3733
Journal of Cell Science 112, 3733-3745 (1999)
Printed in Great Britain © The Company of Biologists Limited 1999
JCS0790
A key function of non-planar membranes and their associated microtubular
ribbons in contractile vacuole membrane dynamics is revealed by
electrophysiologically controlled fixation of Paramecium
Takashi Tominaga*, Yutaka Naitoh and Richard D. Allen‡
Pacific Biomedical Research Center, Snyder Hall 306, University of Hawaii at Manoa, 2538 The Mall, Honolulu, Hawaii 96822,
USA
*Present address: Laboratory for Brain-Operative Devices, Brain Science Institute (BSI), The Institute of Physical and Chemical Research (RIKEN), 2-1 Hirosawa,
Wako, Saitama 351-01, Japan
‡Author for correspondence (e-mail: [email protected])
Accepted 25 August; published on WWW 18 October 1999
SUMMARY
The contractile vacuole complex of the fresh water
protozoan Paramecium multimicronucleatum exhibits
periodic exocytotic activity. This keeps cytosolic osmolarity
at a constant value. The contractile vacuole, the central
exocytotic vesicle of the complex, becomes disconnected
from its surrounding radial arms and rounds before its
fluid content is expelled. We previously proposed a
hypothesis that the rounding of the contractile vacuole
corresponds to an increase in its membrane tension and
that a periodic increase in membrane tension governs the
exocytotic cycle. We also proposed a hypothesis that
transformation of excess planar membrane of the
contractile vacuole into 40 nm diameter tubules, that
remain continuous with the contractile vacuole membrane,
is a primary cause for the tension development in the
planar membrane. In order to investigate tension
development further, we have examined electron
microscopically the contractile vacuole membrane at the
rounding phase. To do this, we developed a computer-aided
system to fix the cell precisely at the time that the
contractile vacuole exhibited rounding. In this system a
decrease in the electrical potential across the contractile
vacuole membrane that accompanied the vacuole’s
rounding was monitored through a fine-tipped
microelectrode inserted directly into the in vivo contractile
vacuole. A decrease in membrane potential was used to
generate an electric signal that activated an injector for
injecting a fixative through a microcapillary against the cell
at the precise time of rounding. Subsequent electron
micrographs of the contractile vacuole membrane clearly
demonstrated that numerous ~40 nm membrane-bound
tubules formed in the vicinity of the vacuole’s microtubule
ribbons when the vacuole showed rounding. This finding
suggested that membrane tubulation was the cause for
topographical isolation of excess membrane from the
planar membrane during the periodic rounding of the
contractile vacuole. This together with stereo-pair images
of the contractile vacuole complex membranes suggested
that the microtubule ribbons were intimately involved in
enhancing this membrane tubulation activity. Electron
micrographs of the contractile vacuole complexes also
showed that decorated tubules came to lie abnormally close
to the contractile vacuole in these impaled cells. This
suggested that the contractile vacuole was capable of
utilizing the smooth spongiome membrane that lies around
the ampullae and the collecting canals to increase its size.
INTRODUCTION
decorated spongiome’s outer periphery The decorated
spongiome is not usually found around ampullae. All
membranes are organized around a cytoskeleton of
microtubular ribbons that originate at the CV pore. The ribbons
pass in pinwheel fashion over the surface of the CV and out to
the tips of the radial arms, one ribbon passes along each arm
(Hausmann and Allen, 1977).
Excess cytosolic water, acquired osmotically, is segregated
as a result of the activity of the decorated spongiome (Ishida
et al., 1993) and is transferred ultimately from the ampullae
into the CV. The CV then expels the water to the exterior of
The contractile vacuole complex (CVC) is an osmoregulatory
organelle of fresh water protozoa. In Paramecium it consists of
a central contractile vacuole (CV) and 5-10 radial arms that fan
out from the CV. A radial arm consists of (1) an ampulla
adjacent to the CV, (2) the collecting canal which is continuous
with the ampulla, (3) the smooth spongiome that branches from
both the ampulla and the collecting canal, and (4) the decorated
spongiome, which is continuous with the smooth spongiome
at its inner periphery and ends blindly in the cytosol at the
Key words: Contractile vacuole, Electrophysiologically controlled
fixation, Membrane potential, Membrane tubulation, Membrane
tension, Microtubular ribbon, Paramecium
3734 T. Tominaga, Y. Naitoh and R. D. Allen
the cell through the periodic opening of the pore. In a single
exocytotic cycle, the CV at first undergoes a relatively slow
swelling process, as the segregated water enters the vacuole
(the fluid filling phase). This is followed by a rapid rounding
process (the rounding phase) and ends by a rapid shrinkage of
the vacuole as the fluid is expelled to the cell’s exterior through
a fixed pore (the fluid expulsion phase) (Allen and Fok, 1988;
Kitching, 1967; Patterson, 1980; Wigg et al., 1967; Zeuthen,
1992).
We previously found that an isolated in vitro CV also
showed rounding-slackening cycles (Tominaga et al., 1998a)
without subsequent fluid expulsion. Such rounding without
expulsion can also occur in the in vivo CV (Patterson and
Sleigh, 1976; Tominaga et al., 1998a). Based on this finding,
we proposed the hypothesis that the CV membrane possesses
its own mechanism by which its tension is periodically
increased. The rounding phase thus corresponds to the
increased membrane tension phase of the CV. Such a periodic
change in membrane tension may also govern the exocytotic
cycle of the in vivo CVC (see also Tani et al., 1999).
Previously we also found (Tominaga et al., 1998b) that
the membrane potential and the input capacitance of the
contractile vacuole complex recorded through a fine-tipped
microelectrode inserted into the in vivo CV markedly
decreased immediately after the start of the CV’s rounding
phase. As input capacitance corresponds to the membrane area,
the capacitance decrease implied that the CV has become
disconnected from the radial arms during the rounding phase.
Also, since the membrane potential is most likely generated in
the tubules of the decorated spongiome which are part of the
radial arm (Tominaga et al., 1998b), the drop in membrane
potential measured by the electrode indicates that the CV is no
longer connected to its electrogenic source (Giglione and
Gross, 1995) when the CV was disconnected from the radial
arms.
Furthermore, we (Tominaga et al., 1998a) also demonstrated
that the roundness of the CV increased while the volume of the
CV remained unchanged at its maximum level after the CV had
been disconnected from the radial arms during the rounding
phase. This implies that the CV itself possesses a mechanism
by which the amount of planar CV membrane (corresponding
to the apparent surface area of the CV membrane) is
effectively reduced. In addition, we demonstrated electron
microscopically that the CV membrane became transformed
into a network of 40 nm tubules (Allen and Fok, 1988; Naitoh
et al., 1997b) when the CV expels its content. We (Tominaga
et al., 1998a), therefore, proposed the hypothesis that the
apparent transformation of the CV membrane by tubulation
(here termed ‘enhanced tubulation activity’) is the primary
cause for the topographical isolation of excess membrane from
the CV’s planar membrane during the rounding phase.
The primary objective of this paper is to determine electron
microscopically whether membrane tubulation is enhanced in
the CV membrane when the CV undergoes rounding and
whether such tubulation is the primary cause of rounding. In
order to fix the cell precisely in the rounding phase of the CV,
we developed a mechanism to electrophysiologically detect a
decrease in the CV’s membrane potential and to use this
decrease to trigger the discharge of fixative against the cell.
This technique has allowed us to detect the early stages of 40
nm tubule formation of the CV membrane and to show that the
tubular enhancement begins adjacent to the microtubule
ribbons when the CV is in its rounding phase. Only relaxed
tubules are present along these ribbons in the late fluid filling
phase. We also observed that the rounded CV membrane away
from the microtubular ribbons does not tubulate. We conclude
that enhanced membrane tubulation during the rounding phase
is somehow promoted by the association of the CV membrane
with the CV’s cytoskeleton of microtubular ribbons.
It has been demonstrated, where laser tweezer techniques
were used to measure the membrane tension produced by
tethers, that the tension of the plasma membrane is an
important factor in controlling the shape of the cell and its
motility (Dai et al., 1998; Sheetz and Dai, 1996). Visualization
of the membrane dynamics exhibited by intracellular
organelles and vesicles is also important for understanding
cellular processes such as membrane recycling and surface
membrane expansion as occurs, for example, in neurons
(Ashery et al., 1996; Dailey and Bridgman, 1993). For such
studies the electrophysiological trigger technique we describe
here and use to fix Paramecium at a precise stage of CV activity
should be adaptable and highly useful to study membrane
dynamics at precise times in other cells and organelles.
MATERIALS AND METHODS
Cells
Cells of P. multimicronucleatum (syngen 2) (Allen et al., 1988) were
grown in an axenic culture medium at 24°C (Fok and Allen, 1979)
and harvested at the mid-logarithmic growth phase. These cells were
washed with saline solution containing (final concentration in mM)
1.0 KCl, 1.0 CaCl2 and 1.0 MOPS-KOH buffer (pH 7.0). The cells
were equilibrated in the solution for more than 4 hours prior to
experimentation (Naitoh et al., 1997b).
Electrophysiological control of the timing of fixation
A diagram including a flow-chart for the computer-assisted control of
the timing of fixation of the in situ CV during the rounding phase is
shown in Fig. 1. An equilibrated cell was placed into a small droplet
of the saline solution under silicone oil on a small (2 × 2 mm) section
of a glass slide. Excess saline was pipetted out of the droplet until the
cell was squeezed sufficiently by the saline-oil boundary to become
immobile.
The tip of a microcapillary electrode filled with 3 M KCl (e1;
approximately 50 MΩ) was inserted into the cytosol and grounded. A
fine-tipped microcapillary electrode filled with 3 M KCl (e2;
approximately 100 MΩ) was inserted into one of the CVs (CV1) in
order to monitor the membrane potential across the CV membrane
(ECVC). This electrode also served to inject square current pulses (0.3
nA) used to monitor the input resistance of the organelle. Input
resistance increased 5 times over that of the cell when the electrode
tip successfully entered the CV (Tominaga et al., 1998b). An electric
oscillation generated by a temporary (approximately 20 milliseconds)
overcompensation of the stray capacitance of the head amplifier was
indispensable for successful penetration of the electrode tip into the
CV (Tominaga et al., 1998b).
The tip of a microcapillary (approximately 10 µm in inner
diameter) filled with fixative containing 2% glutarardehyde in 50 mM
cacodylate buffer was placed in the silicone oil at a distance of
approximately 50 µm from the saline droplet where the cell was held
(fixative pipette). This pipette was connected to the pressure outlet of
an electronic microinjector (IM-200, Narishige USA, Inc., Greenvale,
NY, USA) which was used to squirt the fixative against the cell when
the injector was activated by a timing signal from an A/D-D/A
Membrane dynamics of the contractile vacuole 3735
Fig. 1. Schematic representation
of the procedures for the
electrophysiologically controlled
fixation of a Paramecium
multimicronucleatum cell.
(A) Arrangement of the
electrodes and a flow-chart of
the computer-assisted controls.
e1; a grounded microcapillary
electrode has its tip inserted into
the cytosol. e2; a fine-tipped
microcapillary electrode has its
tip inserted into one of the two
CVs (CV1; another is labeled
CV2) through which the
potential difference across the
CV membrane (ECVC) is
measured. (B) Time course of
change in ECVC. ECVC gradually
decreases as the CV enters the
rounding phase. A horizontal
solid line corresponds to the
level for ECVC during the fluid
filling phase. The upper
horizontal dotted line
corresponds to the threshold
potential level for triggering an
electric signal for activating the
fixative injector. ∆E; the
difference between ECVC during
the fluid filling phase and the
threshold level. The threshold
potential was set by changing
∆E in the control system. The
lower horizontal dotted line
corresponds to the reference
(cytosolic) potential level. The
fixative is squirted against the
cell with a certain delay after the
triggering signal is fed into the
injector.
converter (ITC 16, Instrutech Corp. Great Neck, NY, USA). The
temperature of the experimental vessel was held at 17°C.
ECVC was fed into a computer (Power Macintosh 7600/136, Apple
Computer Inc. Cupertino, CA, USA) through an A/D-D/A converter.
The computer was programmed to send an electric signal to the
microinjector to activate it when ECVC decreased from its normal
value maintained during the fluid filling phase (approximately 60 mV)
(Tominaga et al., 1998b) to a predetermined lower value (threshold
level for activating the injector) that is achieved at the start of the
severing of the radial arms from the CV. The injector that expelled the
fixative against the cell responded with some delay (approximately 90
milliseconds) after it was activated by the signal from the computer
(Fig. 1B). The timing of fixation during the rounding phase was
controlled by changing the threshold level. To fix the cell during the
CV’s much longer fluid filling phase the injector was manually
activated to squirt the fixative against the cell. The hydrostatic
pressure of the inside of the microcapillary had been kept slightly
lower than the atmospheric pressure to prevent leakage of the fixative
through the opening of the capillary before activating the injector.
Also the oil prevented unintentional mixing of the fixative with the
saline solution before injector activation. The amount of fixative
squirted into the droplet was approximately 0.2 µl, which is 10 times
as large as that of the saline solution surrounding the cell. Software
for feeding the electrical signals into the computer and for generating
3736 T. Tominaga, Y. Naitoh and R. D. Allen
the electric signals required for the experiments was developed on the
basis of the IgorPro (WaveMetrics, Inc., Oswego, OR, USA) and
pulseControl XOP software packages (Herrington et al., 1995).
4 hours in 1% glutaraldehyde in 50 mM cacodylate buffer (pH 7.4).
The fixative was microinjected against the cell under the oil. The cell
was video taped for future reference.
Video microscopy
Images of the cell obtained using Nomarski optics (×40 objective lens
on a Leitz-DMIRB microscope, Leica Mikrosk. u. System. GmbH.
Wetzlar, Germany) were monitored on a television screen after capture
by a video camera (CCD-72, Dage MIT Inc., Michigan City, IN, USA;
Fig. 1A) and were recorded continuously by a video cassette recorder
(AG6300, Panasonic Indust. Co., Secaucus, NJ, USA) at 30 frames
second−1 during the experiments. The video images of the contractile
vacuole were fed into the same computer using a frame grabber (LG3, Scion Corp., Frederick, MD, USA) for determining the moment of
fixation of the cell.
Electron microscopy
Fixed cells were washed 2 hours in 50 mM cacodylate buffer (pH 7.4)
and post fixed in 1% osmium tetroxide in 50 mM cacodylate buffer
for 30 minutes. After a 30 minute wash in distilled water the cells
were prestained in a 4% aqueous solution of uranyl acetate for 1 hour,
washed 10 minutes in water and dehydrated in an ethanol series.
Embedment was in Epon 812. All the procedures of electron
microscopic preparation, except curing the Epon 812, were performed
at a room temperature of 24-26°C. Stereo pairs were taken by tilting
the specimen stage in the electron microscope plus and minus 10°
before taking pictures.
Fixation of the ruptured cells
An equilibrated cell was placed in a small drop of a saline solution
on a glass slide which was then covered with silicone oil. Excess
saline solution was removed through a micropipette until the pressure
exerted by the oil-water interface ruptured the cell. The cells were
fixed for varied times of not less than 15 minutes and not longer than
A
RESULTS
Fixation of the cell at the time of the CV’s rounding
A series of consecutive images of a representative cell of
Paramecium and an intermittent trace for the electric potential
of the CVC, with reference to the cytosolic
potential level (ECVC) as recorded simultaneously
before, during and after computer-aided fixation of
the cell, are shown in Fig. 2 (A, ECVC; B, cell
images). ECVC had been almost constant at
approximately 60 mV during the fluid filling phase
(frames 0-6). It then began to decrease as the CV
began to round (frame 7). When ECVC decreased
to a threshold potential level, which was set at
µ
Fig. 2. Changes in the membrane potential (A) and the shape (B) of a CV of a P. multimicronucleatum cell before, during and after fixation of
the cell. Lower dotted line in A corresponds to the reference (cytosolic) potential level (0 for ECVC). Upper dotted line in A corresponds to the
threshold potential level for activating the fixative injector. Arrowheads in A correspond to the time when the images of the cell shown in B
were taken. Numbers next to arrowheads correspond to the frame number in B. In frame 0 in B, the CV containing the fine-tipped
microelectrode, e2, is labeled. This e2 electrode is not in sharp focus. e1 is the microelectrode whose tip is in the cytosol. e1 and e2 correspond to
e1 and e2 in Fig. 1, respectively.
Membrane dynamics of the contractile vacuole 3737
approximately 50 mV in this case, the computer system
generated an electric signal to activate the injector after a
certain delay (approximately 103 milliseconds). The moment
of activation of the injector could be recognized by a downward
spike on the potential trace which occurs as an artifact signal
from the injector. Approximately 97 milliseconds after the start
of fixative injection into the saline droplet, the ECVC quickly
fell to 0. This moment corresponds to frame 8 where the cell
image is blurred by the cell’s displacement caused by the
fixative current against the cell and/or cell contraction. The CV
was at the middle of its rounding phase when the fixative was
injected. ECVC, then, began to increase to a steady level of
approximately 35 mV that was reached in a second or two. This
steady potential level varied case-by-case and is thought to be
an artifactual tip potential caused by fixed membrane or
cytosolic material forming a plug in the tip of the
microelectrode. CV activity ceased immediately after fixative
injection, and the CV remained rounded, as is clearly seen in
frames 9-11. The cell’s ciliary activity also ceased immediately
after fixative injection and ciliary orientation remained in a
reversed direction (cilia are poorly seen in Fig. 2B).
Electron micrographs of the CV of a cell thus fixed
Electron microscopy of a serially sectioned CVC in a cell fixed
as the CV had reached the late rounding phase (Fig. 3A)
demonstrated that the CV’s pore was still closed but the
membrane systems along the radial arms had become separated
from the CV. The CV was rounded following fixation (Fig. 3B)
but due in part to the mechanical stress required to remove the
microelectrode that was used to monitor the CV’s membrane
potential from the CV, its shape had become more oval than
spherical (Fig. 3C). A section passing tangentially through the
pore region at the CV’s peripheral hemisphere is shown in Fig.
3D. The membrane of this rounded CV is for the most part
planar (Fig. 3E) with no sign of extensive tubulation except in
the immediate vicinity of the microtubular ribbons.
These ribbons originate at the surface of the CV pore and
pass over the peripheral hemisphere of the CV (Hausmann and
Allen, 1977). Unlike the membrane that covers most of the CV
the CV membrane near the microtubules takes on a highly
modified topography (Fig. 3F,G). Here the membrane forms
clumps of tubules of 40 nm or so in diameter or a network of
such tubules. It was impossible to determine the amount of
such tubulated membrane but the impression received is that
the amount could represent a significant fraction of that making
up the planar CV membrane.
Against the rounded CV a microtubular ribbon lies flattened
in the plane of the spherical CV membrane. All excess
membrane extends as tubules that form along the edges of the
microtubular ribbons or that protrude from between adjacent
microtubules where groups of microtubules of the ribbon are
separated by a narrow gap. These tubules often extend around
to the cytosolic face of the ribbons where they became
associated with the ribbons’ cytosolic side and, in effect,
partially enclose the ribbon.
Such tubulation occurs all along the ribbons from the pore
to the beginning of the radial arms (Fig. 3G). Such non-planar
membrane is even more pronounced along the extended radial
arms where extensive networks of tubulated membranes occur
depending on the phase of fluid filling or expulsion in the
ampullae. These networks make up what has been called the
smooth spongiome (for terminology see Patterson, 1980). Even
where the ampullae are no longer continuous with the CV
membrane, sectioned tubules can be found along the
microtubular ribbons in the gap between the rounded CV and
the emptied and more-or-less collapsed ampullae.
Fig. 4 shows that the membrane of the ampulla/collecting
canal that is attached to the cytoskeleton of microtubular
ribbons along a radial arm is not continuous with the CV
membrane at the rounded phase. Images of three sections in a
set of serial sections (Fig. 4A-C) of the collecting canal at its
juncture with the CV show a small gap (Fig. 4B) between the
CV and the canal bridged only by the microtubular ribbon
(Fig. 4A and two additional sections not shown between A and
B). As reported earlier (Allen and Fok, 1988) there is usually
no special cytoskeletal material evident at this juncture even
in cells fixed at the rounding phase at the time this connection
is known to be severed. These serial sections, however, do
show that in cells that have had a microelectrode inserted into
their CV the CVC organization can be altered. In Fig. 4 it is
evident that the decorated tubules have come to lie against the
CV membrane. In in vivo cells the decorated tubules are never
found to lie around the ampullae (Allen and Fok, 1988) and
so they are always kept a distance of a few micrometers (∼5
µm) away from the CV membrane. Thus the time needed to
insert the microelectrode into the CV and to begin to obtain
the video images and electrophysiological recordings was
long enough for the CV to have gone through a few filling and
expulsion cycles. During this time the CV often becomes
enlarged which requires significant amounts of additional
membrane to be incorporated into the CV membrane. In Fig.
4 the membrane of the ampulla has apparently been used to
enlarge the CV.
In a cell fixed by activating the fixative injector manually at
its CV’s fluid filling stage, when the CV membrane potential
remained at a relatively constant level of 60 mV or more, the
serially sectioned CV was found to be attached to the radial
arms, the CV pore was closed and the CV’s shape was less than
spherical (Fig. 5A). The microtubular ribbons near the pore
were still attached to the CV membrane but the normal rigidity
of these planar microtubular ribbons that arise from the side of
the pore caused the membrane of the CV to assume a fluted
cross-sectioned profile near the pore (Fig. 5B). The membrane
of the CV appeared to be more relaxed, i.e. it did not have the
same extensive microtubule-associated tubulation seen in the
rounding phase (Fig. 5C,D). Tubules could still be seen near
the microtubular ribbons but these were less extensive,
particularly near the pore, wider in diameter and seemed to be
under less tension as though they might be in a state of
relaxation.
Stereo images of tubulated membrane in ruptured
cells
Some whole cells clearly show a close relationship between the
CV membrane and the microtubular ribbons in the late
expulsion phase (Naitoh et al., 1997b). However, cells ruptured
and flattened at the end of the fluid expulsion phase (Fig. 6),
clearly showed the CV membrane to be extensively tubulated
into uniform 40 nm diameter tubules. These tubules assumed
mostly a two-dimensional plane against the microtubular
ribbons where they came to lie oriented more-or-less
perpendicularly to the long axis of the ribbons. Stereo-pairs
3738 T. Tominaga, Y. Naitoh and R. D. Allen
Fig. 3. A CVC of a P. multimicronucleatum cell fixed during the CV’s rounding phase. The cell was fixed when the membrane potential across
the CV membrane, ECVC, decreased to a threshold potential level set to trigger the discharge of the fixative against the cell. (A) The trace of the
electrical potential difference between e1 and e2, as shown in Fig. 1. The left open arrow indicates the moment when e2 was inserted into the
CV. The potential, hereafter, corresponds to ECVC. A horizontal dotted line corresponds to the threshold potential (approximately 62 mV) used
for triggering injection of the fixative. The right filled arrow indicates the moment when the fixative discharge started. (B) A light micrograph of
the CV taken 500 milliseconds after the fixative discharge. Bar, 40 µm. (C) A section through the middle of the same CV as that shown in B.
Area between brackets is enlarged in E. Bar, 10 µm. (D) A tangential section of the same CV showing the CV pore (p) and the attachment to a
radial arm, boxed areas to left and right, respectively. (E) CV membrane away from microtubular ribbons is planar. Bar, 0.5 µm.
(F) Enlargement of the CV through its surface next to the pore (boxed area to the right of D). Grazing sections of microtubular ribbons
(arrowheads) that are attached to the CV membrane are interspersed with non-planar membrane tubules (arrows) lying to the sides of the
ribbons. Bar, 0.2 µm. (G) Membrane tubules (arrows) are also found along the microtubules (arrowhead) that pass from the CV to the radial
arm (boxed area to the left of D). Non-planar, tubulated membrane is found on both sides of the microtubules. Bar, 0.2 µm.
show this relationship best (Fig. 6, use a stereo viewer for
easier recognition). The tubules are spaced almost equally in
this stereo image. This spatial association is highly suggestive
that microtubules play some essential role during the collapse
of the planar CV membrane into these tubules as their collapse
seems to be centered on the ribbons.
Ruptured cells also clearly demonstrate the threedimensional network of the non-planar smooth spongiome
along the radial arms. The three-dimensional lattice is most
evident in a stereo pair (Fig. 7). The membrane of the lattice
can be described as having a cubic symmetry (see
Discussion).
Membrane dynamics of the contractile vacuole 3739
Regeneration of the CV
Attempts to push the tip of a microelectrode into the CV during
electrical oscillation frequently caused the CV to rupture and
deflate. We observed that a new CV soon appeared (within a
few seconds) after the original CV was ruptured. Moreover, a
third CV could appear again soon after the second CV had been
ruptured by further electrical oscillation. This renewal of the
CV could be repeated several times until the cell deteriorated.
DISCUSSION
Fig. 4. Sections of a serially sectioned CVC of P. multimicronucleatum
that show the relationship of a collecting canal (cc) with the CV (cv)
during the rounding phase. Like the CVC in Fig. 3 this CVC was fixed
after the membrane potential across the CV membrane decreased to
the threshold level for triggering fixative discharge against the cell.
Such microelectrode-impaled CVs tended to become abnormally large.
(A) Section 1 shows the microtubular ribbon (arrowhead) that reaches
from the CV membrane to pass along the collecting canal (cc) which is
out of the plane of this section. The smooth spongiome (ss) that
encircles the collecting canal is present as is the peripherically placed
decorated spongiome (ds). (B) Section 4 is one of the first sections
showing the collecting canal (cc). Thus, the collecting canal is seen to
be separated from the CV by a short gap. (C) Section 14 in this series
shows the collecting canal (cc) with its microtubular cytoskeleton (left
arrowheads) enclosed by the non-planar membranes of the smooth
spongiome (ss) and, peripheral to this, the decorated spongiome (ds)
that lies abnormally close to the membrane of the enlarged CV.
Sectioned trichocysts (t) can be used to vertically align the serial
sections. Bar, 0.5 µm.
Computer-aided electrophysiological control of the
timing of fixation
We previously proposed a hypothesis that the periodic
exocytotic activity of the CV in the Paramecium cell is
primarily governed by periodic development of tension in the
CV membrane which is associated with the retrieval, i.e.
topographical isolation of some of its membrane due to its
transformation into membrane-bound tubules that have a
diameter of approximately 40 nm (enhanced tubulation
activity) (Tominaga et al., 1998a). In order to examine electron
microscopically the beginning of the transformation of the CV
membrane into tubules, we needed to fix the CV precisely in
the very short rounding phase that occurs immediately prior to
fluid expulsion.
At the start of the rounding phase the electrical potential,
which is approximately 60 mV across the CV membrane with
reference to the cytosolic potential, began to decrease toward
0 (Tominaga et al., 1998b). We, therefore, utilized this decrease
in the CV potential for triggering the generation of an electric
signal (controlled by a computer) to activate an injector to
inject fixative into the small saline droplet that contained the
cell (Figs 1, 2). The saline droplet was under silicone oil. The
timing of fixation could be controlled by changing the
threshold potential level for generating the electric signal for
fixative injection. When the threshold potential level was
lowered the electric signal could be generated later fixing the
CV at a later rounding phase.
We conclude that the computer-controlled injector triggering
system we developed required less than three seconds, possibly
as little as half of a second, to fix individual cells as the electron
micrographs show that the impaled CVs are all within the 1 to
3 second window of the rounding phase, i.e. the CV pore is
still closed while the ampullae have already separated from the
CV. This membrane potential-controlled fixation system is
unique and is potentially useful for electron microscopical
examination of cells in relation to a variety of morphological
and physiological activities where physiological parameters
such as membrane potential can be measured.
Tubulation of the CV membrane during the rounding
phase
Electron microscopy of the CV membrane at very precisely
timed stages indicates that the rounding phase is accompanied
by an enhanced membrane tubulation of its excess membrane.
This tubulation only occurs along the margins of the
microtubular ribbons (Fig. 3F,G) and does not occur over the
planar non-microtubule-associated parts of the CV (Fig. 3E).
As tubulation progresses the tubules may collapse into a threedimensional network of membranes that comes to cover the
3740 T. Tominaga, Y. Naitoh and R. D. Allen
Fig. 5. A CVC of a P. multimicronucleatum cell fixed during the
CV’s fluid filling phase. (A) Low magnification of the CV (cv) at the
level of the CV pore (arrowhead). Bar, 5 µm. (B) The CV (cv)
membrane is fluted near the pore (p) due to its regular association
with the microtubular ribbons (arrowheads). The CV membrane
shows little evidence of tubulation near the pore during the fluid
filling phase. Bar, 0.5 µm. (C) Junction of a collecting canal with the
CV which is open to the CV (cv). The non-planar membrane
(arrows) along the microtubular ribbons (arrowheads) at this junction
appears relatively relaxed compared to the tubules at a similar
location in the rounded CV. Bar, 0.2 µm. (D) Another connection of
the CV (cv) to a collecting canal showing the microtubular
cytoskeleton (arrowhead) and related non-planar membranes (long
arrows) next to the ribbon. Openings of tubular invaginations (short
arrows) appear in the gap between two parts of the ribbon in this
section. Bar, 0.2 µm.
that tubulation itself is produced mainly in association with
microtubules. Such a mechanism in which the CV rounds as a
result of the tubulation of excess membrane does not require a
contractile cytoskeleton covering the entire CV membrane,
which we never see. However, a lack of a global cytoskeleton
around the CV does not rule out the possibility that an actinmyosin type of contractile process may indeed be associated
with the microtubules and might be involved in a more
restricted membrane tubulation. In this regard, Doberstein et
al. (1993) reported the presence of myosin I around the
contractile vacuoles of Acanthamoeba. We do sometimes see
a zone of exclusion around the membrane tubules, where
ribosomes and other small cytosolic components are prevented
from approaching the tubules. What lies around the tubules and
between the meshes of the tubular network is currently
unknown.
The appearance of the CV membrane in the fluid filling
phase (Fig. 5) suggests a more relaxed membrane in which the
membrane tubules near its microtubular ribbons are in the
process of expanding into a more planar topography as the CV
fills. This appearance fits a cyclic process of enhanced
membrane tubulation during the short period of rounding and
fluid expulsion which is followed by a much longer period of
membrane relaxation that covers most of the CV cycle.
The membranes of the radial arms also undergo fluid fillingexpulsion cycles that occur out-of-phase with that of the
tubulation-relaxation cycle of the CV membrane. The radial
arm membranes tubulated after they had expelled their content
into the CV when the CV membrane is relaxed. These
membranes then became planar as the ampullae, no longer
connected to the CV, now fill. If enhanced tubulation of the
filled ampullae occurs just before fluid expulsion as it does in
the CV, this might be manifested as a movement of tubules
along the microtubular ribbons where the membranes of
ampullae would be expected to encounter the collapsed CV
membrane bringing about fusion of these two membranes.
Fusion would permit the release of the contents of the ampullae
into the CV.
cytosolic side of the ribbon and thereby partially encloses the
ribbons. Such enclosing networks are commonly found all
along the ribbons of the radial arms.
This observation implies that rounding is brought about by
an enhanced tendency of the membrane to tubulate and that the
membranes assume non-planar symmetries. We now observe
A role for microtubules in membrane tubulation
Electron micrographs of serially sectioned CVs and,
particularly, stereo images of ruptured cells make it clear that
the collapse of the CV membrane is focused on sheets of
microtubules during the fluid expulsion phase. However, how
microtubules promote this membrane tubulation is not known.
Membrane dynamics of the contractile vacuole 3741
Fig. 6 Stereo pair of the CV pore (p) and microtubular ribbons in a section of a ruptured P. multimicronucleatum cell. The CV was at the end of
the fluid expulsion phase. The CV membrane has collapsed into a complex tubular system (arrows) lying parallel to the microtubular ribbons
(arrowhead). Some tubules lie free of microtubules. Bar, 0.5 µm.
What is known is that ER membranes can be pulled out into
long tubules as a result of their attachment to microtubules
(reviewed by Schroer and Sheetz, 1991). In this case the ER
membrane is tethered to the microtubules and is presumably
pulled along the microtubules by motors such as kinesin or
cytoplasmic dynein that can bind the membranes to the
microtubules. It is possible in Paramecium that such
associations may occur between the microtubular ribbons of
the CV cytoskeleton and the CV membrane. Bridges of
unknown composition can always be seen between the ribbons
and the membrane (Hausmann and Allen, 1977) both during
the fluid filling phase when tubulation is not enhanced as well
as in the rounding phase when tubulation is enhanced.
Enhanced tubulation activity begins at the rounding phase
and may persist to the end of the fluid expulsion phase. This
enhanced tubulation activity is initially visualized as an
increase in membrane tubule activity along the microtubular
ribbons and the tightening of the CV membrane around its
content to form a sphere. This precedes and leads to the
electrophysiologically detectable detachment of the radial arms
from the CV membrane (Tominaga et al., 1998b). This
detachment is physically demonstrated in this study and
is shown to occur before the CV pore opens. The
electrophysiological triggering of fixation of the CV at a
specific time has allowed us to view the physical image of the
CV at a very precise phase, a feat impossible to attain by
random sampling of fixed CVs. Thus we show that rounding
is initiated at the exact time that the membrane of the CV shows
enhanced tubulation activity along its microtubular ribbons.
The tubulation activity is initially exhibited as an array of 40
nm tubules lying flat against the microtubular ribbons and at
various angles to the long axis of the ribbon itself. In the radial
arms this tubulation is further enhanced into the formation of
networks of membranes which exhibit cubic morphology such
as that discussed by Landh (1995) of which a bi-continuous
form was recently described in mitochondria of an amoeba
(Deng and Mieczkowski, 1998). Cubic membranes are now
postulated to occur in caveolar membranes (Rietveld and
Simons, 1998). Thus the smooth spongiome membrane
together with the CV membrane itself may fall into a category
of biological membranes of growing interest and of potential
importance that collapse into a membrane that exhibit tubular
and/or cubic symmetry when their bending energy is released.
We have already shown that the CV membranes of
Paramecium potentially store enough bending energy to
account for the work done by the in vitro CV during fluid
expulsion when their membrane reverts from a planar to a
tubular form (Naitoh et al., 1997a).
Utilization of the smooth spongiome for increasing
the CV’s size
Observations of the displacement of the decorated spongiome
3742 T. Tominaga, Y. Naitoh and R. D. Allen
Fig. 7. Stereo pair of the collecting canal
(cc) and smooth spongiome (arrows) of a
radial arm in a section of a ruptured P.
multimicronucleatum cell. The CV was at
the end of the fluid expulsion phase. The
smooth spongiome membrane exhibits
cubic symmetry. Arrowheads, microtubular
ribbons. Bar, 0.5 µm.
toward the CV membrane due to external manipulation of the
CVC, such as is viewed in Fig. 4, or to naturally occurring
conditions that prevent the CV pore from opening (Tominaga
et al., 1998a) show that the smooth spongiome from around the
ampullae and the collecting canals can become incorporated
into the CV membrane. Thus the CV membrane and the
smooth spongiome membrane appear to be interchangeable.
However, we have no morphological or other evidence that the
decorated tubular membrane ever becomes continuous directly
with the CV membrane. The existence of a monoclonal
antibody that we had raised against Paramecium protein that
labels the membranes of the CV, collecting canals and smooth
spongiome, along with the plasma membrane, but not the
decorated tubules (Fok et al., 1995; Naitoh et al., 1997b), had
already suggested that the CV and smooth spongiomes were
similar to each other but different from the decorated tubules.
Thus, rapid growth of the CV appears to be possible by
incorporation of the smooth spongiome into the CV membrane.
Based on their electron micrographs of the contractile
vacuole complex, Fok et al. (1995) estimated the total surface
area of the decorated spongiome of the two CVs of a single
cell with a standard size to be approximately 21×103 µm2. We
previously reported that the interiors of all parts of the
contractile vacuole complex during the fluid filling phase are
electrically isopotential (Tominaga et al., 1998b). The input
capacitance of approximately 180 pF obtained from a single
CVC at this phase indicates the organelle’s total membrane
area is approximately 18×103 µm2, which is calculated based
on the assumption that the specific membrane capacitance of
the CV membrane approximates 1 µF cm−2 as is estimated for
other conventional biomembranes (Cole, 1968). The area of the
smooth membrane of a single CVC, therefore, approximates
7.5×103 µm2. This area is sufficient to surround a CV of 49
µm in diameter in its rounding phase. It has been reported that
the CV, whose normal in vivo size is approximately 12 µm
diameter in standard saline solution, can become as large as 50
µm in diameter when its CV is inhibited from expelling its
content through its pore by exposing the cell to 33 µg l−1
cationized ferritin in the external saline solution (Naitoh et al.,
1997b). This coincidence of the membrane area for a CV of
maximum size with that for the total amount of smooth
membrane of the CVC also supports our idea that smooth
membranes of the contractile vacuole complex can be utilized
by the CV when it is needed for increasing CV size. It will be
necessary to investigate the mechanism by which the smooth
membrane is differentiated into the CV, ampullae, collecting
canals and smooth spongiome, which may differ in their
function depending on their position within the CVC.
Involvement of cytoskeletal factors, such as microtubular
ribbons, in this differentiation would be especially interesting.
Fig. 8. Schematic drawings of the CVC and its exocytotic cycle
giving special attention to the enhanced tubulation activity of their
membranes along the cytoskeleton of the microtubule ribbons in P.
multimicronucleatum. (A) The CVC. CV, the contractile vacuole; AP,
an ampulla; CC, collecting canal; SS, smooth spongiome; DS,
decorated spongiome; MTR, microtubular ribbon; P, pore of the CV
through which its fluid content is expelled to the exterior of the cell.
(B) The enhanced tubulation activity in association with an
exocytotic cycle. The tubulation activity is indicated by the
membrane projections that correspond to the tubules, T. The length
of the projection corresponds to the relative degree of the tubulation
activity. Longer (shorter) projections correspond to higher (lower)
activity. (C) More detailed membrane dynamics associated with
closure of the CV pore after fluid expulsion. See the text for details.
Membrane dynamics of the contractile vacuole 3743
A
DS
CV
SS
AP
CC
MTR
P
B
a
b
AP
CV
AP
CV
T
c
MTR
f
AP
T
P
e
CV
CV
d
AP
AP
CV
AP
P
C
c
b
a
3744 T. Tominaga, Y. Naitoh and R. D. Allen
Exocytotic cycles of the CV with special attention to
the tubulation of the membrane
Based on our present results together with previous published
results (Naitoh et al., 1997a,b; Tominaga et al., 1998a,b) we
have produced a schematic drawing of the exocytotic cycle of
the CV and ampullae giving special attention to the enhanced
tubulation activity of their membranes along the microtubule
ribbons (MTR; Fig. 8B). A more realistic drawing of a portion
of the contractile vacuole complex is presented in Fig. 8A.
During the fluid filling phase (Fig. 8Ba) of the CV the tubulation
activity is high in the ampullary membrane (the degree of the
activity is shown as the relative length of the membrane
projections that represent the tubules) so that the ampullae are
thin. Segregated water enters the CV from the ampullae so that
the CV swells. The tubulation activity of the CV membrane is
predicted to be low in the fluid filling phase (reduced number
of membrane projections). At the end of the fluid filling phase
enhanced tubulation activity in the CV membrane becomes
high, so that the CV membrane starts to actively tubulate. The
tubulation of the CV membrane results in an increase in the
tension in the CV membrane, which leads to severing of the
ampullae from the CV and the rounding of the CV (the rounding
phase; Fig. 8Bb). As the membrane tension reaches a maximum
at the end of the rounding phase, the CV’s pore (p in Fig. 8Bc,d)
opens, so that the segregated fluid in the CV is expelled to the
outside of the cell through the pore. Expulsion is due mainly to
the in vivo cytosolic pressure (Naitoh et al., 1997b) (fluid
expulsion phase; Fig. 8Bc,d). The CV membrane then continues
to tubulate as the segregated water is expelled, although
enhanced tubulation activity may be reduced in this phase (Fig.
8Bd). The pore closes at the end of fluid expulsion as the
membrane tension decreases (Fig. 8Be). The ampullae begin to
swell immediately after the separation of the ampullae from the
CV, as segregated water continues to flow into the ampullae
from the collecting canals (Fig. 8Bc,d,e). The swollen ampullae
reattach to the tubulated CV membrane and fluid from the
ampullae enters the CV so that the portion of the tubulated CV
attached again to ampullae (to the left in Fig. 8Bf) quickly
swells. The tubulation activity in this phase is apparently high
in the ampullary membrane and low in the CV membrane. With
the reattachment of additional ampullae to the CV additional
tubules of the CV swell and fuse with the previously swollen
portion of the CV thus forming a single CV. Segregated water
continues to flow into the CV through the ampullae which are
now thin due to their membrane being tubulated (back to Fig.
8Ba).
Detailed membrane dynamics associated with closure of the
pore, which corresponds to Fig. 8Bd,e, are shown in Fig. 8C
and are based on what normally occurs during fusion and
fission of biological membranes. The plasma membrane,
continuous with the CV membrane after pore opening (Fig.
8Ca), fuses immediately after the CV collapses at the end of
fluid expulsion (Fig. 8Cb). To retain continuous membranes the
longitudinal section of the pore at this stage should show three
lipid bilayers across the pore although we have yet to capture
this stage in our electron microscopic studies. Two of the lipid
bilayer membranes, the two derived from the CV, should then
fuse. The tubulated CV may then be pulled apart along the
microtubular ribbons, so that the pore is now covered only by
the plasma membrane (Fig. 8Cc), the situation we usually
observe at this stage.
As is depicted in Fig. 8B, an enhanced tubulation cycle of
the ampullary membranes may precede that of the CV
membrane. This phase difference in the tubulation cycles of the
CV and the ampullae would require that the membrane
dynamics associated with the exocytotic activity of the
contractile vacuole be well coordinated. The mechanism by
which the timing of tubulation in both membranes is controlled
remains unknown. The possible involvement of the
microtubule ribbons in the timing mechanism should be
investigated.
It is interesting to note here that severing of the ampullae
from the CV, which can be detected as a sudden decrease in
the input capacitance of the contractile vacuole complex, takes
place even when the CV’s pore does not open for one reason
or another (Tominaga et al., 1998a). This implies that the
coordinated tubulation activity does not need to be directly
followed by fluid expulsion from the CV. Moreover, certain
chemicals (such as cationized ferritin) or mechanical agitation
(such as insertion of a microneedle) of the cell can cause
prolongation of the period of the exocytotic cycle, although the
rate of water segregation which is at least partially under the
control of the decorated spongiome (Ishida et al., 1993) is not
affected (Naitoh et al., 1997b). This implies that the water
segregation activity of the contractile vacuole complex itself is
not dependent on the enhanced and coordinated tubulation
activity of the CVC membranes. Rather enhanced tubulation
of the CVC membrane controls the gates in the plumbing
system for moving the segregated water from its site of fluid
collection to the CV and ultimately out of the cell.
This work was supported by NSF Grants MCB 95 05910 and MCB
98 09929. We thank Dr Tomomi Tani for his valuable comments. The
Biological Electron Microscope Facility is supported in part by NIH
grant RR-03061 and by NSF instrumentation grants
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