doi:10.1016/j.jmb.2005.12.041

J. Mol. Biol. (2006) 357, 73–81

Electron Cryomicroscopic Visualization of PomA/B
Stator Units of the Sodium-driven Flagellar Motorin Liposomes
Koji Yonekura1,2*, Toshiharu Yakushi3, Tatsuo Atsumi3
Saori Maki-Yonekura2 , Michio Homma3 and Keiichi Namba1,2
1

Graduate School of Frontier
Biosciences, Osaka University
1-3, Yamadaoka, Suita, Osaka
565-0871, Japan
2
Dynamic NanoMachine
Project, ICORP, JST, 1-3,
Yamadaoka, Suita, Osaka
565-0871, Japan
3

Division of Biological Science
Graduate School of Science
Nagoya University, Chikusa-ku
Nagoya 464-8602, Japan

A motor protein complex of the bacterial flagellum, PomA/B from Vibrio
alginolyticus, was reconstituted into liposomes and visualized by electron
cryomicroscopy. PomA/B is a sodium channel, composed of two
membrane proteins, PomA and PomB, and converts ion flux to the rotationof the flagellar motor. Escherichia coli and Salmonella have a homolog called
MotA/B, which utilizes proton instead of sodium ion. PomB and MotB
have a peptidoglycan-binding motif in their C-terminal region, andtherefore PomA/B and MotA/B are regarded as the stator. Energy filtering
electron cryomicroscopy enhanced the image contrast of the proteinsreconstituted into liposomes and showed that two extramembrane
domains with clearly different sizes stick out of the lipid bilayers on
opposite sides. Image analysis combined with gold labeling and deletion of
the peptidoglycan-binding motif revealed that the longer one, w70 Å long,is likely to correspond to the periplasmic domain, and the other, about halfsize, to the cytoplasmic domain.
q 2005 Elsevier Ltd. All rights reserved.

*Corresponding author

Keywords: bacterial flagellum; flagellar motor; stator; liposome; energy
filtering electron cryomicroscopy

Introduction
The bacterial flagellum is a large macromolecularcomplex, composed of about 25 different proteins. It
is used for bacterial motility by rotating the flagellar
filaments, which grow to 10–15 mm long. The rotary
motor at the base of the filament drives its rotation.
The motor structure called the flagellar basal bodycrosses both the cytoplasmic and outer membrane
and continues to the extracellular structure called
the hook and the filament.1–4 The motor is poweredby the electrochemical potential of specific ions
Present addresses: K. Yonekura and S. Maki-Yonekura,
The W. M. Keck Advanced Microscopy Laboratory,
Department of Biochemistry and Biophysics, Universityof California, San Francisco, 1700, 4th Street, San
Francisco, CA 94143-2532, USA. T. Yakushi, Departmentof Bioscience and Biotechnology, Faculty of Agriculture,
Shinshu University, Minamiminowa, Nagano 399-4598,
Japan.
Abbreviations used: LDL, low-density lipoprotein;
CTF, contrast transfer function.
E-mail address of the corresponding author:
yone@msg.ucsf.edu

across the cytoplasmic membrane. Genetic andphysiological studies of the proton driven motors
of Escherichia coli and Salmonella typhimurium haveshown that two membrane proteins, MotA and
MotB, form a complex responsible for torquegeneration 5–8 and presumably form a proton
channel.9–11 MotA and MotB have four and one
transmembrane helices, respectively. The MotA/B
complex also functions as the stator by being
anchored to the peptidoglycan layer of thecell through a binding motif in the periplasmic
domain of MotB.12,13 Electron micrographs offreeze-fractured samples showed that the stator
complexes appear as a circular array of particles on
the cytoplasmic membrane, surrounding therotor.14,15 For torque generation, MotA/B is thought
to undergo a conformational change coupled to theproton flux while it interacts with FliG, which is acomponent of the rotor and the motor switch
complex. The switch complex, which is made ofthree proteins, FliG, FliM and FliN, forms the C-ring(Figure 1(a)), and is required for torque generation
and regulation of the switching probability of therotation direction of the motor.16–18

0022-2836/$ - see front matter q 2005 Elsevier Ltd. All rights reserved.

74

Figure 1. Schematic diagrams of two types of flagellar
motors and membrane topology of PomA and PomB.
(a) A hypothetical model for the proton-driven motor in
E. coli and Salmonella (left) and the sodium-driven motorin V. alginolyticus (right). The sodium-driven polar
flagellum of Vibrio is sheathed. The energy source forthe flagellar motor rotation is provided by an electrochemical potential gradient across the inner membrane.
Hypothetical functional units of the stator are (MotA)4(MotB)2 and (PomA)4(PomB)2.33–36 MotX and MotY colocalize to the outer membrane and interact with PomA/
B.28,29 IM, inner membrane; PG, peptidoglycan; OM,
outer membrane. (b) Predicted membrane topology of
PomA and PomB. Black columns within the inner
membrane represent the transmembrane regions. Small
white open circles in PomB indicate the positions of threecysteine residues (Cys8, Cys10 and Cys31). Large
extramembrane domains are represented by ellipsoids.

Some bacteria, such as alkalophilic Bacillus andVibrio species use the electrochemical potential of
sodium ions to generate torque.19,20 In Vibrio, suchas Vibrio alginolyticus, Vibrio parahaemolyticus, andVibrio cholerae,19,20 which normally have only one
flagellum at their cellular pole, four proteins,
PomA, PomB, MotX, and MotY, have been identified and shown to be necessary for the rotation of
the polar flagellar motor. The maximum rotational
speed of the sodium-driven motor in V. alginolyticus
is w1700 Hz,21 while that of the proton-driven
motor in E. coli is w300 Hz.22,23 PomA and PomB
are homologous to MotA and MotB, respectively,
whereas paralogs of MotX and MotY24–27 are notfound in E. coli. Although the functions of MotX
and MotY are not yet clear, recent studies show thatthey are located in the outer membrane and interact
with the PomA/B complex.28,29 PomA and PomB
are thought to form a sodium channel complex.

Electron Cryomicroscopy of PomA/B in Liposomes

Channel blockers for mammalian epithelial sodium
ion channels such as amiloride and phenamilspecifically inhibit the rotation of the sodium-type
motor.30,31 MotA/B from E. coli can drive the polar
flagellum of V. cholerae26 and V. alginolyticus32 byproton motive force. Also, a complex of PomA and achimera protein made of PomB and MotB segments
can convert a proton-driven motor into a sodiumdriven one.32 Hence, PomA/B or MotA/B determines ion specificity of the motor, and the two typesof motors must share a similar mechanism for iondriven torque generation. Several studies suggested
that the functional unit is (PomA)4(PomB)2 or(MotA)4(MotB)2.33–36 Figure 1 shows a schematic
drawing of the proton-type motor in E. coli and
Salmonella and the sodium-type motor in Vibrio anda topology diagram of the PomA/B complex.
Despite the importance of its function, thestructural information of the stator complex isvery limited and observation of the structure has
not been reported to date except for the freezefracture views.14,15 Here, we report the observationof a purified PomA/B complex reconstituted into
liposomes by energy filtering electron cryomicroscopy. The image showed that two extramembrane
domains stick out of the lipid bilayers on both sides
but asymmetrically in size. Image analysis combined with gold labeling and deletion of thepeptidoglycan-binding motif allowed domain
assignment of the complex.

Results
Sample preparation
The purification protocol was changed from theprevious one,37 and Chaps, instead of sucrose
monocaprate, was used to solubilize the PomA/B
complex, because Chaps was able to keep thecomplex more stably (T.A. et al., unpublishedresults). Gel-filtration gave a clear single peak at
550 kDa as calibrated with soluble globular proteinsused as standards. This indicates that PomA/B
forms a large complex, but smaller than that formedin sucrose monocaprate (900 kDa).37 SDS-PAGE andimmunoblots revealed that PomA forms a stable
dimer that is resistant to SDS (the second and fifthlane in Figure 2).38 The dimer band disappeared
when the sample solution was dialyzed against
non-detergent buffer solutions for a few days toreconstitute PomA/B into liposomes as shown in
the third and sixth lane in Figure 2. Therefore, thedimer formation might be caused by the detergent
Chaps.
We first observed negatively stained samples ofpurified PomA/B dissolved in detergent solutions
by electron microscopy, but found only aggregationof the proteins. We also examined the same samples
quickly frozen and embedded in vitreous ice,
but we could not see any structure, presumably
because the size of the predicted functional unit
(PomA)4(PomB)2, of which the molecular mass is

75Electron Cryomicroscopy of PomA/B in Liposomes

testing various conditions for reconstitution into
unilamellar liposomes with sizes suitable forelectron cryomicroscopic observation, the protocol
described in Materials and Methods, Sample
preparation, gave good and reproducible results.
Electron cryomicroscopy of proteoliposomes

Figure 2. SDS-PAGE patterns of the purified PomA/B
complex by silver staining. Lanes: 1, marker; 2, PomA/B;
3, PomA/B with bound Nanogold after reconstitution
into liposomes with no reducing agent; 4, the same as lane
3 except that this was developed with a gold staining kit;
5, PomA/BDC,43 the complex of PomA and a truncation
variant of PomB missing the C-terminal 120 residues; and6, PomA/BDC after reconstitution into liposomes with no
reducing agent. Note that the bands of the PomA dimer
disappear and the bands of PomB and PomBDC are
replaced by those of their dimers, respectively, in lanes 3
and 6. The band of the PomA monomer moved faster thanthat of the PomBDC monomer despite that the molecularmass of PomA is larger.

170 kDa,33–36 may be too small to visualize byelectron cryomicroscopy when the complex isdispersed in solution. Also, visualization of themembrane-spanning structure of the complex
would be more desirable. Therefore, reconstitutionof the complex into liposomes was tried. After

We examined proteoliposomes by electronenergy filtering microscopy, which can produce
the image formed by zero-energy loss (zero-loss)
electrons by removing inelastically scattered electrons, which contribute to noises in image formation. Most of the unilamellar liposomes prepared
as described above had the size range from a few
hundreds to a few thousands Å in diameter. Zeroloss images resolved the proteoliposomes significantly clearer than conventional electron cryomicrographs due to reduced noise and increased
amplitude contrast.39–41 This effect was more
remarkable for larger liposomes because they areembedded in thicker ice. The phosphate head groupof lipids in the lipid bilayers showed the highest
contrast in the images even without energy filtering,but the contrast enhancement by zero-loss imaging
was also most prominent, probably because phosphor scatters more electrons inelastically thanhydrogen, carbon, nitrogen and oxygen, and energy
filtering can eliminate those electrons (Figure 3).
The purified PomA/B complex was reconstituted
into liposomes by removal of the detergent by
dialysis and quickly frozen for electron cryomicroscopic observation. There were aggregations, probably derived from PomA/B, but we found in many
liposomes rod-shaped objects sticking out of thelipid bilayer plane on both sides (Figure 3). These
two extramembrane domains were asymmetric inlength but had a similar diameter of w20 Å. The
longer one was w70 Å long and the shorter one wasapproximately one-half of it. There seemed to be no
preferred orientations for insertion of the complex(Figure 4(a)). Although the orientation was different

Figure 3. Zero-loss image of a
PomA/B proteoliposome. (a) Typical electron cryomicrograph of aunilamellar proteoliposome with adiameter of w1000 Å. The lipid
bilayers are clearly visible. Circles
enclose elongated particles sticking
out of the vesicle surface. The bar
represents 500 Å. (b) Zoom-up
view of the particle encircled in
the upper left circle in (a). These
images clearly show that two
extramembrane domains of distinctly different sizes stick out of
the lipid bilayers on both sides.
The longer one is w70 Å long. The
bar represents 100 Å.

76

Electron Cryomicroscopy of PomA/B in Liposomes

Figure 4. Individual images of PomA/B inserted into the lipid bilayers and two-dimensional average. (a) Individual
images. (b) Two-dimensional average of those shown in (a). The bar represents 50 Å. (c) The same image as in (b), but
overlaid with hypothetical domain envelopes.

from image to image, and conformations of themolecules appeared to have variations, 18 images(Figure 4(a)) were averaged roughly based on the
manually picked points (see Materials andMethods). The resulting average is shown in
Figure 4(b), and the hypothetical molecular envelope is drawn in Figure 4(c). The longer domain canbe divided into two parts of similar dimension, aproximal and a distal domain. The density spanning the lipid bilayers shows an asymmetric
distribution, and the one on the side of the shorter
domain looks slightly larger than the other(Figure 4(c)), but this may be due to the clearlyhigher density contrast of the inner leaflet of thebilayers than the outer one (Figure 3). Although theshorter domain sticks out of the outer leaflet in
the image shown in Figure 3(b), the orientation of
the particle is opposite in 13 images out of 18 thatwere used to obtain the average, which puts theshorter domain on the side of inner leaflet in about
two-thirds of the images. However, a different
interpretation could be made based on the size ofthe cytoplasmic domains of PomA as described in
Discussion. The size of the shorter domain in the
average is smaller than those in individual raw
images and probably due to its conformationalvariation.
Gold-labelingTo ensure that the particles we observed above
were the PomA/B complex, a gold label was
introduced. PomB has three free cysteine residues,
two in the N-terminal region (Cys10 and Cys8) and
one in the transmembrane region (Cys31)(Figure 1(b)), whereas PomA has no cysteine. We
used monomaleimido Nanogold with a 1.4 nm gold
particle to label these cysteine residues. Because any

reductants would prevent the reaction and destroy
gold particles, no reducing agent was added to thedialysis buffer. Under this condition, PomB forms a
dimer with a disulfide bond as shown in the thirdand sixth lanes in Figure 2, but it is known that thisbond formation does not affect the motor functionof the motor.36 An LI silver staining kit, which is astaining reagent specific for gold, was used to detectbound Nanogold as shown in the fourth lane in
Figure 2. The band corresponding to the PomB
dimer is clearly stained. The region corresponding
to the smeared band of the PomA monomer and theregion below are also stained, but these areprobably due to non-specifically bound andunbound gold, respectively, as the stain density ismuch weaker compared to the amount of theprotein shown in the third lane.
Individual gold particles are visible at relatively
small defocus, e.g. at w7000 Å (Figure 5(a)), but aremuch harder to see at larger defocus, e.g. a few
mm.42 In contrast, proteins on the surface ofliposomes can be barely seen at small defocus
even by energy filtering (Figure 5(a)). Recording apair of identical views at two different defocus
levels seemed to be a good solution, but in practicethe second shot frequently destroyed the lipid
vesicles, particularly larger ones even at relatively
low dose. Figure 5(b) shows the poor contrast ofbound Nanogold in individual images recorded at adefocus of w25,000 Å. However, averaging of about
ten images clearly revealed the bound gold particle,
as shown in Figure 5(c). The density distribution of
Nanogold is broad due to the rough alignment ofheterogeneous images as described in Materials andMethods, Image analysis. Still, the high-density
region is clearly located on the surface of themembrane on the shorter domain side. Since
Cys31 is predicted to be located near the center of

77Electron Cryomicroscopy of PomA/B in Liposomes

Figure 6. Average of 23 images of PomA/BDC.43 The
longer domain is missing.

side, where the peptidoglycan-binding domain of
PomB is located.
Figure 5. Individual and averaged images of PomA/B
with bound Nanogold in comparison with the one
without Nanogold labeling. (a) Image taken at a relatively
small defocus level (w7000 Å underfocus). (b) Images
taken at w25,000 Å underfocus. (c) Average of 12 images
including those in (b). The contrast of the image isadjusted to show the peak corresponding to the averaged
Nanogold density clearly. (d) The image without Nanogold, as already shown in Figure 4(b) but with its contrast
adjusted to (c) for direct comparison. Arrows in (a) and (c)indicate a single bound Nanogold and the averaged
density, respectively.

the membrane and not on the membrane surface(Figure 1(b)), this is not the one labeled. Since it is
known that Cys31 is not responsible for the dimer
formation,36 either Cys8 or Cys10 must be forming adisulfide bond and the other labeled with Nanogold. Therefore, this result indicates that the side of
the shorter domain is most likely the PomB
N-terminal side.
Deletion of peptidoglycan-binding motif
We also examined a mutant with deletion of the
C-terminal 120 residues of PomB, which includes
the peptidoglycan-binding motif (PomA/BDC).43
This mutant protein was purified in the same way
without any changes in the elution profile and thepeak position in gel-filtration, despite the change inits molecular mass. The purified protein wasreconstituted into liposomes in the same way.
Although it was more difficult to find them inelectron cryomicrographs because of the smallersize, we were able to record their images andaveraged them in the same way as for the fulllength protein complex. Figure 6 shows an averageof 20 images. The particle does not have the large
extramembrane domain, indicating that the side ofthe longer domain is most likely the periplasmic

Discussion
Despite the important roles that PomA/B and
MotA/B play in torque generation as the stator of thebacterial flagellar motor, even their overall shapes
were not known. Here, we report the observation of
the shape and domain arrangement of the PomA/B
complex and locations of domains relative to themembrane bilayers, which was made possible byenergy filtering electron cryomicroscopy of thepurified complex reconstituted into liposomes.
Since conventional negatively stained or iceembedded specimens of the purified complex
dissolved in Chaps solutions did not show any
structural features by electron microscopy, thecomplex was reconstituted into lipid vesicles, andthe proteoliposomes embedded in vitreous ice wereobserved. A similar observation has been reported fora membrane protein, the low-density lipoprotein(LDL) receptor (w115 kDa), which has a large
glycosylated extracellular domain and one transmembrane helix,42 but without energy filtering.
Energy filtering can better resolve details of large
samples such as liposomes embedded in relatively
thick ice and produce images of higher amplitudecontrast, because it removes inelastically scatteredelectrons, which otherwise give high backgroundnoise and increase as the specimen thickness
increases. In particular, the phosphate head groupsof lipids become distinctly clear (Figure 3(a)). Using
energy filtering we were able to identify PomA/B
clearly, even though its extramembrane regions are
smaller than that of the LDL receptor.42
The images of liposomes with reconstituted
PomA/B clearly resolved two extramembrane
domains of different sizes sticking out of the lipid
bilayers on opposite sides. The longer one is w70 Å
long and the shorter one is roughly one-half of it.

78
Although individual images of PomA/B may varyin their orientation, and probably conformation, a
two-dimensional image average further revealed
that the longer domain is divided into two parts.
Nanogold labeling of either Cys8 or Cys10 of PomB
showed that the N-terminal region of PomB is
located within the shorter domain, and in agreement with this, the C-terminal deletion of 120
residues of PomB made the longer domain disappear, indicating that the longer domain contains
the peptidoglycan-binding domain. Although thesize of this C-terminal deletion is less than half ofthe periplasmic domain of PomB (w280 residues),
predicted from the amino acid sequence, deletion of
the peptidoglycan-binding domain might havedestabilized the remaining half of the periplasmic
domain, resulting in smearing out of its density inthe averaged image. Therefore, it is likely that thelonger domain corresponds to the predicted periplasmic domain of PomB. The distance between theouter surface of the cytoplasmic membrane and theinner surface of the outer membranes is w220 Å,and the peptidoglycan layer extends within thisspace with its inner surface approximately 150 Å
apart from the outer surface of the cytoplasmic
membrane.44 The length of the longer domain istherefore not sufficient to reach the peptidoglycan
layer. The peptidoglycan-binding domain, which isabout a half of the whole periplasmic domain of
PomB, may not yet be visualized in the averaged
image due to its flexible orientation relative to therest of PomA/PomB. Or, peptidoglycan may be
extended out of the layer so that the longer domain
can reach, as depicted in Figure 7.

Figure 7. Interpretation of the PomA/B images
obtained here combined with all available information
including the model proposed by Brown et al.48 Charged
residues in the C-terminal domain of FliG are thought tointeract with critical residues in the large cytoplasmic
loop between transmembrane helix 2 and 3 of PomA. The
atomic model of the C-terminal and the middle domainsof FliG is from Brown et al.48 The half cut density map of
the MS ring is from Suzuki et al.49 The position of FliG was
adopted from Francis et al.,44 Brown et al.48 and Suzuki
et al.49 The peptidoglycan layer and its branches are
represented in yellow.

Electron Cryomicroscopy of PomA/B in Liposomes

The major part of the shorter domain sticking
into the cytoplasm would correspond to the long
loop between transmembrane helix 2 and 3 andthe C-terminal chain of PomA (Figure 1(b)),
which are predicted to reside in the cytoplasm.
The loop and the C-terminal chain are w100 and
w50 amino acid residues long, respectively, while
the periplasmic domain of PomB is w280 aminoacid residues. Many studies have suggested that
(PomA)4(PomB)2 is the functional unit.33–36 SDSPAGE also showed that PomB forms a dimer (thethird and sixth lanes in Figure 2). The fact thatno double-headed structure was observed inindividual images of PomA/B suggests that thetwo periplasmic domains of the PomB dimer
are tightly packed together. Assuming that
(PomA)4(PomB)2 is the complex we observed,
the longer domain should contain the two
periplasmic domains of PomB consisting of 560(280!2) residues, and the shorter domain is
supposed to consist of 600 (150!4) residues offour cytoplasmic loops and four C-terminal
chains of PomA. In the averaged image, however,the size of the shorter domain is only half that of
the longer domain. This may partly be due toaveraging out of a significant amount of densityof the cytoplasmic domains of PomA for its
variable conformation and orientation, but it may
also be because a large part of the cytoplasmic
domains is located close to the surface of themembrane, as suggested by the higher density of
the half transmembrane domain on the cytoplasmic side than the periplasmic side
(Figure 4(b)). The cytoplasmic loop of PomA
contains critical amino acid residues that arethought to interact with FliG,45–47 which forms
the switch complex of the flagellar motor andmakes up the C-ring together with FliM and FliN
(Figure 1(a)). The crystal structures of the
C-terminal and middle domains of FliG from
Thermotoga maritima have been solved by X-ray
crystallography, revealing that important charged
residues are lined up on one side of FliG48 so as
to interact with the cytoplasmic loop of PomA or
MotA. In Figure 7, we illustrate our interpretationof the PomA/B image obtained in this study in
combination with all other available information44,49 including the model proposed byBrown et al.48
We observed the shape of the PomA/B complex
reconstituted into liposomes, but still little is known
about its structure. Its high-resolution three-dimensional structure is essential for understanding thecoupling mechanism of the motor torque generation
with ion flux through the channel of the complex.

Materials and Methods
Sample preparation
A bacterial strain of V. alginolyticus, plasmid andgrowth condition were described by Yakushi et al.37

79Electron Cryomicroscopy of PomA/B in Liposomes

A plasmid, pKJ301, which overproduces PomA/B with
six histidine residues at the C terminus of PomB wasused.37 Purification of the PomA/B complex was
carried out as described37 with some modifications.
Briefly, the membrane fraction was dissolved in 2.5%(w/v) Chaps, 20 mM Tris–HCl (pH 8.0), 150 mM KCl,
10% (v/v) glycerol, 1 tablet of Complete EDTA-freeprotease inhibitor cocktail (Roche Applied Science,
Indianapolis, IN) and 20 mM imidazole (pH 8.0) at
room temperature for 40 min. After the suspension wascentrifuged at 100,000g for 30 min, the supernatant wasapplied to a column filled with Ni-NTA agarose(QIAGEN, Valencia, CA) and mixed at 4 8C for 1 h.
Then, the column was washed with the same buffer
except for containing 1% (w/v) Chaps. Then, theconcentration of imidazole in the buffer was increased
stepwise to 400 mM. PomA/B-His6 was well eluted at
50 mM–200 mM imidazole. The eluate was concentrated using a spin concentrator with a molecularmass cutoff of 10,000 Da and applied to a gel-filtration
column, Superdex 200HR (Amersham Biosciences,
Pittsburgh, PA). It was run at a flow rate of 0.5 ml/
min in the same buffer but not containing imidazole.
A fraction at the main peak corresponding tow550 kDa was collected and subjected to the BCA
assay (Pierce, Rockford, IL) for measurement of theprotein concentration. Then, it was mixed withpreformed liposomes made from E. coli polar lipid
extract (Avanti Polar Lipids, Alabaster, AL) at a final
concentration of the lipid of 1 mg/ml and a final ratio
of protein to lipid of w0.4 in the buffer containing 2%
Chaps, and incubated at room temperature for 1 h.
Then, the detergent was removed by dialysis against
20 mM Tris–HCl (pH 6.8) or 20 mM Mes (pH 6.5)
(essential for gold labeling), 150 mM KCl, 10% (v/v)
glycerol and 5 mM MgCl2 at 37 8C overnight. PomB has
free cysteine residues (Figure 1(b), white circles),whereas PomA has none. For gold labeling of cysteine,monomaleimido Nanogold with a 1.4 nm gold particle(Nanoprobes, Yaphank, NY) was added to the proteoliposomes at a final concentration of Nanogold at a few
times excess to the protein, and the samples wereincubated at room temperature for 1 h. Then, they weredialyzed again to remove unbound Nanogold at 4 8C
for a few days. For SDS-PAGE, the samples were run
on 12.5% or 15% (w/v) homogeneous polyacrylamide
gel using the Laemmli buffer system.50 To checkmonomaleimido Nanogold bound to the cysteine
residues, the buffer system contained essentially no
reductant. The gels were developed with an LI Silver
stain kit (Nanoprobes, Yaphank, NY) to detect boundNanogold. Western blot was done using anti-peptide
antibodies against PomA and PomB.51 We also
examined a mutant PomA/B complex with deletionof the C-terminal 120 residues of PomB, which includes
the peptidoglycan-binding motif. This mutant (PomA/
BDC) also has six histidine residues at its C terminus,43
and therefore purification and preparation of its
proteoliposomes were carried out as described above.

Figure 8. Schematic representation of the imagealignment method. Three points (X) were marked
manually at the tips of the longer and shorter extramembrane domains and at the center of the lipid bilayers.
Some of the images were inverted into their mirror
images about the axis approximately normal to the lipid
bilayers (paired thick curves) for alignment and averaging.

accelerating voltage of 200 kV. An energy slit below theelectromagnet was adjusted to select only electrons withenergy loss less than 10 eV. Images were recorded on
Kodak SO163 films (Eastman Kodak, Rochester, NY) at anominal magnification of 70,000! using a low-dose kit.
Defocus levels of w25,000 Å were used, which give thefirst zero of the contrast transfer function (CTF) at w25 Å
resolution.
Image analysis
We selected by eye images that showed extramembrane
domains sticking out of the lipid bilayers on both sides.
The images were digitized with a LeafScan 45 linear CCD
densitometer (LaserSoft Imaging, Longboat Key, FL) at a
step size of 10 mm. Magnification was calibrated with thelayer-line spacing of the straight flagellar filaments53 andthe thickness of the lipid bilayers. Then, three points weremanually marked on each PomA/B image, one on thelonger and one on the shorter extramembrane domain
and one on the center of the lipid bilayers, by using the
SPIDER/WEB package.54 Based on the selected points,
each image was cut out into a small box and the density
scales of the images were adjusted. Most of the PomA/B
images were slightly curved in either one of two
directions. To adjust relative positions of the three points
among the images, some images were converted into theirmirrored ones against a line approximately normal to thelipid plane (Figure 8). This procedure corresponds to thechange of the direction of view or image projection by
1808. Then, all the images were aligned and averaged
based on the two points on the longer and shorter
domains. Since it was a manual-based alignment, toassure the reliability and reproducibility of the result, thewhole procedure from marking points to the average wasrepeated three times to make sure the resulting 2D
averages do not show any significant difference. The
numbers of averaged images are 18 for the intact PomA/
B complex, 12 for the one with gold labeling, and 23 forthe deletion mutant.

Electron cryomicroscopy
A few microlitres of the proteoliposome solution wasapplied to a holey carbon grid and rapidly frozen inliquid ethane. These sample grids were mounted on an
Oxford CT3500 cryo-holder (Gatan, Pleasanton, CA) andexamined with a HITACHI EF-2000 electron microscope(Hitachi, Tokyo, Japan), equipped with a g-type energy
filter52 and a cold field emission gun operated at an

Acknowledgements
We thank Masaru Kojima for his help in harvesting bacteria and preparation of the membrane
fraction, and Naoko Hattori for making deletionmutants.

80

References
1. Macnab, R. M. (2003). How bacteria assemble flagella.
Annu. Rev. Microbiol. 57, 77–100.
2. Blair, D. F. (2003). Flagellar movement driven byproton translocation. FEBS Letters, 545, 86–95.
3. Yonekura, K., Maki-Yonekura, S. & Namba, K. (2002).
Growth mechanism of the bacterial flagellar filament.
Res. Microbiol. 153, 191–197.
4. Minamino, T. & Namba, K. (2004). Self-assembly andtype III protein export of the bacterial flagellum.
J. Mol. Microbiol. Biotechnol. 7, 5–17.
5. Dean, G. E., Macnab, R. M., Stader, J., Matsumura, P.
& Burks, C. (1984). Gene sequence and predicted
amino acid sequence of the motA protein, amembrane-associated protein required for flagellar
rotation in Escherichia coli. J. Bacteriol. 159, 991–999.
6. Zhou, J., Fazzio, R. T. & Blair, D. F. (1995). Membrane
topology of the MotA protein of Escherichia coli. J. Mol.
Biol. 251, 237–242.
7. Stader, J., Matsumura, P., Vacante, D., Dean, G. E. &Macnab, R. M. (1986). Nucleotide sequence of the
Escherichia coli motB gene and site-limited incorporation of its product into the cytoplasmic membrane.
J. Bacteriol. 166, 244–252.
8. Chun, S. Y. & Parkinson, J. S. (1988). Bacterial motility:
membrane topology of the Escherichia coli MotB
protein. Science, 239, 276–278.
9. Blair, D. F. & Berg, H. C. (1990). The MotA protein of
E. coli is a proton-conducting component of theflagellar motor. Cell, 60, 439–449.
10. Stolz, B. & Berg, H. C. (1991). Evidence for interactions between MotA and MotB, torque-generating
elements of the flagellar motor of Escherichia coli.
J. Bacteriol. 173, 7033–7037.
11. Tang, H., Braun, T. F. & Blair, D. F. (1996). Motility
protein complexes in the bacterial flagellar motor.
J. Mol. Biol. 261, 209–221.
12. De Mot, R. & Vanderleyden, J. (1994). The C-terminal
sequence conservation between OmpA-related outermembrane proteins and MotB suggests a common
function in both Gram-positive and Gram-negative
bacteria, possibly in the interaction of these domains
with peptidoglycan. Mol. Microbiol. 12, 333–334.
13. Koebnik, R. (1995). Proposal for a peptidoglycanassociating alpha-helical motif in the C-terminal
regions of some bacterial cell-surface proteins. Mol.
Microbiol. 16, 1269–1270.
14. Coulton, J. W. & Murray, R. G. (1978). Cell envelope
associations of Aquaspirillum serpens flagella.
J. Bacteriol. 136, 1037–1049.
15. Khan, S., Dapice, M. & Reese, T. S. (1988). Effects ofmot gene expression on the structure of the flagellar
motor. J. Mol. Biol. 202, 575–584.
16. Sockett, H., Yamaguchi, S., Kihara, M., Irikura, V. &Macnab, R. M. (1992). Molecular analysis of theflagellar switch protein FliM of Salmonella typhimurium. J. Bacteriol. 174, 793–806.
17. Irikura, V. M., Kihara, M., Yamaguchi, S., Sockett, H. &Macnab, R. M. (1993). Salmonella typhimurium fliG andfliN mutations causing defects in assembly, rotation,and switching of the flagellar motor. J. Bacteriol. 175,
802–810.
18. Lloyd, S. A., Tang, H., Wang, X., Billings, S. & Blair, D. F.
(1996). Torque generation in the flagellar motor ofEscherichia coli: evidence of a direct role for FliG but notfor FliM or FliN. J. Bacteriol. 178, 223–231.

Electron Cryomicroscopy of PomA/B in Liposomes

19. Yorimitsu, T. & Homma, M. (2001). NaC-driven
flagellar motor of Vibrio. Biochim. Biophys. Acta, 1505,
82–93.
20. McCarter, L. L. (2001). Polar flagellar motility of theVibrionaceae. Microbiol. Mol. Biol. Rev. 65, 445–462.
21. Magariyama, Y., Sugiyama, S., Muramoto, K.,
Maekawa, Y., Kawagishi, I., Imae, Y. & Kudo, S.
(1994). Very fast flagellar rotation. Nature, 371, 752.
22. Lowe, G., Meister, M. & Berg, H. C. (1987). Rapid
rotation of flagellar bundles in swimming bacteria.
Nature, 325, 637–640.
23. Chen, X. & Berg, H. C. (2000). Torque–speed
relationship of the flagellar rotary motor of Escherichia
coli. Biophys. J. 78, 1036–1041.
24. McCarter, L. L. (1994). MotY, a component of thesodium-type flagellar motor. J. Bacteriol. 176,
4219–4225.
25. McCarter, L. L. (1994). MotX, the channel componentof the sodium-type flagellar motor. J. Bacteriol. 176,
5988–5998.
26. Gosink, K. K. & Häse, C. C. (2000). Requirements forconversion of the NaC-driven flagellar motor of Vibrio
cholerae to the HC-driven motor of Escherichia coli.
J. Bacteriol. 182, 4234–4240.
27. Okabe, M., Yakushi, T., Asai, Y. & Homma, M. (2001).
Cloning and characterization of motX, a Vibrio
alginolyticus sodium-driven flagellar motor gene.
J. Biochem. 130, 879–884.
28. Okabe, M., Yakushi, T., Kojima, M. & Homma, M.
(2002). MotX and MotY, specific components of thesodium-driven flagellar motor, colocalize to the outermembrane in Vibrio alginolyticus. Mol. Microbiol. 46,
125–134.
29. Okabe, M., Yakushi, T. & Homma, M. (2005).
Interactions of MotX with MotY and with the
PomA/PomB sodium ion channel complex of theVibrio alginolyticus polar flagellum. J. Biol. Chem. 280,
25659–25664.
30. Sugiyama, S., Cragoe, E. J., Jr & Imae, Y. (1988).
Amiloride, a specific inhibitor for the NaC-driven
flagellar motors of alkalophilic Bacillus. J. Biol. Chem.
263, 8215–8219.
31. Atsumi, T., Sugiyama, S., Cragoe, E. J., Jr & Imae, Y.
(1990). Specific inhibition of the NaC-driven flagellar
motors of alkalophilic Bacillus strains by the amiloride
analog phenamil. J. Bacteriol. 172, 1634–1639.
32. Asai, Y., Yakushi, T., Kawagishi, I. & Homma, M.
(2003). Ion-coupling determinants of NaC-driven and
HC-driven flagellar motors. J. Mol. Biol. 327, 453–463.
33. Sato, K. & Homma, M. (2000). Functional reconstitution of the NaC-driven polar flagellar motor component of Vibrio alginolyticus. J. Biol. Chem. 275,
5718–5722.
34. Braun, T. F. & Blair, D. F. (2001). Targeted disulfide
cross-linking of the MotB protein of Escherichia coli:
evidence for two HC channels in the stator complex.
Biochemistry, 40, 13051–13059.
35. Kojima, S. & Blair, D. F. (2004). Solubilization andpurification of the MotA/MotB complex of Escherichia
coli. Biochemistry, 43, 26–34.
36. Yorimitsu, T., Kojima, M., Yakushi, T. & Homma, M.
(2004). Multimeric structure of the PomA/PomB
channel complex in the NaC-driven flagellar motorof Vibrio alginolyticus. J. Biochem. 135, 43–51.
37. Yakushi, T., Kojima, M. & Homma, M. (2004).
Isolation of Vibrio alginolyticus sodium-driven flagellar motor complex composed of PomA and PomB
solubilized by sucrose monocaprate. Microbiology, 150,
911–920.

Electron Cryomicroscopy of PomA/B in Liposomes

38. Fukuoka, H., Yakushi, T., Kusumoto, A. & Homma, M.
(2005). Assembly of motor proteins, PomA and PomB,in the NaC-driven stator of the flagellar motor. J. Mol.
Biol. 351, 707–717.
39. Smith, M. F. & Langmore, J. P. (1992). Quantitation ofmolecular densities by cryo-electron microscopy.
Determination of the radial density distribution of
tobacco mosaic virus. J. Mol. Biol. 226, 763–774.
40. Zhu, J., Penczek, P. A., Schröder, R. & Frank, J. (1997).
Three-dimensional reconstruction with contrast
transfer function correction from energy-filtered cryoelectron micrographs: procedure and application to the
70 S Escherichia coli ribosome. J. Struct. Biol. 118, 197–219.
41. Angert, I., Majorovits, E. & Schröder, R. R. (2000). Zeroloss image formation and modified contrast transfer
theory in EFTEM. Ultramicroscopy, 81, 203–222.
42. Jeon, H. & Shipley, G. G. (2000). Vesicle-reconstituted
low density lipoprotein receptor. Visualization bycryoelectron microscopy. J. Biol. Chem. 275,
30458–30464.
43. Yakushi, T., Hattori, N. & Homma, M. (2005). Deletion
analysis of the carboxyl-terminal region of the PomB
component of the Vibrio alginolyticus polar flagellar
motor. J. Bacteriol. 187, 778–784.
44. Francis, N. R., Sosinsky, G. E., Thomas, D. &DeRosier, D. J. (1994). Isolation, characterizationand structure of bacterial flagellar motors containing
the switch complex. J. Mol. Biol. 235, 1261–1270.
45. Zhou, J. & Blair, D. F. (1997). Residues of thecytoplasmic domain of MotA essential for torquegeneration in the bacterial flagellar motor. J. Mol. Biol.
273, 428–439.
46. Zhou, J., Lloyd, S. A. & Blair, D. F. (1998). Electrostaticinteractions between rotor and stator in the bacterial
flagellar motor. Proc. Natl Acad. Sci. USA, 95, 6436–6441.

81
47. Yorimitsu, T., Sowa, Y., Ishijima, A., Yakushi, T. &Homma, M. (2002). The systematic substitutions
around the conserved charged residues of thecytoplasmic loop of NaC-driven flagellar motor
component PomA. J. Mol. Biol. 320, 403–413.
48. Brown, P. N., Hill, C. P. & Blair, D. F. (2002). Crystal
structure of the middle and C-terminal domains of theflagellar rotor protein FliG. EMBO J. 21, 3225–3234.
49. Suzuki, H., Yonekura, K. & Namba, K. (2004).
Structure of the rotor of the bacterial flagellar
motor revealed by electron cryomicroscopy andsingle-particle image analysis. J. Mol. Biol. 337,
105–113.
50. Laemmli, U. K. (1970). Cleavage of structural proteinsduring the assembly of the head of bacteriophage T4.
Nature, 227, 680–685.
51. Yorimitsu, T., Sato, K., Asai, Y., Kawagishi, I. &Homma, M. (1999). Functional interaction between
PomA and PomB, the NaC-driven flagellar motor
components of Vibrio alginolyticus. J. Bacteriol. 181,5103–5106.
52. Taya, S., Taniguchi, Y., Nakazawa, E. & Usukura, J.
(1996). Development of g-type energy filtering TEM.
J. Electron Microsc. 45, 307–313.
53. Maki-Yonekura, S., Yonekura, K. & Namba, K. (2003).
Domain movements of HAP2 in the cap-filamentcomplex formation and growth process of thebacterial flagellum. Proc. Natl Acad. Sci. USA, 100,
15528–15533.
54. Frank, J., Radermacher, M., Penczek, P., Zhu, J., Li, Y.,
Ladjadj, M. & Leith, A. (1996). SPIDER and WEB
processing and visualization of images in 3D electron
microscopy and related fields. J. Struct. Biol. 116,
190–199.

Edited by Sir A. Klug
(Received 13 September 2005; received in revised form 7 November 2005; accepted 9 December 2005)
Available online 4 January 2006