Download Cell Surface Membrane Remodelling and Mitochondrial

Modern Research and Educational Topics in Microscopy.
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Cell Surface Membrane Remodelling and Mitochondrial
Remodelling in Live Neurons: Practical Approaches for
Studying Dynamic Processes Using Confocal Microscopy
N. Shulyakova, P. Kaifosh, D. Diec and L.R. Mills*,1,2
1
Genetics and Development Division, Toronto Western Research Institute, University Health Network
11-430 Toronto Western Hospital 399 Bathurst St Toronto ON M5T 2S8
2
Department of Physiology, University of Toronto, Toronto, ON, Canada
Neurite outgrowth and growth cone motility are dynamic processes that involve a consequent remodelling
of the cell surface membrane. Plasma membrane remodelling in neurons is also essential for successful
recovery from insults that trigger cell swelling e.g., ischemia and excitotoxicity. Confocal microscopy, by
facilitating high resolution studies on live neurons in dissociated cultures and in slice cultures, has had a
major impact on our understanding of these events. During previous studies on plasma membrane
remodelling induced by osmotic perturbation, we also observed striking and reversible changes in
mitochondrial morphology. This article describes experimental protocols we have developed, and
practical solutions for problems encountered when examining cell surface membrane remodelling and
mitochondrial remodelling in live neurons.
Keywords neurons; PC12 cells; slice cultures: plasma membrane remodelling; mitochondria; VLDs
confocal microscopy; biolistic transfection; mtGFP; lck-GFP; lipophilic dyes; Lucifer Yellow
1. Introduction
Remodelling of the cell surface membrane occurs whenever neurons change their shape. As neurons
grow during development such activity is virtually continuous. In mature neurons remodelling occurs
during regeneration, and in response to insults that trigger neuronal swelling. Our previous studies
experimentally induced plasma membrane remodelling by using transient osmotic shocks to trigger cell
surface area regulation (SAR) [1-4]. (SAR) is the homeostatic process by which the plasma membrane is
reorganized in response to stimuli that cause cell swelling and shrinking. Swelling (induced by
hyposmotic medium), recruits internal membrane stores to the plasma membrane. During shrinking
(induced by an abrupt return to normosmotic medium), the now excess plasma membrane is retrieved,
generating transient vacuole–like dilations (VLDs) at the adherent surface. VLD recovery, the process by
which VLDs are re-internalized, involves cortical F-actin [2], and is partially dependant on ATP [2,3,4].
Neurons in hippocampal slice cultures demonstrate SAR, suggesting it is an intrinsic property of
neurons, and may be a key element in the brain’s response to ischemic insults [3]. During the course of
our studies on SAR, we observed concomitant changes in mitochondrial morphology. Unlike the
mitochondrial swelling observed in response to stresses associated with apoptosis [5], or necrosis (e.g.,
Ca2+ overload [6], glutamate toxicity [7], Cd2+ toxicity [8], loss of membrane potential [9], complex-I
inhibition [10]), the changes we observed were rapid, global, and reversible within minutes. Given the
complex morphology of neurons, this ‘mitochondrial remodelling’ may be essential for mitochondrial
trafficking in neurites, particularly under conditions of increased bioenergetic demands (e.g. during
regeneration, or at synaptic sites). This chapter describes some practical approaches for imaging cell
membrane remodelling and mitochondrial remodelling in live neurons.
*
Corresponding author: e-mail: [email protected], Phone: 416-603-5420
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2. Plasma membrane remodelling in live neurons
2.1
Experimental models
Confocal microscopy is the method of choice for monitoring dynamic events in live cells [11,12] To
examine remodelling of the plasma membrane we use osmotic perturbations to evoke neuronal swelling
and shrinking. Swelling is induced by replacing culture medium with a hyposmotic solution (medium to
water ratio 1:5 to 1:9) for 2-4 minutes; replacing this solution with normosmotic medium (290 mOsm)
causes rapid shrinking. To monitor these events we use fluorescent dyes to label the plasma membrane,
or the bath. Our previous experience with live cell imaging encompasses studies on molluscan neurons
[1,2], differentiated PC12 cells [4,13], sympathetic neurons [14], acute hippocampal slices [15, acute
spinal cord slices [16], and hippocampal slice cultures [3].
2.2 Optimizing imaging conditions
To reduce complications associated with cell overlap, PC12 cells and primary neurons are plated at low
density (5×104 cells/ 35 mm dish). Undifferentiated PC12 cells are treated with NGF for 4-6 days to
induce differentiation or can be are differentiated in tissue culture flasks for up to 10 days prior to
plating. Hippocampal slice cultures are cultured on porous membrane inserts (0.4µm, Millicell-CM), for
at least 7 days, and preferably longer, during which time they gradually ‘thin out’, optimizing
transfection and imaging conditions [17].
High resolution imaging requires high numerical aperture objectives, in our cases a 40X fluor or a
60X planApo. The short working distances of these objectives effectively limit imaging to the central
region of a 35 mm dish. To maximize cell attachment in this area, we plate neurons in a 75-100 µL drop
of medium in the centre of the dish, and allow them to attach for several hours before topping up the
medium. Our confocal is on an inverted microscope platform so in order to achieve high resolution
images we must grow cells on glass. Glass-bottomed dishes can be made by drilling a 10-20 mm
diameter hole in the centre of a 35 mm plastic dish and gluing a #1 coverslip (0.17 mm), to the bottom
surface of the dish, effectively creating a small well. The choice of glue, coverslip glass, and cleaning
agents, can be problematic, especially for hippocampal neurons, and we frequently use a commercial
source (WillcoWells). Alternatively, neurons can be cultured on #1 coverslips in plastic dishes, and the
coverslips transferred to specialized chambers for imaging; this approach is not ideal for experiments that
require cultures be transferred back to the incubator. Since PC12 cells and primary neurons do not
attach/grow as well on glass compared to plastic, the glass is coated with a double layer of 25% collagen
or 1.5 µg/ml poly-L-ornithine (PLO). PC12 cell growth is optimal on collagen, but for frequent media
exchanges PLO offers greater adhesivity. Laminin, while a good substrate for primary neurons, is not
optimal for PC12 cell differentiation. Note: Confocal imaging of live cells grown on plastic can be done
on an inverted microscope, using 10X and 20X fluor objectives (e.g. [14]), but there is considerable loss
of signal. For fixed cells however, high resolution imaging is possible on an inverted microscope using
the following modifications. A #1 coverslip is mounted over the culture (using clear nail polish), and the
dish is inverted face-down over the objective, attaching it to the underside of the microscope stage by
double-sided tape. Paraformaldehyde (2%) is the fixative of choice as glutaraldehyde is autofluorescent.
Where an up-right microscope platform is available, a water immersion lens permits high resolution
imaging of live cells growing on plastic. However, rapid medium exchanges are more difficult in this
configuration, the continued culture of now contaminated cells is not an option, and fluorescent reagents
necessitate very careful cleaning of the objective.
For imaging hippocampal slices the feet on the culture inserts are removed (use scissors), and the
inserts transferred to 50mm glass bottomed dishes. Under these conditions, organotypic cultures growing
on the inserts can be imaged using high numerical aperture objectives, in our case a 40X fluor, and with
some care, a 60X planApo [3,4,23]. Note: Intact inserts can used for long-term studies by transferring
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them to new culture dishes using pieces of capillary tube (diameter 1.5 mm) placed underneath the
membrane insert to substitute for the membrane feet (which normally serve to prevent flooding of the
tissue). When imaging in intact inserts is difficult, a section of the membrane (with the slice), can be cut
out, inverted in a glass bottomed dish, and immobilized by reducing the medium volume and partially
covering the central well with a coverslipPrior to imaging, culture dishes are affixed to the microscope
stage to eliminate movement; blue tack, or double-sided tape are inexpensive and highly effective. Relocating an individual neuron is difficult if a culture dish has been moved even slightly, so it is useful to
mark the original orientation with a permanent marker pen. Note: These pens should not be used on the
bottom of the dish since, despite their name, the ink is not indelible after contact with emersion oil.
Photo-etched gridded coverslips have limited use for tracking individual cells since they are too thick
(1.5mm). For repeated imaging fiduciary marks can be made prior to plating by very gently scratching
the glass surface with a diamond knife. Note: Deep scratches can fatally weaken the glass. The
requirements of live neurons for perfusion with oxygenated media vary considerably. In our studies the
duration of each cycle of osmotic perturbations was relatively short and continuous perfusion was not
required. Since rapid medium changes were essential and our perfusion system has a relatively slow
clearance rate (2 ml/min), exacerbated by the presence of the central well in glass bottomed dishes, we
typically removed the medium by pipette attached to a vacuum suction device. Medium was reintroduced
as a bolus, also by pipette. We usually elected to work at a (warm) room temperature (28 C), since it
slows plasma membrane remodelling slightly, facilitating the capture of fast events. Note: In previous
studies of calcium dynamics in acute spinal cord preparation [16], sympathetic neurons [14] and acute
hippocampal slices [15] imaging at room temperature improved cell survival.
We use an inverted (Nikon based) scanning confocal microscope (Bio–Rad MRC-600) equipped with
an argon-ion laser. To minimize photo-bleaching and photo-damage, potential cells are initially selected
in epi-fluorescence mode and confocal imaging parameters optimized with cells outside this field. To
reduce photobleaching we use a neutral density filter of 3 (1% transmission), a zoom factor below 3, and
single scans (rates ranging from 1/s to 1/30s). Simultaneous fluorescent and phase contrast images can be
taken using the second channel for phase (non-confocal, transmitted light) images. VLDs can also be
monitored in 3-D by optically sectioning neurons; optical series are taken using one single scan per
section and step sizes (the distance between two optical sections) of 0.5–3.0 µm. Note: Optical series are
only practical if the object of interest does not move and/or change shape during series collection.
2.3
Imaging plasma membrane remodelling
VLD formation and recovery were monitored by confocal microscopy using a variety of reagents that
label the plasma membrane, and by adding dyes to the bath. In studies on SAR high resolution phase
images, are also crucial to establish and maintain the focal plane, and to interpret the confocal images.
FM 4-64 (T13320, Molecular Probes, Invitrogen) is an lipophilic styryl dye (MW 60, excitation 558nm,
emission 734nm). This water-soluble dye is non-toxic, and virtually nonfluorescent in aqueous media,
but becomes intensely fluorescent upon insertion into the outer leaflet of the surface membrane. FM4-64
is reasonably photostable, after 50 frames in cells (pulsed-labelled with FM4-64), less than 10% of the
signal is lost to photobleaching [18]. We used 10µl/ml of a 1mg/ml FM 4-64 stock in methanol,
maintaining this concentration during solution exchanges and imaged with the BHS (fluorescein) filter
blocks. FM4-64 effectively labelled the plasma membrane (Fig.1). Images taken at the level of the
substratum showed that VLDs appeared as fluorescent rings that corresponded closely with VLDs visible
in phase mode (Fig. 1b,c). During recovery VLDs gradually shrank and then disappeared as they were
absorbed into the plasma membrane (see ref [1] for details). While easy to use FM4-64 was rapidly
internalized by endocytosis and labelled other membranes. For this reason, it is most useful for imaging
single cycles of SAR.
Trials of other membrane dyes indicated that if a dye stained both the plasma membrane and
endomembranes, distinguishing VLD membranes was difficult. The carbocyanine dye 1,18-dioctadecyl3,3,38,38-tetramethylindocarbocyanine perchlorate (DiI) in ethanol or in DMSO (final concentration 2.5
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µM) proved to be the best choice for selective and rapid labelling of the plasma membrane. DiI labelled
neurons were imaged using the GHS (rhodamine) filter block. Since DiI is relatively insoluble and forms
a fine, intensely fluorescent precipitate in the bath, it was added for only 30 sec in the swelling phase,
Fig. 1 Osmotic changes in PC12 cells labelled with FM 4-64: Swelling in hyposmotic medium for 4 min. (a) and
subsequent shrinking in normosmotic medium (b) induces formation of VLDs (arrow) to accommodate the decrease
in cell surface area. VLDs form as membrane invaginations at the adherent surface (see [1]). (c) phase contrast
image of the cell in (b). (d,e) VLDs spontaneously recover in normosmotic medium.
and cultures were washed repeatedly during the shrinking phase (triggered by normosmotic medium), to
eliminate the bath signal. Note: This protocol affords no opportunity to image the plasma membrane
during the initial swelling phase, but swelling can be monitored in subsequent cycles. DiI labelled
VLDs formed within 4 sec of the onset of cell shrinking [1], initially appearing as small fluorescent
‘points’. During the next minute, VLDs rapidly expanded and then gradually shrank. Comparison of
Figs. 2a and 2b shows that some VLDs visible in confocal mode were obscured in
Fig. 2 VLDs in snail neurons labelled with DiI. (a) Shrinking in normosmotic medium induces numerous VLDs
(arrow) visible in phase contrast mode. (b) Confocal image shows numerous DiI labelled VLDs some of which are
undetected by phase optics. (c) VLDs at different stages of recovery. Magnification, 60x.
phase images by the nucleus. At the level of the substratum DiI revealed a remarkable degree of
complexity in the morphology of VLDs during recovery. While some VLDs appeared to collapse into
the plasma membrane (flattened circles, Fig. 2c), others appeared as irregular annuli of motile, adherent,
processes (arrows in Fig. 2c). Such profiles were in marked contrast to the smooth membrane profiles
observed for the same VLDs at focal planes deeper in the cytoplasm (not shown). Strikingly, VLDs
formed in second or third cycles were similar in size, shape and site of origin to those formed in the first
cycle.
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Fig. 3 Bath dyes taken up by VLDs as they form label VLD contents. (a) Profiles of LY filled VLDs in PC12 cells,
image taken at the level of the adherent surface. (b) RD3000 filled VLDs in a snail neuron. This image, taken 5.0
µM above the substratum shows that VLDs extend part way into the cytoplasm.
2.4
Bath dyes and VLD dynamics
Lucifer yellow (L-453, Molecular Probes, Invitrogen) is a water-soluble, plasma membrane impermeable
dye (excitation/emission 428/536 nm) that we used to label the bath contents. While it is commonly
believed that Lucifer yellow (LY), due to its low absorbance at 488 nm, is insufficiently excited with the
argon-ion laser, our experience shows otherwise [1,15]. Rhodamine-Dextran 3000 (RD3000), was also
used to label the bath. Note: In bath labelling experiments, the capacity of the confocal microscope to
reject out-of-focus fluorescence is essential (in epi-fluorescence mode the bath signal predominates and
cells are not visible). In confocal images neurons appear black objects against an intense bath signal.
Within seconds of adding normosmotic medium to previously swollen cells, numerous VLDs of diverse
size and shape were detected (see Figure 3a,b). VLDs imaged via their fluorescent aqueous contents
behaved like VLDs imaged via membranes labelled with DiI and FM4-64; they formed rapidly (<1 min)
within the bounds of the cytoplasm as neurons shrank, and gradually disappeared during recovery. By
washing out the bath after VLD formation we determined that some VLDs were pinched off (i.e. they
retained their dye contents), while others remained open to the bath (i.e. these lost their dye content).
2.5
Biolistic transfection
While lipophilic dyes offer a reliable way to monitor SAR their use can complicate time lapse studies
due to dye internalization. With bath dyes, signal loss after specimen fixation is a limitation. Critically,
despite considerable effort, neither type of dye proved suitable for monitoring SAR in slice cultures. As
an alternative we explored biolistic transfection with a plasma membrane targeted green fluorescent
protein lckGFP. (lckGFP was a gift from Dr. Eubanks, see [18] for details about this plasmid). Using an
approach analogous to our use of bath dyes which labels VLD contents but not their membranes, we also
transfected with an EGFP plasmid that labels the cytoplasm but not the plasma membrane [3].
Gene gun cartridges are prepared according to the Helios gene gun manufacturer protocol (BioRad),
[3]. Briefly, 50µg of plasmid DNA is precipitated onto 25mg of gold particles (1.6 µm in diameter),
spread over the inner surface of the Tefzel tubing and dried. Note: While tungsten beads are less
expensive we found transfection yields were higher using gold particles. Cartridges are loaded into the
cartridge holder and a diffusion screen attached to the tip of the gene gun spacer. To prevent tissue
damage, and particle clustering, a membrane filter (Isopore polycarbonate membrane filter, 3µm pore
size, 8.0x105 pores/cm2, Millipore, TSTP04700) is inserted between the culture and the gene gun nozzle
[3, 19]. The membrane filter is glued to the bottom of a 50 mm tissue culture dish (WillCo Wells) with
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the glass bottom removed (~40mm diameter hole). Optimal shooting parameters vary for helium gas
pressure (80-200 psi) and distance between gun nozzle and the tissue (10-25 mm). To transfect
dissociated cultures the medium was aspirated from the culture dish, cells were transfected with a single
shot, and the medium restored. To transfect hippocampal slices (7-10 days in vitro), membrane inserts
were removed from 6-well culture dishes, placed on a plastic stage for transfection, and then returned to
the 6-well dishes. Cells expressing GFP or lck-GFP were visualized using fluorescein filter sets.
VLD profiles in transfected PC12 cells were similar to those observed using FM4-64 or DiI (compare
Figs. 1, 2 with Fig. 4). In slice cultures biolistic transfection labelled 10-50 cells per slice throughout the
CA1-3 and DG hippocampal neuronal layers. The number of labelled cells within different
Fig. 4 Visualization of VLD formation and recovery in PC12 cells expressing lckGFP in the plasma membrane after
biolistic transfection. Corresponding pairs of phase contrast and fluorescent images show cell before osmotic
changes (a, a’), during swelling in hyposmotic solution (b, b’). Return to normosmotic medium causes the cell to
shrink and VLDs, (e.g. arrow) form. (c, c’). VLDs spontaneously recover in (d, d’). Magnification, 60X.
regions varied, likely due to uneven distribution of DNA carriers. This low transfection yield creates
ideal conditions for imaging individual neurons which can be repeatedly imaged for up to 4 weeks.
EGFP transfected neurons (Figure 5c), had uniformly labelled soma and neurites whereas those
transfected with lckGFP (Figure 5d,e), had intensely labelled soma and less intensely labelled neurites.
Cells were selected for SAR experiments based on their morphology (the presence of dendrites and a
distinct axon) and location, subsequently cell identity could be confirmed by immunohistochemistry.
Transfected cells underwent marked morphological changes in response to osmotic perturbation [3],
and these changes were reversible (see Figure 6a-c). Both EGFP and lckGFP labelled neurons formed
VLD-like structures, indicating that neurons in slice cultures do demonstrate SAR. Transfected neurons
swelled and shrank repeatedly in response to multiple cycles of osmotic perturbation (normosmotichyposmotic-normosmotic) [3], in contrast to previous lower resolution studies on slices which showed
neuronal swelling but no recovery (shrinking), post-reperfusion [20]. This difference may reflect the fact
that our approach focuses on changes at the level of individual neurons.
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3. Imaging mitochondria and mitochondrial remodelling
Although confocal microscopy has been used for high resolution imaging of mitochondria in live
neurons for well over a decade [11], little is known about mitochondrial remodelling. What is clear is
that sustained morphological changes are implicated in a large number of disease pathways [5,21] and
that under physiological conditions mitochondria undergo constant oscillatory and directional motion
[10] as well as frequent fission and fusion [5]. To study mitochondrial remodelling we used osmotic
perturbations that mirrored those used to study SAR.
Fig. 5 Biolistic transfection of hippocampal slices with EGFP and lckGFP and mtGFP: a) EGFP transfection labels
the cytoplasm of numerous cells in hippocampal slice. b, c) EGFP labels the cytoplasm, so the cell body and neurites
(arrows) are readily visible; d, e) transfection with lckGFP labels plasma membrane and organelles membrane; areas
where organelles accumulate (arrows at soma in e) show intense fluorescence. f) Hippocampal neuron in
hippocampal slice culture transfected with mtGFP; g) fine details of the mitochondria (arrow) in f)
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3.1
Experimental models and optimizing imaging conditions
To image mitochondria we used two approaches; differentiated PC12 cells stably transfected with a
Fig. 7 Confocal images of mtGFP-labelled mitochondria. a) Prior to swelling, mitochondria are elongated and
tubular. b) Two minutes after introduction of hyposmotic medium (see text), mitochondria have rounded up and
some appear swollen. Arrows label mitochondria in the same neurite in (a) and (b).
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Fig. 8 Selected images from a 3 minute time series demonstrating mitochondrial remodelling in a PC12 cell. a)
elongated mitochondria prior to osmotic perturbation. Hyposmotic solution was introduced at 10 s and norm-osmotic
solution was reintroduced at 90 s. (b-d) mitochondrial rounding. (e-f) recovery of elongated morphology.
plasmid encoding an inducible mitochondrially targeted GFP fusion protein (mtGFP see [13]), and (in
cells where mtGFP synthesis is inhibited by tetracycline [13]), two commonly used mitochondrial
probes, Mitotracker Green FM and Rhodamine-123 (both Molecular Probes). For dye labelling, cells
were incubated for 20 min in 20-150nM Mitotracker (1mM stock in DMSO) or 0.5-2µM Rhodamine-123
(2.6mM stock in DMSO). Cells were selected for imaging based on their size, mitochondrial
morphology, and fluorescent signal intensity. Flatter cells and neurites are ideal since they restrict
mitochondrial movement into and out of the plane of focus. In any given culture, the degree of
differentiation, mitochondrial morphology, and signal intensity varied. Only well-differentiated cells
with neurites and intensely fluorescent and elongated mitochondria were selected. Isolated cells were
preferred because they were easier to distinguish from their neighbours.
Optimal results were obtained when images were focused on mitochondria in the bottom-most plane,
i.e., at the adherent surface. Using this focal plane permits comparison of mitochondria in neurites and
growth cones with those in the cell body. It also provides fiduciary marks (necessary for the refocusing
necessitated by rapid medium exchanges), since the overall cell footprint at the adherent surface remains
relatively static during swelling and shrinking.
Given the requirement for frequent images, photobleaching is a major concern. MtGFP is remarkably
stable; under our imaging conditions (see 2.1), signal intensity can be maintained over three sequential
cycles of mitochondrial remodelling (100 images over 15 minutes). Intervals of four or five seconds
between images were usually sufficiently frequent for tracking the dynamics of individual mitochondria.
In contrast, Mitotracker Green was highly susceptible to photobleaching and most of our experiments
with this dye were prematurely terminated for this reason. Although more stable than Mitotracker,
Rhodamine-123 (Rhod-123) bleached substantially especially in ‘unquenched’ mode (see below, and
[22]). Note: Faster scanning speeds can alleviate photobleaching, but at the expense of image resolution.
A more effective remedy was to scan less frequently using smaller osmotic perturbations which induce
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slower changes. For stressors that induce slower mitochondrial remodelling, slower acquisition speeds
may be sufficient; in these cases Rhod-123 may prove more useful.
Introduction of hyposmotic medium caused rapid changes in mitochondrial morphology. Within two
minutes, mitochondria lost their elongated form and rounded up (Fig. 7a,b). The response was global,
with similar changes occurring in the soma and in neurites. Upon re-introduction of normosmotic
medium, mitochondria rapidly reverted to elongated forms. Automated imaging at five second intervals
revealed the intermediate steps of mitochondrial shape changes, selected images from a time series are
shown in Fig. 8. Repeated cycles of swelling and shrinking evoked multiple rounds of mitochondrial
remodelling, but did not affect neuronal survival for up to 4 days. Although we did not collect time series
data with Mitotracker, or Rhod-123, mitochondria labelled with these dyes also demonstrated rapid and
reversible changes in mitochondrial morphology in response to osmotic perturbations [23]. Ongoing
experiments are using biolistic transfection of mtGFP to examine mitochondrial remodelling in slice
cultures (Fig. 5f,g and [24]) where the low transfection yield creates ideal conditions for imaging
mitochondria within single neurons.
Rhod-123 accumulates in mitochondria as a function of mitochondrial membrane potential; when
mitochondria depolarize, it is released into the cytoplasm [reviewed, 22]. We looked for both short term
changes (e.g. flickering), and sustained declines in mitochondrial membrane potential. Note: For these
studies we rinsed extensively post loading with Rhod-123 to eliminate bath dye. Under non-quench
mode conditions (loading concentration 0.5µM), no change in the mitochondrial signal was observed in
response to hyposmotic medium. In contrast, under quenching conditions (loading concentrations
>2.0µM), hyposmotic medium caused mitochondria to release a small amount of Rhod-123 into the
cytoplasm. The mitochondrial signal itself appeared unchanged. These results suggest that our swelling
regime caused only a modest and transient depolarization of the inner mitochondrial membrane. They
also emphasize the importance of selecting appropriate loading protocols for Rhod-123 (see [22], for an
excellent discussion of these issues). Western blots of cytoplasmic fractions collected 5 min and 2hrs
post 2 cycles of swelling and shrinking showed zero to trace amounts of cytochrome c, confirming that
our protocols did not result in a sustained depolarization of mitochondria, or activate apoptotic pathways.
3. Summary
Neurons in disociated cultures display a considerable capacity for cell surface remodelling [1,2,4,25].
Experiments using biolistic transfection of membrane targeted plasmids have shown similar events ocurr
in the complex 3-D environment of the hippocampal slice [3]. Studies on SAR using differentiated PC12
cells transfected with a mitochondrially targeted mtGFP revealed dramatic and reversible changes in
mitochondrial morphology [23, and unpublished observations]. These rapid morphological
transformations suggest that mitochondria have a hitherto unappreciated capacity to remodel in response
to stress. While the mechanisms that underlie this mitochondria plasticity are unknown, it is clear that
such transitions are not associated with decreased neuronal survival. Further, while the role of
mitochondrial remodelling in physiological conditions remains to be elucidated, the capacity to remodel
may play a key role in neuronal ability to recover from insults that target mitochondria.
Acknowledgements. NSERC and CIHR to LRM; and OGS to NS and MSA to NS. The authors thank Alexandra
Haggard for her help with the mitochondrial experiments and Chloe Shniffer for her help in preparing this
manuscript.
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