Download The measurement of exocytosis in plant cells

Journal of Experimental Botany, Vol. 47, No. 299, pp. 717-728, June 1996
Journal of
Experimental
Botany
REVIEW ARTICLE
The measurement of exocytosis in plant cells
Nicholas Battey1'3, Andrew Carroll 2 , Pirn van Kesteren 1 , Alison Taylor2 and Colin Brownlee2
1
2
Plant Science Laboratories, The University of Reading, Whiteknights, Reading RG6 6AS, UK
Marine Biological Association, Citadel Hill, Plymouth PL12PB, UK
Received 3 October 1995; Accepted 12 February 1996
Abstract
Exocytosis is of vital importance to the growth and
development of plant cells. It is a dynamic process in
which vesicles bearing polysaccharide precursors and
proteins fuse with the plasma membrane and release
their contents. Equally important, new plasma membrane is delivered by exocytosis as secretory vesicle
membrane becomes incorporated. The requirements
for polysaccharides, proteins and plasma membrane
are very different in different cell types, so there must
be sophisticated mechanisms for ensuring delivery of
these materials to the correct cellular locations at the
appropriate time and, particularly in the case of membrane, their recovery and recycling. Currently, little is
known of these mechanisms in plants, but new
methods for measuring exocytosis are under development, and existing techniques have already contributed data of considerable relevance. Here the methods
for measuring exocytosis are described and evaluated,
with emphasis on the electrophysiological measurement of capacitance as a relatively non-invasive
method, and on cell-free assays because of their
potential importance in the identification of proteins
and other factors that control exocytosis in plant cells.
Key words: Exocytosis, vesicle traffic, vesicle fusion,
polysaccharides, cell wall, cell plate, root cap, secretion,
patch-clamping.
Introduction
Exocytosis is the process by which plant cells secrete
polysaccharide precursors for cell wall elaboration and
hence cell growth. Extracellular proteins also cross the
plasma membrane by exocytosis, following synthesis on
the endoplasmic reticulum and transport through the
3
To whom correspondence should be addressed. Fax: +44 1734 750630.
© Oxford University Press 1996
Golgi apparatus. Further, membrane necessary for
plasma membrane growth and for its differentiation
during the life of the cell is incorporated from the
secretory vesicle membrane during exocytosis. For each
of these processes (polysaccharide, protein and phospholipid delivery) there may be alternative routes, but the
available evidence suggests that the exocytotic pathway
is of major importance.
The processes of cell development that depend on
exocytosis, and the molecular mechanisms that may be
involved in its control have been reviewed elsewhere
(Battey and Blackbourn, 1993). The emphasis of this
review will, therefore, be on methods for measuring
exocytosis in plants. This stress on techniques is justifiable
because although the study of exocytosis is in its infancy
in plants, a wide variety of methods is available from
work on animals and yeast. The presence of a cell wall
and high turgor pressure creates problems specific to
plant cells so that careful application of these methods is
required. Yet it is this wall and the importance of cell
growth in plant development that makes quantification
of exocytosis of such especial interest in plants.
Exocytosis releases secretory product and increases
plasma membrane area. Methods that measure either of
these processes are, therefore, potentially able to report
exocytosis, but because of limitations in both approaches,
the ideal solution is to try and estimate secretion and
membrane area simultaneously. A good deal of effort is
currently focused on the estimation of membrane area by
capacitance, so this technique has been given some prominence here. Additionally, cell-free assays in which secretory vesicles and plasma membrane are isolated and then
mixed in a fusion assay, are useful as a means to isolate
and purify regulators of exocytosis. They may also contribute valuable information on mechanisms, if interpreted with caution and in parallel with measurements of
718 Battey et al.
exocytosis in vivo. The opportunities for development
of these assays in plant cells have, therefore, been
emphasized.
Measurement of secretory product release
Polysaccharides
Changes in secretory product release are likely to be
accompanied by changes in the rate of exocytosis.
However, there can clearly be many causes of such
changes in flux, including alterations in rates of product
synthesis, packaging, and vesicular transport through and
from the Golgi, as well as in the exocytotic step itself. In
the root cap of many plants hypersecretory cells exocytose
large quantities of polysaccharide slime (Plate 1). Many
measurements of this secretion have been made and there
is much evidence for major control at the level of synthesis
(as reflected in Golgi activity; Rougier, 1981). However,
the analysis carried out by Bowles and Northcote (1974)
shows that it is possible to calculate flux specifically
for the post-Golgi (presumably exocytotic) step. Radioactivity from 14C-glucose saturates in the membrane compartments after 30 min, but continues to increase in the
exported polysaccharides. These workers were, therefore,
able to calculate the rate of export of polysaccharide
across the plasma membrane and to deduce that slime
components would need to be displaced from the Golgi
apparatus every 20 s, whereas polysaccharides for cell
wall formation would only be displaced every 150 s.
Presumably, application of this method would allow
measurement of the effects of putative agonists and
antagonists on exocytosis of root cap slime. A similar
approach, labelling secreted polysaccharide with 3Hfucose or 14C-arabinose, has been employed to study the
factors regulating exocytosis from sycamore suspension
cells (Morris and Northcote, 1977). This work showed
that addition of various cations caused a very rapid
increase in polysaccharide secretion, and it was speculated
that this was an effect on vesicle fusion mediated by a
rise in intracellular Ca 2+ .
In pea stem segments, rate zonal centrifugation has
been used to separate Golgi stacks and putative secretory
vesicles and, in conjunction with pulse-chase labelling
(with 3H-glucose), transit of polysaccharide from Golgi
to secretory vesicles to cell wall has been followed
(Robinson et al, 1976; Griffing and Ray, 1985). One
conclusion is that nigericin may inhibit exocytosis by
inhibiting zlpH-driven secretory vesicle swelling (Griffing
and Ray, 1985). This technique seems to have some
potential, for instance for study of the effects of auxin on
the exocytotic step (Napier and Venis, 1995), but it has
not been very widely employed.
A variety of probes can give more qualitative information on polysaccharide secretion. The fucose-specific lectin
UeA I (Ulex europaeus agglutinin I; Rougier et al, 1979;
Roy and Vian, 1991) shows the polar nature of fucoserich slime secretion from root cap cells. Calcofluor White
is useful for detection of /3-glucans (Galbraith, 1981;
Griffing et al, 1986). In the special case of cell plate
formation, fusion of exocytotic vesicles leads to the formation of the new cross-wall at the end of telophase (see
Battey and Blackbourn, 1993, for review). Aniline blue
staining shows that /91-3 glucan (callose) is an early
product of the newly formed cell plate, and that callose
synthesis continues in cell plates after their isolation from
BY2 cell protoplasts (Kakimoto and Shibaoka, 1992).
This provides an excellent means of identifying isolated
cell plates: aniline blue is reasonably specific for callose
(Wood and Fulcher, 1984), and this is reflected in specific
staining of the cell plate (Plate 2).
Proteins
Plate 1. Exocytosis from the maize root cap. Polysaccharide 'slime' is
the visible product of vigorous exocytosis from the hypersecretory cells
of the maize root cap. Hypersecretory activity is confined to only a few
cell layers, is polarized, and is very sensitive to external factors (x 35).
(Photograph: courtesy of Dave Maw, Jealott's Hill Research Station,
Bracknell.)
Secretion of a-amylase is perhaps the best characterized
protein secretion in plants (see Jones and Jacobsen, 1991,
for review). As a measure of exocytosis it is very useful
because secretion can be measured from protoplasts (Bush
et al, 1986). It occurs over a well-defined period (36-72
h), and is Ca2+-dependent. Further, secretion of a-amylase from protoplasts is controlled by both GA and ABA,
which seem to exert their effects by changing intracellular
Measurement of exocytosis
719
Plate 2. Cell plates can be detected by aniline blue staining. The procedure is based on that of Kakimoto and Shibaoka (1992); the cells are
pretreated for 1 h with 3 mM UDP-glucose (a callose precursor) and then stained with aniline blue. Left: Nomarski optics. Right: UV fluorescence
image of the same field. Upper panel: intact tobacco BY-2 cells showing synchronized cell plate formation after treatment with aphidicolin and
propyzamide using a method based on that of Yasuhara et at. (1993). A minor component in commercial preparations of aniline blue shows
yellow-green fluorescence when bound to callose in the cell plate. An example of a cell plate is arrowed. Lower panel: Cell plates isolated from
tobacco BY-2 cell protoplasts according to the method of Kakimoto and Shibaoka (1992), DNase and cold treated, and further purified by sucrose
density gradient centrifugation. Aniline blue fluorescence indicates that the cell plates are typically intact after this procedure, but the Nomarski
image suggests that some cytoplasm still remains associated with some of the cell plates (x 145).
720 Battey et al.
Ca2+-levels (Gilroy and Jones, 1992). Zorec and Tester
(1992, 1993) have measured exocytosis as a change in
membrane capacitance in aleurone protoplasts (see
below). A parallel study of a-amylase secretion and
exocytosis would be very informative, particularly in view
of the predicted effects of GA and ABA on (Ca 2+ dependent) exocytosis in this system.
There are other cell types where further study of protein
secretion would be rewarding. Peroxidase secretion from
spinach cell suspensions is stimulated rapidly by the
addition of 1 mM Ca 2+ to the medium (Sticher et al,
1981). Although the possibility of release of cell wall
peroxidases could not be ruled out, the evidence was in
favour of an effect on release from the intracellular
compartment and, hence, presumably exocytosis. It would
be very interesting to study this process in protoplasts.
plasma membrane must take place in hypersecretory root
cap cells, to allow the vesicle fusion required for delivery
of polysaccharide slime in the absence of cell growth (for
discussion see Morre and Mollenhauer, 1983). One estimate puts this at 10% of the plasma membrane recycled
per minute (Shannon and Steer, 1984). Even in the
rapidly elongating cells of the coleoptile more than sufficient membrane can be delivered by secretory vesicles
(Quaite et al, 1983; Phillips et al, 1988).
Clearly estimates of exocytosis that depend on measurement of plasma membrane area (such as membrane
capacitance-see below) must take account of this active
recycling. This presents some difficulty because the mechanism of recovery is still unclear. In the root cap, coated
vesicles appear to retrieve membrane not only from the
plasma membrane, but also from secretory vesicles in
transit (Mollenhauer et al, 1991). However, it is not
clear that endocytosis is quantitatively sufficient to
account for the required rate of membrane recovery
Measurement of plasma membrane area
(Shannon and Steer, 1984; Emons and Traas, 1986; Steer,
The maximum elastic stretching of the plasma membrane
1988). The close association of cortical ER with the
is about 2% of its surface area (Wolfe and Steponkus,
plasma membrane may facilitate molecular recycling of
1981, 1983; Wolfe et al., 1985; Dowgerte/ al., 1987). The
membrane lipids (Craig and Staehelin, 1988), a mechanauthors concluded that further increases in plasma memism that has yet to be discounted and may be favoured
brane area can only be accounted for by the addition of
by the high turgor pressure acting on the plant plasma
new membrane material from intracellular stores. The
membrane
(Cram, 1980; Staehelin and Chapman, 1987).
problem is, therefore, to estimate accurately the rate of
Grabski
et
al.
(1993) obtained evidence for rapid transfer
this addition. Micropipette aspiration techniques, in
of
fluorescently-labelled
lipids from the plasma membrane
which negative pressures are applied to patches of memto
the
ER,
and
have
interpreted
this as consistent with
brane through a micropipette, reveal an equilibrium mem1
such
a
recycling
mechanism.
It
is also possible that
brane tension (mean value 120 /xN m" ) above which
exocytosis
leads
to
formation
of
a
pore,
release of vesicle
new material is added to the plasma membrane until the
contents,
and
then
vesicle
reformation,
so that vesicle
tension is released (Wolfe et al., 1985). Such exocytotic
membrane is only temporarily incorporated into the
addition of membrane can accommodate surface area
plasma membrane. This has been proposed to explain
changes resulting in tensions of up to 4 mN m" 1 ; above
the current 'flicker' detected in nerve terminals (Spruce
this protoplast lysis occurs. This approach illustrates the
et al, 1990).
exocytotic delivery of new membrane. However, it does
The most complete descriptive model for the exocytotic
not allow quantitative determination of the kinetics of
event in plant cells has been presented by Staehelin, based
membrane flux.
on observation of disc- and horseshoe-shaped infoldings
of the plasma membrane in suspension culture (Staehelin
Estimation of vesicle supply and recycling using microscopy
and Chapman, 1987) and root cap cells (Craig and
This topic has been comprehensively reviewed by Steer
Staehelin, 1988) (Plate 3). These are interpreted as steps
(1988). Specific aspects of immediate relevance to the
in the fusion of exocytotic vesicles, which after fusion are
measurement of exocytosis are focused on here. The most
flattened due to the high turgor pressure of plant cells,
detailed transmission electron microscope (TEM) anaand then tip over sideways, presumably as a result of
lysis is of pollen tube growth in relation to secretory
movement of the cytoplasm relative to the plasma memvesicle formation/delivery rates (Picton and Steer, 1983;
brane (Fig. 1). This model has been confirmed in work
Steer and Picton, 1984). The principal conclusion is that
on carrot suspension cells (Emons et al, 1992). A decline
vesicle production rates are very similar in fast- and slowin the number of exocytotic configurations in elongated
growing tubes, suggesting that vesicles are produced in
cells between 1 and 14 d after subculturing may reflect a
excess to allow maximum growth under favourable condidecrease in transport of hemicelluloses and pectins due to
tions (Steer and Picton, 1984). This means that an excess
exhaustion of sugar in the medium (Emons et al, 1992).
of membrane is delivered to the pollen tube tip, so that
It is interesting that these important observations of
some form of recycling must normally occur. Similar
Staehelin and Emons are not necessarily inconsistent with
considerations indicate that very active removal of the
the idea of transient vesicle fusion discussed by Spruce
Measurement of exocytosis
721
Plate 3. Exocytotic configurations in the plasma membrane of a sycamore suspension cell. Freeze-fracture electron microscopy was used on the
plasma membrane of cells ultra-rapidly frozen by liquid propane. This view of the exoplasmic face of the membrane shows horseshoe-shaped
membrane infoldings that are believed to reflect exocytotic vesicle fusion. For the mechanism proposed to account for these shapes, see Fig. 1.
Reproduced from Staehelin and Chapman (1987) by permission of Springer-Verlag, and courtesy of the authors (x40 000).
Fig. 1. The Staehelin/Chapman model for exocytosis from plant cells. The diagrams illustrate the stages in vesicle fusion that Staehelin and
Chapman (1987) suggest lead to the horseshoe-shaped configurations shown in Plate 3 Top row diagrams: plasma membrane viewed from the
inside (plasmatic face, PF) as the vesicle approaches (I), and docks and forms a fusion pore (II) At stage III the fusion pore is deformed to a slit
because the vesicle is flattened under the forces of turgor following content release, and at stage IV the vesicle topples over and the fusion slit
expands along the margins of the vesicle to give the horseshoe-shape. Bottom row diagrams: side view of the same series of events. Stages III and
IV appear to be unique to plants and probably reflect the effects of high turgor. The model implies that the plasma membrane can not immediately
expand to accommodate the new vesicle membrane after fusion, presumably because it is constrained by the cell wall. Redrawn from Staehelin and
Chapman (1987).
et al. (1990): it is not clear what happens after flattening
of the vesicle (Stage IV in Fig. 1). Recovery of the spent
vesicle could be the next step (Craig and Staehelin, 1988;
Staehelin and Chapman, 1987). However, TEM pictures
of hypersecretory root cap cells certainly give the impression of complete vesicle incorporation into the plasma
membrane (Plate 4), so it is possible that both mechanisms operate.
722
Battey et al.
Plate 4. Transmission electron micrograph of a hypersecretory cell from
the maize root cap. The section was double stained with uranyl
acetate/lead citrate according to standard procedures. The polysacchande slime is unstained and is massed along the upper edge of the cell.
The profile of a recently fused secretory vesicle is visible (large arrow),
and vesicles apparently about to fuse can also be seen (small arrows).
Note the cytoplasm contains both stained and unstained vesicles
(x9200).
Estimation of plasma membrane area by
electrophysiological measurements of membrane
capacitance: theory
Biological membranes function as a barrier between two
electrolyte solutions, and so they have a number of
electrical properties. One of these is capacitance (Cm),
which in all membranes has a constant value per unit
surface area xl ^Fcm~ 2 . The current flowing through
a capacitor is proportional to the rate of change of the
applied voltage, i.e.
T=CxdV/dt
where C=capacitance. Exocytosis involves the addition
of new membrane, which if added at a greater rate than
it is removed will result in an increase in membrane
surface area. Thus measurement of the changes in Cm
provides a valuable, dynamic and geometry independent
probe in the study of exocytosis. The measurement of
capacitance in plant cell membranes depends upon the
application of a changing voltage across the membrane
and detection of the currents induced. This can be done
using the whole cell patch clamp technique (Hammill
et al., 1981). To measure Cm from whole cell recordings,
a sinusoidal command voltage (Kcom) is applied to a cell
inducing a sinusoidal current to flow across the plasma
membrane. This current has both resistive and capacitative components, each with a distinctive phase angle (0)
relative to the command signal. The phase of the capacitative current is orthogonal to the phase of the resistive
current under normal circumstances. Thus, determining
the phase of the resistive current (0 — 90°) allows the
phase angle of the capacitative current (d) to be calculated;
the current in this phase is measured as an output signal
proportional to the membrane capacitance, but independent of membrane resistance (Fig. 2).
Techniques used to monitor the changes in membrane
capacitance following exocytotic events were developed
principally to study stimulus-secretion coupling in excitable animal cells and have been used subsequently in a
wide range of animal cells (Neher and Marty, 1982,
Fernandez et al., 1984; Mason et al, 1989; Schweizer
et al, 1989; Zorec et al., 1991). Two general approaches
have been followed in the measurement of cell membrane
capacitance, identical in theory but different in methodology. These are the phase detection technique using a
lock-in amplifier (Neher and Marty, 1982), and the phasetracking technique (Fidler and Fernandez, 1989). The
principles involved are illustrated in Fig. 2A and B.
There are inherent problems with the phase detection
technique, because the impedance of the cell may change
during the experiment, as plasma membrane ion channels
open and close; or the series resistance (Rs—see Fig. 2)
may increase as the membrane begins to reform over the
ruptured patch between the walls of the pipette. This
causes the current from the cell (7C) to drift out of phase,
so that the amplifier is not necessarily reading current at
a phase directly proportional to membrane capacitance
(Fig. 2C). In response to these problems, Fidler and
Fernandez (1989) described a phase detection technique
whereby the phase angle (9) is automatically tracked and
continuously adjusted. This means that changes in the
electrical properties of the cell during the course of the
experiment can be accommodated. This approach is currently being used in studies of exocytosis from root cap
protoplasts.
The electrophysiological measurement of exocytosis in
plant cells offers a number of opportunities. It allows
exocytosis and membrane retrieval to be monitored directly; further, because the whole cell patch clamp configuration results in direct communication between the
cytosol and the microelectrode filling solution, cytosolic
Ca2+ can be buffered and putative regulators of exocytosis
perfused into the cytosol and their effects recorded. There
is also the potential to distinguish effects on exocytosis
from those on membrane retrieval. In animals this has
been achieved where there is clear temporal separation of
the two processes (e.g. in neurons; von Gersdorff and
Matthews, 1994; Heidelberger et al., 1994), or where
secretory vesicles are sufficiently large to allow individual
fusion events to be monitored (e.g. 0.5-1 /nm in mast
cells; Breckenridge and Aimers, 1987). In plants, even
where secretory vesicles are relatively large (e.g. about
500 nm in root cap hypersecretory cells; Plate 4) , this
size is at the lower end of the detectable range; and, in
Measurement of exocytosis
general, exocytosis- might be expected to be accompanied
by continuous membrane retrieval (although more
information is needed here). These are important considerations because they imply that as it stands the patch
clamp method will be useful for measuring the net result
of exocytosis and membrane retrieval, but not for
distinguishing between the two processes.
Estimation of plasma membrane area by membrane
capacitance: practical expectations and observations
Considerations of membrane recycling (see above) suggest
that secretory vesicle fusion should be followed by membrane recovery. In protoplasts, unless swelling is occurring, Cm might, therefore, be expected to increase
following vesicle fusion and then return to its original
value. However, what is actually observed will depend
critically on the number of vesicles arriving at the plasma
membrane per second, vesicle size in relation to plasma
membrane area, and the actual rate of membrane recovery
in the specific cell type. Typically, secretory vesicles are
about 150nm in diameter (Picton and Steer, 1981) and
a protoplast about 30 /nm in diameter. If each is assumed
to be spherical then a vesicle will contribute only 0.0025%
of the plasma membrane area. Capacitance changes of
1 pF can be detected readily (equivalent to about 3000
secretory vesicle fusions, but an increase of only 2% in
plasma membrane area), and so quite a large amount of
vesicle fusion can occur without a visible increase in
protoplast volume. On the other hand, in hypersecretory
root cap cells it has been estimated that an area equivalent
to the whole plasma membrane is delivered every 10 min
(Shannon and Steer, 1984); this gives an idea of the rates
of membrane delivery that are possible. However, even
in this case, if membrane recovery occurs as an immediate
consequence of vesicle fusion, capacitance will increase
only very transiently. It should also be noted that capacitance would be expected to increase if the membrane
thins significantly due to elastic stretching. This
exocytosis-independent increase in capacitance is generally thought to be small, but this is an assumption that
remains to be verified.
Cm has now been measured in protoplasts from the
barley aleurone layer (Zorec and Tester, 1992, 1993), and
maize coleoptile (Thiel et al., 1994). The results are
consistent in showing an increase in Cm in response to
approximately 1 yM free Ca 2 + in the patch-pipette, and
a decrease in response to < 3 0 n M free Ca 2 + . This suggests that elevated Ca 2 + causes an increase in the rate of
exocytosis (equivalent to about 10 vesicles s" 1 in the
aleurone protoplast; Zorec and Tester, 1992), relative to
the rate of membrane recovery. Further, the effect of
reduced Ca 2 + concentrations suggests that even basal
exocytosis (that needed to compensate for membrane
recovery) is Ca 2+ -dependent. Application of pressure
723
through the patch-pipette causes protoplast swelling, followed by a (5-fold slower) contraction to the original
diameter (Zorec and Tester, 1993). This is interesting,
because it suggests that exocytosis can be triggered (or
membrane recovery inhibited) directly by pressure, and
that plant cells have greatly enhanced sensitivity to turgor
pressure compared with animal cells. It is also significant
that pressure appeared to stimulate fusion of a different
population of vesicles than that involved in the Ca 2 + dependent response (Zorec and Tester, 1993).
The response of protoplasts to pressure is likely to be
different from the response of walled cells. It remains an
intriguing possibility that membrane recovery could be
activated under circumstances in which the cell wall
forbids further expansion in response to exocytosis. This
is in a sense the corollary of the proposal that tension
generated in the plane of the plasma membrane by cell
wall yielding provides the driving force for increase in
plasma membrane surface area (Kell and Glaser, 1993).
Accurate measurements of membrane capacitance, turgor,
and wall extension would indicate how the regulatory
hierarchy between these processes is maintained,
and might, in time, make the mechanism of action of
regulators such as auxin more intelligible.
Cell-free assays for exocytosis
Two cell-free approaches have been used previously to
assay for exocytosis in plants. Baydoun and Northcote
(1980) isolated Golgi-rich and plasma membrane-rich
fractions by density gradient centrifugation of 14C-choline
labelled and unlabelled root tips of maize. After mixing,
the fractions were re-purified and the transfer of 14C-lipid
was measured. Evidence was obtained for fusion between
the Golgi and plasma membrane fractions; Ca 2 + and
a membrane protein of 36 kDa were demonstrated to
be key factors for the fusion process (Baydoun and
Northcote, 1981). This pioneering work depended solely
on marker enzymes for identification of Golgi and plasma
membranes, and there was no indication of the intactness
or otherwise of the secretory vesicles and the Golgi.
Nevertheless, the results provided the first indication that
cell-free assays for exocytosis were possible in plants, and
that Ca 2 + might have a significant role. Morre et al.
(1991) immobilized spinach plasma membrane on nitrocellulose filters, and then measured the transfer of 14 Clipid from a Golgi fraction to the plasma membrane
acceptor. Transfer was temperature-sensitive and there
was a suggestion that it might be stimulated by ATP.
However, it is not clear that aggregation would be distinguished from fusion using this approach. These assays
point the way for future work, and highlight the following
areas for improvement.
724 Battey et al.
A.
B
R
R
I
Computer
control
c.
v
commV
I c pA
Ill
IcpA
Fig. 2. Schematic diagrams of a patch-clampcd cell illustrating the two methods of determining the phase angle d. In both systems the patch clamp
amplifier measures the current flowing in response to a sinusoidally varying command voltage ( l ^ ) and the equivalent circuit model of a patchclamped cell consists of series resistance R,, the cell membrane resistance Rm and the membrane capacitance Cm (A). The lock-in or stationary
phase detection technique, in which current from the cell / c is balanced by the current from the amplifier's transient cancellation circuitary / a at a
summing junction ( + ) . R^ and Ca are, respectively, variable series resistance and whole cell capacitance settings. R, and C2 are fixed resistance and
capacitance elements in parallel with Ra and C a . After making adjustments of Ra and C, so that / a = / c , Ri or C 2 is switched in using switches 1 or
2, respectively, to create controlled changes in the amplifier current. By altering the phase angle detector the amplifier is able to measure / c at a
phase where switching in /?[ or C 2 produces the largest current response; this phase angle is 9-90° if /?, is switched in, and 9 if C2 is switched in.
The proportionality of the induced current to the membrane capacitance is strongly dependent upon the accuracy with which the phase angle is
determined. Increases in cell capacitance are calibrated by making incremental increases of the whole cell capacitance compensation on the amplifier
(simulating increases in membrane capacitance) and noting the current response, in mammalian cells the phase detection technique has been used
to monitor accurately capacitance changes in the range 0.4-80 fF, sufficient to observe the fusion of a single secretory granule. (B). The phase-
Measurement of exocytosis
Purification of intact secretory vesicles, and distinguishing
these from vesiculated Golgi
A marker is needed that distinguishes post-Golgi vesicles
from Golgi. The absence of the Golgi enzyme xyloglucan
xylosyl transferase from secretory vesicles, while the fucosyl transferase remains active, may provide one such
marker (Brummel et al, 1990). The polysaccharide content of secretory vesicles changes as they mature, at least
in hypersecretory root cap cells (Battey and Blackbourn,
1993). This might provide a marker for sectioned
membrane pellets under the TEM.
Purification of cytoplasmic side-out plasma membrane
vesicles
Aqueous two-phase partitioning produces a population
of plasma membrane vesicles which are mainly 'outsideout' in orientation (Larsson et al, 1984). The vesicles
can be turned into the required cytoplasmic side-out
orientation by snap freezing and thawing (Brightman and
Morre, 1992; Larsson et al, 1994). Separation of vesicles
remaining in the outside-out orientation can then be
achieved using the countercurrent distribution procedure
(Widell et al, 1982; Akerlund and Albertsson, 1994). An
alternative, and potentially much more rapid method
involves incubating phase-partitioned plasma membrane
vesicles with the detergent Brij 58, which turns the vesicles
inside-out (Johansson et al, 1995). The only potential
problem with this very convenient method is if the
detergent alters the fusigenic properties of the plasma
membrane vesicles.
Distinguishing secretory vesicle docking (membrane
aggregation) from fusion with the plasma membrane
The assays developed by Baydoun and Northcote (1980,
1981) and Morre et al (1991) do not necessarily distinguish between docking and fusion, although that
described by Baydoun and Northcote would eliminate
Ca 2+ -dependent docking. Assays that measure membrane
or content mixing should reflect only fusion, although
even these methods need to be very carefully controlled.
It is reasonably well established that vesicle fusion is
accompanied by mixing of the lipids from the two membranes. This can be measured using lipid-soluble fluorescent probes that can be added to native membrane
vesicles after purification. In the first approach one population of vesicles is saturated with a probe such as
octadecylrhodamine B chloride ( R i 8 ) . On mixing with
725
the target membrane (unlabelled), the probe is diluted
and fluorescence dequenching occurs; this can be quantified and used as a measure of fusion (Hoekstra et al,
1984; Mac Lean and Edwardson, 1992; Hoekstra and
Klappe, 1993; Lee et al, 1994). This type of assay has
been the subject of some criticism and debate, still currently unresolved (Stegman et al, 1995; Edwardson,
1995).
In the second approach, resonance energy transfer
between two lipid-soluble fluorophores, with the emission
spectrum of one fluorophore partly overlapping the
excitation spectrum of the other, is used to monitor
membrane mixing (Comerford and Dawson, 1988; Keller
et al, 1977). Energy transfer depends on close proximity
between the lipid probes. Mixing a population of donor
vesicles with both probes loaded in their membranes, with
unlabelled acceptor vesicles, will therefore result in
a decrease in resonance energy transfer efficiency.
Alternatively, each probe can be loaded into a different
membrane; fusion is then indicated by an increase in
transfer efficiency. The advantages of energy transfer
assays for monitoring fusion have been described
(Hoekstra and Duzgiines, 1993), and include high sensitivity at low probe labelling rates, continuous monitoring,
and wide applicability involving both artificial liposomes
and native membranes. The relevance of this assay for
measuring fusion in biological membranes is indicated by
the good agreement between measurements of cell fusion
(induced by high PEG concentrations) made by syncitia
counting and by the resonance energy transfer assay
(Partearroyo et al., 1994). A particularly attractive feature
of the energy transfer assay is its potential to make visible
the process of membrane fusion (Uster, 1993).
The resonance energy transfer assay depends on probe
mixing and factors that affect diffusion of lipids in the
membrane bilayer are, therefore, likely to affect the assay.
Further, because native membranes vary in composition,
more variation in labelling efficiency can be expected than
with artificial liposomes. However, assays can be standardized by diluting labelled donor or target membranes
with unlabelled membranes to a set ratio of fluorescence
intensities, or, if the fluorophores are in separate membranes, to a set ratio of membrane amounts. As with
other assays, resonance energy transfer experiments
should include proper positive (e.g. PEG-induced fusion)
and negative controls (e.g. the use of EGTA to exclude
Ca 2 + -induced aggregation) in order to assess real fusion
events. Figure 3 shows an emission spectrum from a
tracking detection technique. Here a 1 Mfi resistor is in series with the cell. In parallel to this resistor is a computer-controlled reed relay switch.
During the course of an experiment this series resistance is switched in and the phase detector finds the phase of lc at which a maximal change
occurs, this phase angle being 8. (C). Illustration of the change in phase of lc with respect to the phase of l^,,,,, when either /? t (lock-in method) or
the 1 MQ resistor (phase-tracking method) is switched in. Traces (i) and (ii) show the phase of Vcom and /„ respectively; temporarily increasing a
resistive element (trace iii) causes Ic to shift relative to the phase angle (8-90°) of the lock-in amplifier (broken line). This causes an error in the /,.
reading and requires correction using the phase detector.
726
Battey et al.
125
o
u
a
v
u
o
500
S2S
550
S7S
600
Emission wavelength (nm)
Fig. 3. Resonance energy transfer assay for vesicle fusion Emission
spectrum of fluorescein-labelled plasma membrane vesicles mixed with
rhodamine-labelled Golgi membrane vesicles, after irradiation at the
excitation wavelength of fluorescein (460 nm). The increased emission
of rhodamine at 592 nm indicates energy transfer from fluorescein to
rhodamine and thus fusion between the differently labelled membranes.
Polyethylene glycol (PEG 3350) promotes vesicle fusion by removing
water from the vicinity of the membranes. Triton X-100 dissolves the
membranes and is a negative control.
resonance energy transfer assay of fusion between maize
Golgi-enriched vesicles and plasma membrane vesicles.
Content mixing assays provide an ideal method of
measuring vesicle fusion. However, even in this case
problems can arise due to leakage of contents during the
assay (Hoekstra and Diizgunes, 1993). Some content
mixing methods depend on incorporation of the probes
during the preparation of artificial liposomes used in the
assay; this is the case with the terbium/dipicolinic acid
technique developed by Diizgunes and Wilschut (1993).
These assays are therefore not so useful for studies of
exocytosis, in which native vesicles are essential.
Conclusions
In vivo quantification of exocytosis can be achieved by
capacitance measurements, and this approach has the
great advantage that potential regulators of exocytosis
can be tested after dialysis through the patch pipette.
However, the current limitation of patch-clamping for
plant cells is that it is necessary to use protoplasts. The
intact cell wall certainly has regulatory influences on
plasma membrane expansion and hence on exocytosis
and membrane recovery; it is the interplay between these
biophysical effects and their translation into the biochemical domain that is really crucial to the understanding of
exocytosis in plant cells. The potential importance of
turgor pressure for exocytosis is illustrated by the apparently unique character of the fusion process in plants
(Staehelin and Chapman, 1987; Emons et al, 1992; see
above). These observations emphasize the difficulties con-
fronting the attempt to quantify exocytosis, particularly
in a cell-free context.
Nevertheless, a cell-free approach is probably essential,
to provide biochemical evidence of the nature and function of regulatory molecules. This needs to be complemented by further work towards the cloning and
characterization of homologues of known regulators such
as syntaxin (Bassham et al, 1995). The identification of
exocytotic mutants in Arabidopsis, the use of epitopetagged secretory products, and advances in our knowledge
of polysaccharide and protein processing en route to the
plasma membrane will provide major opportunities for
improvements in the currently available techniques for
measuring exocytosis in plant cellls.
Acknowledgements
We are grateful to the BBSRC, Leverhulme Trust, Royal
Society, and The University of Reading Research Endowment
Trust Fund for financial support. Thanks to Paul Le Miere for
help with Figure preparation, and to Richard Napier (HRI,
Wellesbourne) and two anonymous referees for their invaluable
comments on the manuscript.
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