response to Ca
in ux can reverse very rapidly when Ca in ux is termi-
nated, as expected if fast Ca dissociation from anionic
phospholipids favors C1 binding to the membrane.
At present we have not entirely eliminated one alter-
native explanation for the Ca-dependent movement
of C1 domains. As shown by mass measurements of
phospholipids, mono- and/or diacylglycerols increase
with high cytoplasmic Ca, even after PI(4,5)P
2
deple-
tion by receptor activation or removal of ATP ( Fig. 9 B ).
If DAG increases markedly when Ca rises, DAG on inter-
nal membranes might draw C1 domains away from the
surface membrane. One argument against this idea is
that total basal DAG levels appear to be higher than
PI(4,5)P
2
levels, as in other cell types ( Callender et al.,
2007 ), and therefore DAG concentrations in internal
membranes may already be high in the basal state. A
second argument against this idea is that the C1 domains
appear uniformly distributed in the cytoplasm after ac-
tivating Ca in ux, as expected if they are not associated
with speci c membrane compartments.
From the extracellular side, it is striking that the af n-
ity of cationic agents for the extracellular cell surface in-
creases profoundly and rapidly with membrane fusion
( Figs. 11 and 12 ). As pointed out in Results, there are
multiple possible explanations. If PS is translocating
from the cytoplasmic lea et to the extracellular mem-
brane lea et, for example as a result of nonbilayer struc-
tures occurring during membrane fusion, one would
expect that agents that bind PS would slow and/or in-
hibit the process when applied from the cytoplasmic
side. As shown in Fig. 12 that was not the case for either
polylysine or the PS-binding protein lactadherin. An-
nexin-V binds much more slowly to the extracellular sur-
face than the cationic agents in whole cell recording
( Fig. 13, A and B ), suggesting that PS may appear more
slowly than membrane fusion occurs. However, the an-
nexin results are not de nitive. First, whole-cell experi-
ments are limited by the fact that Ca must be present on
the outside to allow annexin binding. Using cell perfu-
sion of Na to activate Ca in ux, it is certain that Ca tran-
sients are slower and of lower magnitude than for rapid
activation of reverse exchange currents. Second, kinetic

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