Báo cáo khoa học: The proximity between C-termini of dimeric vacuolar H+-pyrophosphatase determined using atomic force microscopy and a gold nanoparticle technique ppt

The proximity between C-termini of dimeric vacuolar
H
+
-pyrophosphatase determined using atomic force
microscopy and a gold nanoparticle technique
Tseng-Huang Liu
1
, Shen-Hsing Hsu
1
, Yun-Tzu Huang
1
, Shih-Ming Lin
1
, Tsu-Wei Huang
2
,
Tzu-Han Chuang
3
, Shih-Kang Fan
4
, Chien-Chung Fu
3
, Fan-Gang Tseng
2,
* and Rong-Long Pan
1,
*
1 Department of Life Sciences and Institute of Bioinformatics and Structural Biology, College of Life Sciences, National Tsing Hua Univer-
sity, Hsin Chu, Taiwan, ROC
2 Department of Engineering and System Science, National Tsing Hua University, Hsin Chu, Taiwan, ROC
3 Institute of NanoEngineering and MicroSystems, National Tsing Hua University, Hsin Chu, Taiwan, ROC

4 Institute of Nanotechnology, National Chiao Tung University, Hsin Chu, Taiwan, ROC
Introduction
Vacuolar H
+
-pyrophosphatase (V-PPase; EC 3.6.1.1)
is a homodimeric protein with a monomeric molecular
mass of 71–80 kDa [1]. V-PPase catalyzes electrogenic
proton translocation from the cytosol to the vacuolar
lumen to generate an inside-acidic and inside-positive
membrane potential for the secondary transport of
ions, metabolites, and toxic substances [1–3]. The
cDNAs of V-PPase have been cloned from several
higher plants, some protozoa, and several species of
eubacteria and archeubacteria, and are highly similar
(86–91% deduced amino acid identity) [1,3,4]. V-PPase
requires Mg
2+
as a cofactor, and the binding of Mg
2+
Keywords
atomic force microscopy; proton
translocation; tonoplast; vacuolar
H
+
-pyrophosphatase; vacuole
Correspondence
R L. Pan, Department of Life Sciences and
Institute of Bioinformatics and Structural
Biology, College of Life Sciences, National
Tsing Hua University, Hsin Chu 30013,

Taiwan, ROC
Fax: +886 3 5742688
Tel: +886 3 5742685
E-mail:
*These authors contributed equally to this
work
(Received 1 March 2009, revised 17 May
2009, accepted 10 June 2009)
doi:10.1111/j.1742-4658.2009.07146.x
Vacuolar H
+
-translocating inorganic pyrophosphatase [vacuolar H
+
-pyro-
phosphatase (V-PPase); EC 3.6.1.1] is a homodimeric proton translocase; it
plays a pivotal role in electrogenic translocation of protons from the cyto-
sol to the vacuolar lumen, at the expense of PP
i
hydrolysis, for the storage
of ions, sugars, and other metabolites. Dimerization of V-PPase is neces-
sary for full proton translocation function, although the structural details
of V-PPase within the vacuolar membrane remain uncertain. The C-termi-
nus presumably plays a crucial role in sustaining enzymatic and proton-
translocating reactions. We used atomic force microscopy to visualize
V-PPases embedded in an artificial lipid bilayer under physiological condi-
tions. V-PPases were randomly distributed in reconstituted lipid bilayers;
approximately 43.3% of the V-PPase protrusions faced the cytosol, and
56.7% faced the vacuolar lumen. The mean height and width of the cyto-
solic V-PPase protrusions were 2.8 ± 0.3 nm and 26.3 ± 4.7 nm, whereas
those of the luminal protrusions were 1.2 ± 0.1 nm and 21.7 ± 3.6 nm,

respectively. Moreover, both C-termini of dimeric subunits of V-PPase are
on the same side of the membrane, and they are close to each other, as
visualized with antibody and gold nanoparticles against 6·His tags on
C-terminal ends of the enzyme. The distance between the V-PPase C-termi-
nal ends was determined to be approximately 2.2 ± 1.4 nm. Thus, our
study is the first to provide structural details of a membrane-bound
V-PPase dimer, revealing its adjacent C-termini.
Abbreviations
AFM, atomic force microscopy; DDM, n-dodecyl-b-
D-maltoside; GNP, gold nanoparticle; SD, standard deviation; V-PPase, vacuolar
H
+
-pyrophosphatase; TEM, transmission electron microscopy.
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4381
can stabilize and activate the enzyme [1,5]. Relatively
high concentrations of K
+
stimulate the proton-trans-
locating function of V-PPase, whereas excess amounts
of PP
i
,Ca
2+
,Na
+
and F
)
inhibit its enzymatic activ-
ity [6–8]. It is conceivable that the V-PPase provides
specific binding domains for the substrate and the

above-mentioned ions, as well as proton translocation.
Truncation of the C-terminus induces a dramatic
decline in V-PPase enzymatic activity, proton translo-
cation, and coupling efficiency [9]. In addition, deletion
of the C-terminus of V-PPase increases its susceptibil-
ity to heat stress and substantially increases the appar-
ent K
+
binding constant. It is thus likely that the
C-terminus plays an essential role in sustaining the
physiological functions of V-PPase.
Interactions between the subunits of the V-PPase
dimer have been studied [1,2,10–12]. Radiation inac-
tivation analysis demonstrated that the proper
dimeric structure of V-PPase on tonoplastic mem-
branes is a prerequisite for both enzymatic activity
and PP
i
-supported proton translocation [2,11,12].
Further target size measurements revealed that only
one subunit of the purified dimeric complex was suf-
ficient for the enzymatic reaction of V-PPase,
although proton translocation requires the presence
of both subunits [2]. Moreover, high hydrostatic
pressure was employed to inhibit V-PPase through
subunit dissociation of the enzyme, resulting in inac-
tive forms [10]. The physiological substrate and sub-
strate analogs enhance the high-pressure inhibition of
V-PPase, indicating the vulnerability of the subunit–
subunit interaction [10]. The above lines of evidence

illustrate explicitly the importance of dimer forma-
tion for V-PPase function, and suggest nonrandom
and sequestered association of V-PPase subunits
within the vacuolar membrane. Furthermore, the
structures of purified V-PPases from pumpkin (Cu-
curbita sp. Kurokawa Amagur) and Thermotoga mar-
itime have been examined by electron microscopy
[13,14]. Notwithstanding this, structure–function rela-
tionships within this proton-translocating complex
require further study.
Atomic force microscopy (AFM) is a powerful
tool used for nanoscale structural analysis of protein
complexes [15,16], and of supported lipid bilayers in
particular [17–20]. For instance, AFM has provided
marvelously high-resolution images of purified
dimeric membrane proteins in 2D crystals and of
densely packed proteins in native membranes [21–23].
In the present study, we used AFM to directly
observe purified V-PPases reconstituted into planar
lipid bilayers under physiological conditions. Our
images unambiguously reveal a dimeric complex for
this proton-transporting V-PPase. Furthermore, the
molecular volume of V-PPase calculated from AFM
images suggests the presence of two identical subun-
its, verifying the notion of the homodimeric V-PPase
enzyme. A gold nanoparticle (GNP) technique com-
bined with transmission electron microscopy (TEM)
analysis was utilized to determine the distance
between C-termini within a membrane-bound
V-PPase dimer, and indicated that the C-termini are

located at the interface of subunits.
Results and discussion
AFM analysis of purified V-PPase adsorbed onto
mica
Recombinant DNAs for overexpression of V-PPases
containing a 6·His tag at either the C-terminus or
N-terminus were prepared and transformed into a
yeast host. Recombinant V-PPase containing a 6·His
tag at the C-terminus (Fig. 1C) was overexpressed in
yeast and successfully purified from microsomes.
Unfortunately, V-PPase containing a 6·His tag at
the N-terminus was poorly expressed in yeast and
was therefore excluded from the study (data not
shown). SDS/PAGE analysis of the purified C-termi-
nal 6·His-tagged V-PPase followed by Coomassie
Blue staining or western blotting showed that it was
highly purified, comprising a single major band with
a molecular mass of 73 kDa (Fig. 1A), as expected
from the known structure of the V-PPase monomer
[1,2,10]. During size exclusion chromatography,
V-PPase was eluted with an apparent molecular mass
of  145 kDa, similar to its native form and in
agreement with previous studies suggesting a dimeric
conformation [2,10–12].
The purified V-PPase was then reconstituted into
liposomes by a detergent removal method using Bio-
Rad SM-2 beads combined with freeze–thaw sonica-
tion [13]. On addition of PP
i
to the proteoliposome

solution containing Mg
2+
, a dramatic decrease in pH
was generated in the interior of the liposomes (Fig. 1B,
lower trace). The acidic pH was eliminated by the
addition of the ionophore gramicidine D (5 lgÆmL
)1
),
indicating the integrity of the membrane (data not
shown). The liposomes alone (without V-PPase) did
not exhibit proton-translocating activity (Fig. 1B,
upper trace).
Individual V-PPase molecules were adsorbed
randomly on the mica surface and exhibited a proto-
typical globular structure under physiological condi-
tions (Fig. 2A). Figure 2B shows the heights of the
adsorbed particles along the cross-section in Fig. 2A.
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4382 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
The width of individual protein molecules was
measured at half the vertical height. The mean width
and height [± standard deviation (SD), n = 21] of
purified V-PPase were 22.5 ± 3.2 nm and 1.6 ±
0.4 nm, respectively. Furthermore, major peaks on
height and width histograms for the AFM images
also concurred with those parameters obtained above
for V-PPase molecules (Fig. 2C,D). The flattening of
particles was presumably caused by the interaction

between the polar surface of the protein molecules
and the charged surface of the mica [24]. These
images represent the first direct nanoscale observation
of V-PPase.
Determination of molecular volume provides the
stoichiometry of subunit components for functional
enzymes [24]. In this study, the volume of V-PPase
was calculated using Eqn (1) and determined to be
302.4 ± 40.6 nm
3
(V
s
)(n = 21), which was slightly
larger than the theoretical value (V
prot
; 274.5 nm
3
)of
the protein (Table 1). This slight overestimation in
volume probably arose from the broadening effect of
the AFM tip [24]. It is also likely that variations in
volume measurements might arise from distinct inter-
actions of the tip with the individual purified
V-PPase particles [25]. Nonetheless, these results
unambiguously demonstrate the feasibility of this
technique for nanoscale investigation of purified
V-PPase molecules.
AFM analysis of V-PPase reconstituted into
liposomes
The homodimeric structure of V-PPase in a planar

lipid bilayer was imaged directly by AFM (Fig. 3).
Purified V-PPase was first reconstituted into a sup-
ported lipid bilayer, as confirmed by immunofluores-
cence imaging (Fig. 3A). Figure 3A1 shows a planar
lipid bilayer reconstituted with V-PPases and ana-
lyzed by immunofluorescence using a primary anti-
body against His followed by a Cy3-conjugated
secondary antibody; no fluorescence was detected in
bilayers without immunofluorescence labeling of the
protein (Fig. 3A2). In addition, no immunofluores-
cence was observed when the reconstituted sample
was incubated directly with the Cy3-conjugated sec-
ondary antibody in the absence of primary antibody
against His (Fig. 3A3). Lipid bilayers lacking
V-PPases also did not exhibit detectable fluorescence
(Fig. 3A4). These results indicated successful incorpo-
ration of V-PPase into a lipid bilayer, allowing for
subsequent AFM analysis.
To obtain high-resolution AFM images of individ-
ual V-PPases within reconstituted membranes, the
proteoliposomes prepared above were fused into a
large planar lipid bilayer for direct observation. The
thickness of the lipid bilayer without any protein was
approximately 4.6 ± 0.5 nm (n = 12), determined
Fig. 1. Purification and proton transport
activity of V-PPase. (A) Analysis of purified
V-PPase by western blotting (top) and SDS/
PAGE and Coomassie Blue staining (bot-
tom). Lane 1: V-PPase-enriched microsome.
Lane 2: purified V-PPase. Lane 3: reconsti-

tuted V-PPase. Molecular mass (kDa) mark-
ers are indicated on the left. (B) PP
i
-
associated proton translocation of reconsti-
tuted V-PPase. Proton transport was initi-
ated by adding 1.0 m
M PP
i
. At the end of
each reaction, 5 lgÆmL
)1
gramicidin D was
added to stop the fluorescence quenching
of acridine orange. (C) Topological model of
V-PPase. Cylinders 1–16 indicate mem-
brane-spanning domains.
T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4383
from a cross-section of the lipid bilayer. The bilayer
thickness was consistent with previous AFM mea-
surements of a lipid bilayer composed of a phospha-
tidylcholine/cholesterol mixture and prepared in a
similar aqueous environment [26]. V-PPases reconsti-
tuted into the lipid bilayer protruded from the
bilayer surface in a diffuse pattern with a random
distribution. The V-PPase images fell within two cat-
egories according to the extramembranous protrusion

height. These height differences reflect two distinct
populations of individual V-PPases facing the recon-
stituted membrane surface (Fig. 3D,E). Three-dimen-
sional analysis of individual V-PPases randomly
distributed on the membrane surface indicated that
56.7% of the protrusions were small, with a mean
height of 1.2 ± 0.1 nm (n = 20) (Fig. 3B, solid cir-
cles), and the remainder of the protrusions (43.3%)
were large, with a mean height of 2.8 ± 0.3 nm
(n = 17) (Fig. 3B, dotted circle). The uneven distri-
bution and/or orientation of V-PPases on the mem-
brane suggests that targeting of the V-PPase into the
vacuolar membrane of plant cells may follow a spe-
cific pattern, as previously suggested [10]. Figure 3C
shows a cross-section along the line in Fig. 3B. The
widths and heights of the reconstituted V-PPase pro-
trusions in Fig. 3B are listed in Table 1. The AFM
image of reconstituted V-PPase shows a ratio of
approximately 2.40 : 1 for the height values of the
cytosolic and luminal sides. In addition, the theoreti-
cal ratio of the total amount of amino acids on the
cytosolic and luminal sides was calculated as 2.31 : 1
(data not shown), verifying the efficacy of this tech-
nique.
The high protein density in 2D crystals or in native
membranes allows high-resolution AFM topographs
and the elucidation of protein subunit organization
[21–23]. However, it is presently difficult to obtain
V-PPase reconstituted in 2D crystals or packed at high
density into a membrane (data not shown). Notwith-

standing this, current AFM techniques suffice to
provide unambiguous images of the dimeric structure
of V-PPase. Four representative examples exhibiting
minor variations are shown in Fig. 4A. The small
differences in topography of the individual V-PPases in
A
B
D
C
Fig. 2. AFM analysis of purified V-PPase. (A) Three-dimensional
AFM image of purified V-PPase adsorbed onto mica. (B) Profile of
peak heights along the cross-section shown in (A). Purified V-PPase
protrudes 1.6 ± 0.4 nm (n = 21) from the mica surface. (C) Histo-
gram of V-PPase height determined using the AFM image in (A).
(D) Histogram of V-PPase width determined using the AFM image
in (A).
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4384 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
the reconstituted lipid bilayer have probably resulted
from contact with the AFM tip during scanning.
Nevertheless, these AFM images are adequate for
nanoscale resolution of the structural details of
V-PPase [27,28]. Moreover, the resolution of the
images from V-PPases in reconstituted membranes was
Table 1. Dimensions of free and membrane-bound recombinant V-PPase determined by AFM. Values represent means ± SD. n = number
of observations. Observed and predicted volumes were determined from AFM analysis using Eqn (1) and from theoretical analysis using
Eqn (2).
Protein (145 kDa) Height (nm) Width (nm)

Volume (nm
3
)
Observed Predicted
Purified V-PPase (n = 21) 1.6 ± 0.4 22.5 ± 3.2 302.4 ± 40.6 274.5
Reconstituted V-PPase 332.9 ± 46.9 274.5
Lumen side (n = 20) 1.2 ± 0.1 21.7 ± 3.6
Cytosolic side (n = 17) 2.8 ± 0.3 26.3 ± 4.7
Lipid bilayer (n = 12) 4.6 ± 0.5
Fig. 3. Reconstitution of V-PPase into proteoliposomes. (A) Immunofluorescence imaging of V-PPases reconstituted into lipid bilayers. (1)
Sample treated with primary and secondary antibodies. (2) Sample not treated with either antibody. (3) Sample treated with only secondary
antibody. (4) Lipid bilayer lacking V-PPases but treated with primary and secondary antibodies. (B) AFM image of V-PPase extramembranous
protrusions on the luminal and cytosolic sides of the membrane. Solid circle, luminal side; dotted circle, cytosolic side. Inset: section of a
lipid bilayer with thickness of 4.6 ± 0.5 nm (n = 12). (C) Profile of protrusion heights along the cross-section shown in (B). Two populations
of V-PPase protrusions were observed: one with a mean height of 1.2 ± 0.1 nm (n = 20), and one with a mean height of 2.8 ± 0.3 nm
(n = 17). (D) Histogram of V-PPase protrusion heights determined using the AFM image in (B). (E) Histogram of V-PPase peak widths deter-
mined using the AFM image in (B).
T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4385
typically higher than that of those directly adsorbed
onto a mica surface.
The volume of the V-PPase homodimers (V
m
) in the
reconstituted membrane was estimated using the height
of the protein protrusion and the thickness of the lipid
bilayer as the parameters for the volume of a sphere
(Fig. 4B). The volume of reconstituted V-PPase was

measured as 332.9 ± 46.9 nm
3
(n = 17). The V
prot
of
a V-PPase homodimer with a molecular mass of
145 kDa, calculated on the basis of the amino acid
composition, was determined to be 274.5 nm
3
[29].
This theoretical volume correlates very well with that
measured from the AFM images. Note that these
images were obtained by AFM scanning in a fluid,
and therefore probably provide an authentic illustra-
tion of V-PPase structure under physiological condi-
tions. The AFM images indicate the dimeric structure
of V-PPase reconstituted in a lipid bilayer. This study
provides the first 3D representation of individual
V-PPases protruding from the cytosolic and luminal
sides of a membrane in aqueous solution.
Proximity of V-PPase C-termini in reconstituted
membranes
Topology studies examining heterologous V-PPase
expression in yeast suggested that both the C-termini
and the N-termini of each subunit are located on the
lumen side and are opposite the catalytic domain on
the cytosolic side of the vesicular membrane [1].
Because V-PPase is homodimeric, there are two possi-
ble configurations for association of the two subunits;
the C-termini of both subunits may protrude from the

same side or from opposite sides of the membrane
Fig. 4. High-resolution AFM image of
V-PPase dimers in a reconstituted
membrane. (A) AFM analysis of
extramembranous protrusions on the
cytosolic side of proteoliposomes containing
V-PPase (top panels) and those on the
luminal side (bottom panels). (B) Topological
model of the homodimeric structure of
V-PPase.
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4386 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
[30]. The present study demonstrated two distinct types
of protrusions randomly distributed in reconstituted
lipid bilayers. If the C-termini of each V-PPase subunit
within a dimer protruded from opposite sides of the
membrane, the measured heights of these two types of
protrusion should presumably be the same. The nega-
tive results above thus indicate that the C-termini of
the individual subunits of the enzyme are facing the
same side of the membrane.
The relative positions and proximity of the V-PPase
C-termini on the surface of the reconstituted mem-
branes were examined using an IgG antibody against
the C-terminal His tag of the enzyme (Fig. 5). The
AFM image of the immunolabeled V-PPase showed
that protrusions of different heights and widths were
randomly distributed on the lipid bilayer (Fig. 5A). The

antibody could bind to V-PPase on either one or two
molecules (Fig. 5B). Clearly, Fig. 5B2 depicts that two
antibodies bind respectively to a single V-PPase mole-
cule in close vicinity. AFM image analysis using spip
software was used to generate histograms delineating
the distribution of protrusion heights and widths
(Fig. 5C,D), and this revealed three major groups of
protrusions: (a) lower peaks (peak 1; 1.4 ± 0.2 nm
mean height, n = 10) for structures of V-PPase on the
lumen side of the membrane lacking bound antibody;
(b) intermediate peaks (peak 2; 2.9 ± 0.2 nm mean
height, n = 20) for those on the cytosolic side of the
membrane; and (c) higher peaks (peak 3; 4.2 ± 0.3 nm
mean height, n = 10) for antibodies bound presumably
to the lumen side. The ratio of the sum of integrals for
peak 1 and peak 3 (free lumen side and antibodies bind-
ing to the lumen side) to peak 2 (cytosolic side) is con-
sistent with our prior results (approximately 5.6 : 4.4).
Fig. 5. AFM analysis of V-PPase in a reconstituted lipid bilayer immunolabeled with an antibody against His to detect the C-terminal 6·His
tag of V-PPase. (A) Image of a large section of immunolabeled lipid bilayer reconstituted in the presence of V-PPase. (B) High-resolution
images of immunolabeled protrusions in (A). (1) Protrusion showing a single antibody bound to V-PPase. (2) Protrusion showing two
antibodies bound to V-PPase. (C) Histogram of protrusion height determined using the AFM image in (A). (D) Histogram of protrusion width
determined using the AFM image in (A).
T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4387
Previous AFM imaging studies have demonstrated
that the height of a single IgG molecule is
2.4 ± 0.1 nm [24]. Taking this value into account, the

height of peak 3 protrusions (4.2 ± 0.3 nm, n = 10)
would be that of IgG molecules (2.4 ± 0.1 nm) sitting
on V-PPase at the lumen side (1.2 ± 0.1 nm, n = 20).
There were also two major groups in the histogram
representing the distribution of protrusion widths,
probably for those of the single IgG molecule (mean
width = 41.5 ± 1.8 nm, n = 12; 38.5% of protru-
sions) and those of two IgG molecules (mean width =
51.3 ± 0.8 nm, n = 20; 62.5% of protrusions) bound
to a V-PPase in the lipid bilayer, respectively
(Fig. 5D). It is well established that the hinge region of
IgG links the two Fab arms to the Fc portion, provid-
ing global flexibility to the IgG. The flexibility of the
IgG molecule results in Fab ‘elbow bending’, Fab ‘arm
waving’ and rotation, and Fc ‘wagging’ [31]. The
observed variations in the number of IgG molecules
bound to the 6·His-tagged C-termini of V-PPase
subunits have presumably arisen from such antibody
flexibility. Therefore, the space between the two anti-
body molecules could not be precisely determined
using current techniques. As a result, we were also
unable to accurately determine the proximity of the
V-PPase C-termini using the antibody-binding
technique.
We hence employed Ni
2+
–nitrilotriacetic acid GNP
labeling as an alternative technique to evaluate the
proximity of C-termini within V-PPase homodimers.
Extremely small Ni

2+
–nitrilotriacetic acid GNPs were
bound to the 6·His tags of V-PPase C-termini recon-
stituted in lipid bilayers in aqueous solution, resulting
in two major types of protrusion as observed with
AFM: the cytosolic side of V-PPase, and the particles
bound to the lumen side of V-PPase, respectively
(Fig. 6). The solid circle in Fig. 6B indicates GNP
bound to V-PPase C-terminus protruding from the
surface of the lipid bilayer, whereas the dotted circle,
V-PPase protrusion at the lumen side lacking bound
GNP (Fig. 6B). More than 70% of V-PPases were
covered by GNPs on the luminal side (data not
shown). The height distribution histogram indicated
that the heights of the lower V-PPase protrusions
(peak 1) were consistent with those of its cytosolic por-
tions, whereas the heights of the higher ones (peak 2)
represented those of the GNPs bound to the C-termini
of the enzyme (Fig. 6C). The height of the latter
protrusions (4.9 ± 0.1 nm) reflects the sum of Ni
2+
–
nitrilotriacetic acid GNP heights (mean height =
2.0 ± 0.1 nm, n = 16) and V-PPase heights on the
luminal side (mean height = 1.2 ± 0.1 nm, n = 20).
In contrast, the lower V-PPase protrusions represent
those of its cytosolic sides alone. Moreover, in the
width distribution histogram, the higher peaks (peak
1) represent either cytosolic protrusions of V-PPase
lacking GNPs (mean width = 26.3 ± 4.7 nm, n = 17)

or the luminal side containing GNP-bound C-termini
(mean width = 28.2 ± 1.4 nm, n = 48). Other peaks
(peaks 2 and 3) in the width distribution histogram
(> 50 nm) probably reflect GNP clusters, because
 20% of GNPs in solution are visualized as collec-
tions after sonication (data not shown).
The number of GNPs bound to V-PPase C-termini
was then predicted using a Microscope Simulator (Com-
puter Integrated Systems for Microscopy and Manipula-
tion, University of North Carolina, Chapel Hill, NC,
USA) (Fig. 6E). The width of the image for a single
GNP on mica was empirically determined as
21.2 ± 1.1 nm (n = 27, data not shown); the theoreti-
cal width of a single GNP on the surface of V-PPase was
24 nm (Fig. 6E, solid rhombus). The mean width of
GNPs on the surface of V-PPase was empirically mea-
sured as 28.2 ± 1.4 nm (n = 48), suggesting that more
than one GNP was present on the surface of V-PPase.
Because V-PPase is a homodimeric enzyme, it is conceiv-
able that one GNP was bound to each C-terminus.
Moreover, the distance between two GNPs (reflecting
that between two V-PPase C-termini) was extrapolated
from a simulation plot (Fig. 6E, solid circles). The solid
triangle in Fig. 6E reflects the mean width of GNPs on
the surface of V-PPase, corresponding to a GNP dis-
tance of 2.2 ± 1.4 nm. Our results suggest explicitly that
the two C-termini of V-PPase are in close proximity.
To validate the prediction that the V-PPase C-termini
are adjacent, a TEM analysis was used to directly mea-
sure the distance between two GNPs bound to the C-ter-

mini of purified V-PPase. The TEM image displays the
bound GNPs as solid spheres with a diameter of
2.0 ± 0.2 nm (n = 18) (Fig. 7A). In addition, GNPs
bound to V-PPase C-termini occurred in pairs (Fig. 7B),
indicating the dimeric structure of the enzyme. The his-
togram showing the distribution of distances between
GNP pairs observed from the TEM image yields a mean
distance of 1.9 ± 0.8 nm (Fig. 7C), concurring with the
result generated by AFM analysis of GNP-labeled
V-PPase (Fig. 6E; distance = 2.2 ± 1.4 nm). The slight
fluctuation in distances between GNP pairs most likely
arose from the flexibility of the V-PPase C-termini. For
instance, the shorter distance observed indicates two
closed GNPs on the C-termini of the enzyme. In con-
trast, the longer distance indicates a probable extension
of the C-termini of V-PPase. Verification of these possi-
bilities requires further investigation.
The C-terminus of V-PPase has been determined to
be relatively conserved among various plant V-PPases,
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4388 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
and is presumed to be proximal to the catalytic site
[32]. In addition, the importance of the V-PPase C-
terminus in sustaining enzymatic and proton-translo-
cating function and for indirect regulation of K
+
binding has been demonstrated [9]. Moreover, inter-
subunit interactions of V-PPase are critical for proper

enzyme function [10], suggesting that the interface
between the two subunits may participate in enzy-
matic and proton-pumping reactions. In the present
study, AFM measurements and single nanoparticle
analysis using TEM further demonstrated that the
two C-termini of V-PPase homodimers are approxi-
mately 1.9–2.2 nm apart. In conclusion, our study
provides high-resolution images of single V-PPase
molecules within a membrane, allowing analysis of
the architecture, size and structure of V-PPase in a
physiologically relevant environment. We propose a
working model in which the proton channel lies at
the interface between the C-termini of the V-PPase
homodimer (Fig. 8).
Fig. 6. AFM analysis of Ni
2+
–nitrilotriacetic
acid GNPs bound to the C-terminal 6·His
tag of V-PPase. (A) Lipid bilayer reconsti-
tuted in the presence of Ni
2+
–nitrilotriacetic
acid GNP-bound V-PPase. (B) High-resolu-
tion image of individual dimeric structures of
V-PPase labeled with GNPs in reconstituted
membrane. Solid circle, GNP-bound V-PPase
protrusion on the luminal side of the recon-
stituted membrane; dotted circle, V-PPase
protrusion lacking a GNP label on the lumi-
nal side of the membrane. (C) Histogram of

the protrusion heights determined using the
AFM image in (A). (D) Histogram of the pro-
trusion widths determined using the AFM
image in (A). (E) Simulation of potential
V-PPase protrusion widths based on dis-
tances between GNP pairs bound to the
V-PPase C-termini. Solid rhombus, predicted
protrusion width based on a single GNP
molecule bound to the C-terminus of
V-PPase; solid circle, predicted protrusion
widths based on the distance between two
GNP molecules bound to V-PPase C-termini;
solid triangle, actual protrusion width of
GNP-bound V-PPase determined by AFM.
Data represent the mean ± SD.
T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4389
Experimental procedures
Cloning, expression, and purification
The mung bean (Vigna radiata L.) V-PPase cDNA (VPP;
accession number P21616) [33] was cloned into the yeast
expression vector pYES2 (Invitrogen, Carlsbad, CA, USA),
and the two synthesized oligonucleotides P
his
(5¢-CCTCG
AGCCATCATCATCATCATCATTAGGGCCGCATCAT
GTAATTAGTTATGT-3¢) and P
MluI

(5¢-GTACACGCG
TCTGATCAG-3¢) were inserted into the 3¢-end of the
pYES2–VPP plasmid to generate the V-PPase–(His)
6
tail.
The pYES2–VPP–(His)
6
cDNA was transferred into the
Saccharomyces cerevisiae strain BJ2168 (MATa, prc-407,
prb1-1122, pep4-3, leu2, trp1, ura3, GAL) according to the
method described previously [34]. The yeast microsomal
membranes enriched in 6 · His-tagged V-PPase were pre-
pared as described by Kim et al. [35], with minor modifica-
tions. Finally, V-PPase-enriched membrane fractions were
resuspended in the storage buffer [10 mm Tris/HCl (pH
7.6) and 10% (w/v) glycerol] and stored at )70 °C for fur-
ther use. The V-PPase (1 mgÆmL
)1
)-enriched microsomal
membrane fraction was solubilized in an extraction buffer
[10 mm Tris/HCl (pH 8.0), 400 mm KCl, 15% (w/v) glyc-
erol, 1 mm phenylmethanesulfonyl fluoride, 0.1% (w/v)
n-dodecyl-b-d-maltoside (DDM)] by adding the detergent
DDM dropwise, to a final concentration of 6 mgÆmL
)1
,
and gently stirred for 30 min on ice. The solution was
diluted with the extraction buffer described above three-
fold to five-fold, and unsolubilized materials were removed
by ultracentrifugation at 75 000 g at 4 °C for 1 h. The

supernatant was incubated with Ni
2+
–nitrilotriacetic acid
beads prewashed with the extraction buffer for 1 h. The
Ni
2+
–nitrilotriacetic acid beads were injected into the
empty column and eluted at a flow rate of 0.5 mLÆmin
)1
with the elution medium [10 mm Tris/HCl (pH 8.0), 15%
(w/v) glycerol, 10 mm b-mercaptoethanol, 1 mm phen-
ylmethanesulfonyl fluoride, 0.1% (w/v) DDM] with a step
gradient of 20, 40, 60 and 250 mm imidazole, respectively.
The fractions with highest PP
i
hydrolysis activity at
250 mm imidazole were pooled and dialyzed against med-
ium containing 10 mm Tris/HCl (pH 8.0), 15% (w/v) glyc-
erol, and 0.1% (w/v) Triton X-100, and then stored at
)70 °C for further studies.
Fig. 7. TEM analysis of Ni
2+
–nitrilotriacetic acid GNP-labeled
V-PPase. (A) TEM image of Ni
2+
–nitrilotriacetic acid GNP-labeled
purified V-PPase. (B) A gallery of zoomed images for the GNP pairs
labelled on purified V-PPases. (C) Histogram of the distances
between GNP pairs determined using the TEM image in (A).
Fig. 8. A working model of V-PPase. The distance between

C-termini of V-PPase is approximately 2.2 nm.
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4390 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
Enzyme assay and protein determination
PP
i
hydrolytic activity was measured as the release of P
i
from
PP
i
in a reaction medium [30 m m Tris/Mes (pH 8.0), 1 mm
MgSO
4
, 0.5 mm NaF, 50 mm KCl, 1 mm PP
i
, 1.5 lgÆmL
)1
gramicidin D, 20–30 lgÆmL
)1
microsome protein] at 37 °C
for 15–20 min. The rate of pyrophosphate hydrolysis is lin-
ear with respect to the concentration of V-PPase and reac-
tion time under these conditions (data not shown). After
incubation, the reaction was terminated with a stop solution
[1.7% (w/v) ammonium molybdate, 2% (w/v) SDS, 0.02%
(w/v) 1-amino-2-naphthol-4-sulfonic acid]. The released P
i

was determined spectrophotometrically as described previ-
ously [36,37]. The protein concentration was calculated by a
modified Bradford method with BSA as the standard [38].
Measurement of proton translocation
Proton translocation was measured as the initial rate of flu-
orescence quenching of acridine orange (excitation wave-
length, 495 nm, emission wavelength, 530 nm) as described
previously [34,39–41]. The reaction mixture for proton
translocation contained 5 mm Tris/HCl (pH 8.0), 1 mm
EGTA/Tris (pH 7.6), 400 mm glycerol, 100 mm KCl,
1.3 mm MgSO
4
,5lm acridine orange, and 100 lgÆ mL
)1
microsomes. The reaction was initiated by adding 1 mm
sodium pyrophosphate (pH 7.6). The initial rate of fluores-
cence quenching was calculated as the proton transport
activity [34,39–41]. The ionophore, gramicidin D
(5 lgÆmL
)1
), was then included at the end of each assay to
confirm the integrity of the membrane.
SDS/PAGE and western analysis
SDS/PAGE was performed according to Laemmli [42].
Denatured proteins were subjected to SDS/PAGE on a
Phast System (Pharmacia, Uppsala, Sweden) with a 12.5%
(w/v) polyacrylamide PhastGel. The gels were stained with
Coomassie Blue or electrotransferred to a poly(vinylidene
difluoride) membrane by using semidry electrotransblotting
apparatus (Nova Blot, Amersham Pharmacia Biotech,

Piscataway, NJ, USA). The blots were incubated with the
rabbit polyclonal antibody raised against the MAP (Mito-
gen-activated protein kinase)-conjugated synthetic peptide
of the sequence KVERNIPEDDPRNPA, which corre-
sponds to positions 261–275 of the substrate-binding
domain of mung bean V-PPase. Bands of immunoblots
were visualized using a chemiluminescence kit (New
England Nuclear, Boston, MA, USA), according to the
manufacturer’s recommendations.
Reconstitution of V-PPase into the lipid bilayer
Purified V-PPase was reconstituted into the lipid bilayer
according to the protocol described by Sato et al. [13], with
minor modifications. One hundred milligrams of soybean
phosphatidylcholine and 20 mg of cholesterol were dissolved
in 1 mL of chloroform. The lipid mixture was dried in a vac-
uum chamber, and then suspended in 1 mL of suspension
medium [0.25 m sorbitol, 1 mm MgSO
4
, 0.1 mm EGTA,
2mm dithiothreitol, 10 mm Tricine-Na (pH 7.5)]. The sus-
pension was sonicated in a bath-type sonicator for 5 min at
4 °C. The sonicated lipid mixture (15 lL) was added to 1 mL
of 50 lgÆmL
)1
V-PPase solution, and SM-2 Bio-Beads (Bio-
Rad Laboratories, Hercules, CA, USA) were added to the
mixture at 0.25 mgÆlL
)1
. The mixture was stored for 1 h on
ice with gentle agitation, and the beads were then removed by

filtration through a paper filter. The mixture was applied to a
Sephadex G-50 column equilibrated with 0.25 m sorbitol,
10 mm Tricine-Na (pH 7.5), 1 mm MgSO
4
, 0.1 mm EGTA
and 2 mm dithiothreitol to remove NaCl and glycerol. The
proteoliposome fraction was then frozen in liquid nitrogen,
thawed on ice, and sonicated for 20 s at 4 °C in a bath-type
sonicator. The proteoliposomes thus obtained were immedi-
ately used for measurement of proton-translocating activity.
Immunofluorescence microscopy
V-PPase proteoliposomes were prepared on glass coverslips
and then fused into the lipid bilayer. After being washed with
NaCl/P
i
, the coverslips were placed in blocking solution
[NaCl/P
i
containing 3% (w/v) BSA] for 30 min at room tem-
perature. Samples were then incubated with mouse monoclo-
nal antibody against the C-terminal 6·His tag of V-PPase in
NaCl/P
i
(1 lgÆmL
)1
) for 2 h at room temperature. After
being rinsed with NaCl/P
i
, samples were subsequently incu-
bated with carboxymethylindocyanine 3-coupled goat anti-

(mouse IgG) as secondary antibody (1 lgÆ mL
)1
) for 2 h at
room temperature [43]. Samples were then washed again with
NaCl/P
i
. Fluorescence images were captured using an Olym-
pus BX51 microscope with a 100· oil lens (Olympus, Tokyo,
Japan). Immunofluorescence images were collected with the
green channel filter set (excitation wavelength, 525 nm; emis-
sion wavelength, 585 nm) for V-PPases. Bilayers reconsti-
tuted under several conditions were also imaged as controls.
Immunolabeling of reconstituted V-PPase
The lipid bilayer reconstituted with V-PPases on mica was
incubated for 2 h at room temperature with a mouse mono-
clonal antibody against the C-terminal 6·His tag of
V-PPase (1 lgÆmL
)1
) and then washed twice with NaCl/P
i
.
Finally, the specimen was imaged by AFM.
Ni
2+
–nitrilotriacetic acid GNP labeling of
reconstituted V-PPase
The lipid bilayer reconstituted with V-PPases was incubated
for 2 h at room temperature with Ni
2+
–nitrilotriacetic acid

T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4391
Nanogold solution (Nanoprobes, NY, USA), washed twice
with NaCl/P
i
, and then subjected to AFM imaging.
AFM
AFM was performed in the contact mode using a Nano-
scope IIIa Multimode atomic force microscope equipped
with an E-type scanner (Digital Instruments, Santa Bar-
bara, CA, USA) and a Picoplus instrument (Molecular
Imaging, MI, USA). Muscovite mica (Electron Microscopy
Sciences, Hatfield, PA, USA) was freshly cleaved and
immediately covered with adsorption buffer [10 mm Tris/
HCl (pH 7.8), 300 mm KCl, 25 mm MgCl
2
]. Subsequently,
10 lgÆmL
)1
of protein solution was dropped onto the mica
surface. After 1 h, the sample was rinsed with imaging buf-
fer [10 mm Tris/HCl (pH 7.8), 150 mm KCl]. V-shaped sili-
con nitride (Si
3
N
4
) cantilevers with spring constants of
0.08 NÆm

)1
(Olympus, Tokyo, Japan) were used for all
AFM image scanning. The set point was adjusted to the
highest setting without significant noise to minimize the
force applied to the sample. All AFM imaging of lipid
bilayers with/or without reconstituted V-PPase was
performed in the imaging buffer at room temperature
(22–23 °C). All images were captured at 512 · 512 pixel
resolution and processed using nanoscope III software
(Digital Instruments, Santa Barbara, CA, USA) and spip
software (Scanning Probe Image Processor; Image Metrol-
ogy, Lyngby, Denmark). In general, AFM images were
low-pass filtered, and single protein images were further
passed through an additional Gaussian filter to reduce pixi-
lation. Sizes and statistics of purified and reconstituted pro-
teins were obtained from 20–50 protrusion features for each
molecule in the height image.
Molecular volume measurements of membrane
proteins
The volume of a single membrane protein or protein sub-
structure (V
prot
) was calculated using the volume equation
for a sphere [V
m
= (4/3)pr
3
]. In addition, the volume of a
single soluble or purified protein was calculated by regard-
ing the molecule as a segment of a sphere, using Eqn (1)

[29]:
V
s
¼ðph=6Þð3r
2
þ h
2
Þð1Þ
where V
s
is the molecular volume, and h and r are the
height and the radius (half of the measured width) of the
protein, respectively.
In addition, molecular volume based on molecular mass
was calculated using the Eqn (2):
M
o
¼ðN
A
V
prot
Þ=ðV
1
þ dV
2
Þð2Þ
where N
A
is the Avogadro constant (6.022 · 10
23

Æmol
)1
), V
1
is the partial specific volume of the protein (0.74 cm
3
Æg
)1
),
V
2
is the specific volume of water (1 cm
3
Æg
)1
), and d is a
factor describing the extent of hydration for air-dried pro-
teins (0.4 mol H
2
O/mol protein) [44].
TEM image analysis
Twenty microliters of C-terminal 6·His tag of V-PPase
(10 lgÆmL
)1
) was incubated with 5 lL (0.01 nm)ofNi
2+
–
nitrilotriacetic acid Nanogold for 24 h at 4 °C. Samples
were then centrifuged at 13 400 · g for 1 min. The speci-
mens were examined with a Philips Tecnai F20 high-resolu-

tion transmission electron microscope (Fa. Philips,
Eindhoven, the Netherlands) operating at 200 keV.
Acknowledgements
This work was supported by grants from National
Science Council, Republic of China: NSC 96-2627-
M-007-003 and NSC 97-2627-M-007-003 to R. L. Pan,
NSC 96-2627-M-009-001 to S. K. Fan, NSC 96-2627-
M-007-004 to C. C. Fu, and NSC 96-2627-M-007-003
to F. G. Tseng.
References
1 Maeshima M (2000) Vacuolar H
+
-pyrophosphatase.
Biochim Biophys Acta 1465, 37–51.
2 Tzeng CM, Yang CY, Yang SJ, Jiang SS, Kuo SY,
Hung SS, Ma JT & Pan RL (1996) Subunit
structure of vacuolar proton pyrophosphatase as
determined by radiation inactivation. Biochem J 316,
143–147.
3 Drozdowicz YM & Rea PA (2001) Vacuolar proton-
pyrophosphatases: from the evolutionary backwaters
into the mainstream. Trends Plant Sci 6, 206–211.
4 Rea PA, Kim Y, Sarafian V, Poole RJ, Davies JM &
Sanders D (1992) Vacuolar H
+
-translocating pyrophos-
phatases: a new category of ion translocase. Trends
Biochem Sci 17, 348–353.
5 Gordon-Weeks R, Steele SH & Leigh RA (1996) The
role of magnesium, pyrophosphate, and their complexes

as substrates and activators of the vacuolar H
+
-pump-
ing inorganic pyrophosphatase (studies using ligand
protection from covalent inhibitors). Plant Physiol 111,
195–202.
6 Walker RR & Leigh RA (1981) Mg
2+
-dependent, cat-
ion-stimulated inorganic pyrophosphatase associated
with vacuoles isolated from storage roots of red beet
(Beta vulgaris L.). Planta 153, 150–155.
7 Maeshima M (1991) H
+
-translocating inorganic pyro-
phosphatase of plant vacuoles: inhibition by Ca
2+
,
stabilization by Mg
2+
and immunological comparison
with other inorganic pyrophosphatases. Eur J Biochem
196, 11–17.
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4392 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS
8 Baykov AA, Dubnova EB, Bakuleva NP, Evtushenko
OA, Zhen RG & Rea PA (1993) Steady-state
kinetics of substrate hydrolysis by vacuolar

H
+
-pyrophosphatase. A simple three-state model.
FEBS Lett 327, 199–202.
9 Lin HH, Pan YJ, Hsu SH, Van RC, Hsiao YY, Chen
JH & Pan RL (2005) Deletion mutation analysis on
C-terminal domain of plant vacuolar H
+
-pyrophospha-
tase. Arch Biochem Biophys 442, 206–213.
10 Yang SJ, Ko SJ, Tsai YR, Jiang SS, Kuo SY, Hung
SH & Pan RL (1998) Subunit interaction of vacuolar
H
+
-pyrophosphatase as determined by high hydrostatic
pressure. Biochem J 331, 395–402.
11 Maeshima M (1990) Oligomeric structure of H
+
-trans-
locating inorganic pyrophosphatase of plant vacuoles.
Biochem Biophys Res Commun 168, 1157–1162.
12 Sarafian V, Potier M & Poole RJ (1992) Radiation-
inactivation analysis of vacuolar H
+
-ATPase and H
+
-
pyrophosphatase from Beta vulgaris L. Functional sizes
for substrate hydrolysis and for H+ transport. Biochem
J 283, 493–497.

13 Sato MH, Kasahara M, Ishii N, Homareda H, Matsui
H & Yoshida M (1994) Purified vacuolar inorganic
pyrophosphatase consisting of a 75-kDa polypeptide
can pump H
+
into reconstituted proteoliposomes.
J Biol Chem 269, 6725–6758.
14 Lo
´
pez-Marque
´
s RL, Pe
´
rez-Castin
˜
eira JR, Buch-Peder-
sen JB, Marco S, Rigaud JL, Palmaren MG & Serrano
A (2005) Large-scale purification of the proton pumping
pyrophosphatase from Thermotoga maritina:a
‘Hot-Solve’ method for isolation of recombinant
thermophilic membrane proteins. Biochim Biophys
Acta 1716, 69–76.
15 Dufrene YF & Lee GU (2000) Advances in the charac-
terization of supported lipid films with the atomic force
microscope. Biochim Biophys Acta 1509, 14–41.
16 Rinia HA & de Kruijff B (2001) Imaging domains in
model membranes with atomic force microscopy. FEBS
Lett 504, 194–199.
17 Yuan C & Johnston LJ (2001) Atomic force microscopy
studies of ganglioside GM1 domains in phosphatidyl-

choline and phosphatidylcholine/cholesterol bilayers.
Biophys J 81, 1059–1069.
18 Slade A, Luh J, Ho S & Yip CM (2002) Single molecule
imaging of supported planar lipid bilayer-reconstituted
human insulin receptors by in situ scanning probe
microscopy. J Struct Biol 137, 283–291.
19 Saslowsky DE, Lawrence J, Ren X, Brown DA,
Henderson RM & Edwardson JM (2002) Placental
alkaline phosphatase is efficiently targeted to rafts in
supported lipid bilayers. J Biol Chem 277, 26966–26970.
20 Milhiet PE, Giocondi MC, Baghdadi O, Ronzon F,
Roux B & LeGrimellec C (2002) Spontaneous insertion
and partitioning of alkaline phosphatase into model
lipid rafts. EMBO Rep 3, 485–490.
21 Liu LN, Aartsma TJ & Frese RN (2008) Dimers of
light-harvesting complex 2 from Rhodobacter sphaero-
ides characterized in reconstituted 2D crystals with
atomic force microscopy. FEBS J 275, 3157–3166.
22 Bahatyrova S, Frese RN, Siebert CA, Olsen JD, van
Der Werf KO, van Grondelle R, Niederman RA,
Bullough PA, Otto C & Hunter CN (2004) The native
architecture of a photosynthetic membrane. Nature 430,
1058–1062.
23 Fotiadis D, Liang Y, Filipek S, Saperstein DA, Engel
A & Palczewski K (2003) Atomic-force microscopy:
rhodopsin dimers in native disc membranes. Nature
421, 127–128.
24 Schneider SW, La
¨
rmer J, Henderson RM & Oberleith-

ner H (1998) Molecular weights of individual proteins
correlate with molecular volumes measured by atomic
force microscopy. Pflu
¨
gers Arch 435, 362–367.
25 Geisse NA, Cover TL, Henderson RM & Edwardson
JM (2004) Targeting of Helicobacter pylori vacuolating
toxin to lipid raft membrane domains analysed by
atomic force microscopy. Biochem J 381, 911–917.
26 Gallova
´
J, Uhrı
´
kova
´
D, Islamov A, Kuklin A &
Balgavy
´ P (2004) Effect of cholesterol on the bilayer
thickness in unilamellar extruded DLPC and DOPC
liposomes. Gen Physiol Biophys 23, 113–128.
27 Ho
¨
rber JKH & Miles MJ (2003) Scanning probe evolu-
tion in biology. Science 302, 1002–1005.
28 Engel A & Mu
¨
ller DJ (2000) Observing single biomole-
cules at work with the atomic force microscope. Nat
Struct Biol 7, 715–718.
29 Schneider S, Folprecht G, Krohne G & Oberleithner H

(1995) Immunolocalization of lamins and nuclear pore
complex proteins by atomic force microscopy. Pflu
¨
gers
Arch 430, 795–801.
30 Rapp M, Seppa
¨
la
¨
S, Granseth E & von Heijne G (2007)
Emulating membrane protein evolution by rational
design. Science 315, 1282–1284.
31 Brekke OH, Michaelsen TE & Sandlie I (1995) The
structural requirements for complement activation by
IgG: does it hinge on the hinge? Immunol Today 16,
85–90.
32 Takasu A, Nakanishi Y, Yamauchi T & Maeshima M
(1997) Analysis of the substrate binding site and car-
boxyl terminal region of vacuolar H
+
-pyrophosphatase
of mung bean with peptide antibodies. J Biochem 122,
883–889.
33 Hung SH, Chiu SJ, Lin LY & Pan RL (1995) A cDNA
clone encoding the A subunit of the vacuolar H
+
-AT-
Pase from etiolated mung bean seedlings. Plant Physiol
109, 1125–1127.
34 Hsiao YY, Van RC, Hung SH, Lin HH & Pan RL

(2004) Roles of histidine residues in plant vacuolar H
+
-
pyrophosphatase. Biochim Biophys Acta 1608 , 190–199.
35 Kim EJ, Zhen RG & Rea PA (1995) Site-directed muta-
genesis of vacuolar H
+
-pyrophosphatase: necessity of
T H. Liu et al. Adjacent C-termini of dimeric H
+
-pyrophosphatase
FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS 4393
Cys634 for inhibition by maleimides but not catalysis.
J Biol Chem 270, 2630–2635.
36 Fiske CH & Subbarow Y (1925) The colorimetric deter-
mination of phosphorus. J Biol Chem 66, 375–400.
37 Wang MY, Lin YH, Chow WM, Chung TP & Pan RL
(1989) Purification and characterization of tonoplast
ATPase from etiolated mung bean seedlings. Plant
Physiol 90, 475–481.
38 Bradford M (1976) A rapid and sensitive method for
the quantitation of microgram quantities of protein
utilizing the principle of protein dye binding. Anal
Biochem 72, 248–254.
39 Rea PA & Turner JC (1990) Tonoplast adenosine
triphosphatase and inorganic pyrophosphatase. In
Methods in Plant Biochemistry, vol. 3 (Lea PJ ed),
pp 385–405. Academic Press, London.
40 Rottenberg H (1979) The measurement of membrane
potential and deltapH in cells, organelles, and vesicles.

In Methods in Enzymology, vol. 55 (Fleischer S &
Packer L eds), pp 547–569. Academic Press, New York.
41 Clerc S & Barenholz Y (1998) A quantitative
model for using acridine orange as a
transmembrane pH gradient probe. Anal Biochem
259, 104–111.
42 Laemmli UK (1970) Cleavage of structural proteins
during the assembly of the head of bacteriophage T4.
Nature (Lond) 222 , 680–685.
43 Lin H, Zhu YJ & Lal R (1999) Amyloid beta protein
(1–40) forms calcium-permeable Zn
2+
-sensitive
channel in reconstituted lipid vesicles. Biochemistry 38,
11189–11196.
44 Edstrom RD, Meinke MH, Yang R, Yang X, Elings V
& Evans DF (1990) Direct visualization of phosphory-
lase–phosphorylase kinase complexes by scanning
tunneling and atomic force microscopy. Biophys J 58,
1437–1448.
Adjacent C-termini of dimeric H
+
-pyrophosphatase T H. Liu et al.
4394 FEBS Journal 276 (2009) 4381–4394 ª 2009 The Authors Journal compilation ª 2009 FEBS