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\begin{document}

\title{VPS8D, a CORVET subunit, is required to maintain the contractile vacuole
complex in Tetrahymena thermophila}



\author[1]{Chao-Yin Cheng}%
\author[1]{Josefina Hernández}%
\author[1]{Aaron P. Turkewitz}%
\affil[1]{University of Chicago Department of Molecular Genetics and Cell Biology}%


\vspace{-1em}



  
  \date{June 14, 2024}


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\selectlanguage{english}
\begin{abstract}
Contractile vacuole complexes (CVCs) are complex osmoregulatory
organelles, with vesicular (bladder) and tubular (spongiome)
subcompartments. The mechanisms that underlie their formation and
maintenance within the eukaryotic endomembrane network are poorly
understood. In the Ciliate \emph{Tetrahymena thermophila}, six
differentiated CORVETs (class C core vacuole/endosome tethering
complexes), with Vps8 subunits designated A-F, are likely to direct
endosomal trafficking. Vps8Dp localizes to both bladder and spongiome.
We show by inducible knockdown that \emph{VPS8D} is essential to CVC
organization and function. \emph{VPS8D} knockdown increased
susceptibility to osmotic shock, tolerated in the wildtype but
triggering irreversible lethal swelling in the mutant. The knockdown
rapidly triggered contraction of the spongiome and lengthened the period
of the bladder contractile cycle. More prolonged knockdown resulted in
disassembly of both the spongiome and bladder, and dispersal of proteins
associated with those compartments. In stressed cells where the normally
singular bladder is replaced by numerous vesicles bearing bladder
markers, Vps8Dp concentrated conspicuously at long-lived inter-vesicle
contact sites, consistent with tethering activity. Similarly, Vps8Dp in
cell-free preparations accumulated at junctions formed after vacuoles
came into close contact. Also consistent with roles for Vps8Dp in
tethering and/or fusion were the emergence in knockdown cells of
multiple vacuole-related structures, replacing the single bladder.%
\end{abstract}%



\sloppy


\textbf{Introduction}

Contractile vacuole complexes (CVCs) are a morphologically diverse set
of organelles that are found very broadly throughout eukaryotes, and
particularly in eukaryotic protists. Their recognized role is in
osmoregulation, a critical issue for single-celled organisms inhabiting
freshwater and therefore hyperosmotic environments because constant
passive water uptake across the plasma membrane threatens cellular via
viability, including maintaining physiological cytoplasmic ion
concentrations (Naitoh et al., 1997, Tominaga et al., 1998a, Klauke and
Plattner, 2000, Kissmehl et al., 2002, Tani et al., 2002, Reuter et al.,
2013). Notwithstanding their heterogeneity, CVCs all appear built on
similar organizing principles. They are comprised of one or more large
vacuolar compartments that function as water-storing bladders, that are
ringed by a network of tubular extensions called the spongiome (Allen
and Naitoh, 2002). The existing data are consistent with the general
idea that cytoplasmic water crosses the spongiome membranes via
aquaporins or similar channels and driven by gradients dependent in part
on proton-pumping V\textsuperscript{+}-ATPases. Water in the spongiome
lumen diffuses to swell the connected bladder. At some point, the
bladder membrane fuses with but is not integrated into the plasma
membrane, and the soluble contents are rapidly propelled outside the
cell by dramatic contraction of the bladder. This membrane fusion event
in the Amoebozoan \emph{Dictyostelium discoideum} is regulated at least
in part by triggered calcium release from the bladder lumen, which acts
to downregulate a CVC-specific Rab (Parkinson et al., 2014). More
generally, ion mobilization from the CVC lumen and/or excretion during
bladder contraction may play homeostatic and/or signalling roles
(Ladenburger et al., 2006, Malchow et al., 2008, Ladenburger et al.,
2009, Patel and Docampo, 2010, Sivaramakrishnan and Fountain, 2012).
Following bladder contraction, the fusion pore to the plasma membrane
reseals and the bladder is ready for re-filling. Though functionally
similar, the precise structures involved in this cycle and their dynamic
interactions may differ in important details between lineages. For
example, in the Ciliate Paramecium the reticular spongiome is comprised
of two distinct zones, and the contractile cycle includes membrane
fission and fusion between the spongiome and bladder (Allen, 2000),
neither of which features in the CVC of the
Kinetoplastid\emph{Trypanosoma cruzi} (Jimenez et al., 2022).

CVCs are not found in yeasts or other lineages in which cells secrete
and assemble external walls that can counterbalance osmotic pressures
with physical restraints, nor in multicellular organisms like animals
where the tonicity of the extracellular milieu is under organismal
control. Since the most commonly employed model organisms belong to
these groups, CVCs have been relatively understudied and consequently
many central issues remain unsettled. One largely open question is how
the CVC is integrated with other pathways of eukaryotic membrane
trafficking, including the mechanisms that underlie its maintenance in
vegetative cells and inheritance during cell division. A number of
proteins whose homologs are involved in membrane trafficking in fungi
and animals, have been physically and/or functionally associated with
the CVC in diverse eukaryotic lineages. Early work in \emph{D.
discoideum} linked both and the clathrin assembly factor AP180 to CVC
function, suggesting a putative connection between endocytic trafficking
and CVC maintenance (Wen et al., 2009, Macro et al., 2012). The CVC is
nonetheless normally isolated from bulk endocytic trafficking, in
both\emph{Dictyostelium} and the Ciliate \emph{Tetrahymena thermophila}
, based on studies using endocytic tracers (Gabriel et al., 1999, Cheng
et al., 2023). Whether the same trafficking pathways maintain the CVC in
all lineages remains an open question. A related but more far-reaching
question is whether CVCs emerged independently in different lineages, or
whether they stem from a single ancestral origin. The fact that all
known CVCs rely on vacuolar ATPases might argue for the latter, but
vacuolar ATPases function in a very broad range of compartments in the
endomembrane network (Stevens and Forgac, 1997).

In addition to clathrin and AP180, other endosomal protein homologs were
also shown to be physically and/or functionally associated with the CVC
in Dictyostelium, including AP2, epsin, the endosomal synaptobrevin
homolog Vamp7B, Rabs 11 and 8, and an amphiphysin (I-BAR) family protein
(Harris et al., 2001, Stavrou and O'Halloran, 2006, Wen et al., 2009,
Linkner et al., 2014, Manna et al., 2023, Du et al., 2008, Essid et al.,
2012, Macro et al., 2012). Homologs for a subset of these, including a
Rab11 family member, were subsequently found associated with the CVC
in\emph{Trypanosoma cruzi} (Ulrich et al., 2011). The Ciliates are a
third, very distantly related lineage with prominent CVCs, about which
relatively little is known about membrane trafficking determinants.
Comprehensive localization surveys of SNAREs (in \emph{Paramecium
tetraaurelia} ) and of Rabs (in \emph{T. thermophila} ) provided only
ambiguous evidence addressing whether the CVC is an endosome-related
compartment, since in both protein families the most prominent
CVC-localized members did not belong to conserved endosomal subgroups
but instead either belonged to other identifiable subgroups or were
lineage-restricted (Bright et al., 2010, Plattner, 2013, Schonemann et
al., 2013). A possible exception came from the observation that
several\emph{T. thermophila} Rabs belonging to conserved
endocytic/endosomal subgroups showed secondary localization to the CVC,
but with unknown functional significance (Bright et al., 2010). In
contrast, strong evidence of an endosomal connection in ciliates came
from analysis of\emph{T. thermophila} Dop1p, a member of the DOPEY
family of proteins that serve endosome-related roles in other lineages.
In \emph{T. thermophila} , Dop1p localizes strongly to the CVC where it
is required for normal bladder discharge (Cheng et al., 2016, Cheng et
al., 2023).

In this manuscript, we present analysis of \emph{T. thermophila} VPS8D,
a subunit of one of six \emph{T. thermophila} CORVET (class
C\textbf{cor} e vacuole/\textbf{e} ndosome-\textbf{t} ethering)
complexes. In fungi, animals, and plants, CORVET and the related HOPS
(\textbf{ho} motypic fusion and \textbf{p} rotein \textbf{s} orting)
complexes are key determinants in endo-lysosomal trafficking (van der
Beek et al., 2019). Current data support models in which the
hetero-hexameric complexes tether two vesicle membranes by binding Rab
GTPases in each of the adjoining membranes, and catalyze fusion by
interacting with SNAREs (Baker et al., 2015). While many organisms have
one CORVET and one HOPS, in the lineage leading to \emph{T.
thermophila}HOPS was lost and CORVET expanded to a family of six
complexes, each with a unique subunit composition and cellular
localization (Sparvoli et al., 2018). One of these complexes, which
uniquely contains the subunit Vpd8D, was localized in an initial survey
to the CVC, and subsequently shown to be present in a punctate
distribution at both the bladder and spongiome, in dynamic equilibrium
with a cytosolic pool (Sparvoli et al., 2020, Cheng et al., 2023).
Attempts to study the function of Vpd8Dp by gene knockout were
unsuccessful, since unlike the Vps8 subunits in other CORVET complexes
the \emph{VPS8D} paralog appeared to be essential (Sparvoli et al.,
2018). Here we have therefore taken a different approach, using a
gene-specific RNA hairpin to partially deplete the\emph{VPS8D}
transcript. The knockdown cells show a variety of CVC-associated
phenotypes. The severity of the defects appears to correlate with the
intensity of the knockdown, and at the extreme manifests as complete
loss of both the bladder and spongiome. Both our functional and
localization data are consistent with the idea that in\emph{T.
thermophila} a ciliate-adapted CORVET complex directs critical membrane
tethering and fusion cycles in the CVC.

\textbf{Results}

A cartoon illustrating the anatomy of the CVC in \emph{T. thermophila} ,
together with a small set of proteins whose distributions were
determined by \emph{in vivo} localization studies of endogenously tagged
genes, is shown in Fig. 1A. As previously reported (Sparvoli et al.,
2020, Cheng et al., 2023), the CORVET subunit Vps8D, here endogenously
tagged with mCherry, localizes to both the bladder and spongiome (Fig.
1A, right). Similarly, endogenously tagged CORVET subunit Vps11p
localizes to the CVC, but also to other cellular loci as expected since
it is a subunit of multiple CORVET complexes in this organism (Sparvoli
et al., 2020) (Fig. S1).

\textbf{RNAi-mediated knockdown of \emph{VPS8D} results in defects
consistent with a role at the osmoregulatory CVC}

The role of Vps8Dp at the CVC is not known, but the gene appeared to be
essential since we could not recover viable cells when pursuing a gene
knockout strategy (Sparvoli et al., 2018). We therefore chose to pursue
a gene knockdown approach, by constructing an RNA hairpin corresponding
to a part of the VPS8D gene (Fig. 1B) and expressing it
in\emph{Tetrahymena} under the control of the
cadmium-inducible\emph{MTT1} promoter (Shang et al., 2002, Howard-Till
and Yao, 2006). The effectiveness of such hairpins in \emph{T.
thermophila} can be difficult to assess at the level of the targeted
transcripts (Howard-Till and Yao, 2006). To circumvent this issue, we
directly measured protein levels by generating a cell line in which
Vps8Dp was endogenously tagged with the FLAG epitope, and the
\emph{VPS8D} hairpin was inducibly expressed from the \emph{RPL29} gene
locus (Yao and Yao, 1991) (Fig. 1C). Induction of the hairpin expression
led to a significant decrease in the level of Vps8Dp-FLAG, as measured
by immunofluorescence (Fig. 1D). Importantly, after hypoosmotic shock,
the knockdown cells displayed morphological changes consistent with a
defect in maintaining osmotic homeostasis, losing their teardrop shape
and becoming roughly spherical (Fig. 1E) Equivalently-treated wild type
cells could quickly recover their normal shape (Fig. 1E). Similarly,
while the wildtype cells retained full viability after the osmotic
challenge, the \emph{VPS8D} knockdown cells died shortly thereafter
(Fig. 1F). The differences in the cell responses after hypoosmotic shock
between wild type and \emph{VPS8D} -KD cells are diagrammed in Fig. 1G.
Under these stress conditions, the activity of the CVC appears critical
not to prevent osmotic swelling but to allow recovery after such events.

\emph{VPS8D} \textbf{knockdown affects features of both the bladder and
spongiome}

Conventional CORVET in \emph{S. cerevisiae} acts as a determinant in the
maturation of endosomal compartments (Peplowska et al., 2007). To ask
whether knockdown of \emph{T. thermophila} \emph{VPS8D} led to changes
in the CVC, we generated cells expressing the \emph{VPS8D} hairpin in
combination with endogenously tagged copies of either Dop1p, which most
strongly labels the bladder, or tagged Vma4p which decorates the
spongiome (Cheng et al., 2023).

Following 5 hours of hairpin induction, Dop1p-mNeon still shows clear
localization to the CV bladder that is qualitatively similar to that in
non-induced or wildtype cells, and the bladder profiles are also similar
(Fig. 2A, compare left and right panels). However, by 8 hours of hairpin
induction the intensity of Dop1p bladder steady state labeling was
significantly decreased (Fig. 2B and S2). This change in the association
between Dop1p and the bladder membrane upon \emph{VPS8D} knockdown could
also be seen at the level of dynamics. We previously used fluorescence
recovery after photobleaching (FRAP) to discover that Dop1p at the
bladder exchanges with a cytosolic pool (Cheng et al., 2023), as shown
in Figure 2C and 2D. That exchange was inhibited in
\emph{VPS8D}knockdown cells, so that the rate of fluorescence recovery
at the bleached bladder was decreased (Fig. 2E, 2F and 2G). Lastly, the
period between contractions was lengthened in these \emph{VPS8D}
knockdown cells (Fig. 2H). The mechanisms controlling bladder
contraction are not known, but one possibility is that bladder refilling
is less efficient in the mutant. Intriguingly, \emph{VPS8D} hairpin
induction had a clear effect on the spongiome morphology. In particular,
the volume occupied by Vma4p-decorated tubules showed a marked decrease
within 5 hours of\emph{VPS8D} -hairpin induction (Fig. 2I, 2J and S3),
suggesting that spongiome architecture depends more acutely than bladder
structure on the activity of Vps8Dp. Interestingly, these rapid changes
in spongiome structure upon \emph{VPS8D} knockdown are similar to those
seen after cells are exposed to brief hypoosmotic challenge (Fig. S4).

\textbf{Prolonged induction of \emph{VPS8D} hairpin expression leads to
loss of CVC structures}

Following longer periods of \emph{VPS8D} -hairpin induction, the CVC and
associated markers underwent more dramatic changes. Strikingly, many
cells viewed with DIC optics lacked any visible contractile bladders,
strongly suggesting that the bladders are lost during
sustained\emph{VPS8D} knockdown. Not surprisingly, all CVC proteins in
such cells showed a marked change in their distribution compared to in
wildtype. Dop1p no longer localizes to a recognizable bladder but is
instead dispersed throughout the cytoplasm including in many small
puncta (Fig. 3A). Vma4p is also dispersed throughout the cell, visible
as diffuse fluorescence but also present in large, potentially
multi-vesicular structures (Fig. 3B). A second protein that localizes to
the spongiome in wildtype cells, Scr7p, is also delocalized after
extended\emph{VPS8D} -hairpin induction. The delocalization of these two
spongiome markers is likely to reflect the loss of the spongiome, in
parallel to the loss of the bladder.

Taken together, these results strongly suggest that the maintenance of
multiple compartments of the CVC depend on Vps8Dp. The spongiome may be
more acutely dependent on Vps8Dp-assisted membrane trafficking than the
bladder.

\emph{VPS8D} \textbf{knockdown results in a proliferation of large
vesicles or vacuoles bearing CV markers}

If Vps8Dp as part of a CORVET complex is active in promoting fusion
between membranes, then the disappearance of CVC structures
upon\emph{VPS8D} knockdown may be due to a fusion defect between
intermediates in CVC formation. To explore this idea, we analyzed cells
that showed intermediate CVC phenotypes after induction of
the\emph{VPS8D} hairpin. In these experiments, cells were treated with
low concentrations of cadmium and for short time periods. In such
samples, it was easy to find cells displaying CVC features never seen in
wildtype cells, with examples in Figures 4A (wildtype) vs the knockdown
cells (Fig. 4B and 4C). The cell shown in panel B possesses what appear
to be two adjacent CV bladders. These persist for tens of seconds before
one of them contracts rapidly (t = 28sec), which is then followed by
contraction of the second bladder at 29 sec. This asynchronicity implies
that the bladders are linked with two different plasma membrane pores.
Over the next 20 seconds, just one of the bladders appears to refill. In
panel C, the cell contains at least seven medium sized-to-large
Dop1p-labeled vesicles. Over the course of the 15 seconds in the video,
one of the two largest vesicles contracts, followed shortly thereafter
by the second. Within 5-6 seconds, both show signs of refilling and also
appear tightly associated with smaller labeled vesicles at their
periphery. The Dop1p-labeled relatively immobile vesicles in this cell
are spread over a large volume of the cell posterior, so that most
cannot be closely associated with CV pores. The presence in a large
subset of the knockdown cells of heterogeneous Dop1p-labeled
compartments, which include multiple contractile bladders, may be
consistent with the idea that maintenance of the CVC in wildtype cells
requires Vps8Dp-dependent membrane fusion.

\textbf{Vps8Dp is concentrated at junctions between CV-related
membranes}

In budding yeast, the HOPS complex concentrates at the junctions between
membranes destined for fusion during vacuole maintenance (Wang et al.,
2003).To see whether Vps8Dp in \emph{Tetrahymena} behaves similarly, we
took advantage of the observation that many cells exposed to hypoosmotic
stress showed fragmented CVC structures. As shown in Figure 5A, such
cells contain multiple Dop1p-labeled vacuole-like compartments,
concentrated in the cell posterior and undergoing frequent fusion to
form larger compartments, in addition to what appears to be the central
contractile bladder. Dop1p itself appears to be irregularly distributed,
often as puncta, around the periphery of the labeled compartments.
Interestingly, the CVC similarly fragments into multiple Dop1p-decorated
large vacuole-like compartments under conditions of \emph{hyper} osmotic
stress, though with the difference that no contractile central bladder
is visible (Fig. 5B).

Vacuole-like compartments formed under hypoosmotic conditions are also
decorated with Vps8Dp (Fig. 5C). The Dop1p-labeled and Vps8Dp-labeled
vacuoles appear to be the same structures, since there is strong
co-localization of the two proteins at such vacuoles in cells expressing
both Dop1p-mNeon and Vps8Dp-mCherry (Fig. S5). However, a notable
difference between the Dop1p and Vps8Dp distributions is that in many
cases the latter appears strikingly concentrated in regions between
adjoining vacuoles, where the membranes come into close contact (Fig.
5C).

The concentration of Vps8Dp-mNeon between adjoining vacuoles/vesicles
could be seen even more strikingly in some osmotically-stressed samples
in which cells occasionally suffered localized plasma membrane rupture
under the pressure from the microscope cover slip. These breaks allowed
leakage of the large Vps8Dp-labeled vesicles into the surrounding
buffer. In such samples, Vps8Dp could be seen highly concentrated at
junctures between the membranes, consistent with a role in tethering and
fusion (Fig. 5D). Moreover, in some cases we could capture membrane
fusion occurring at Vps8Dp-enriched junctions (Fig. 5D, arrowheads).
Interestingly, we also captured episodes in which two of the leaked
vesicles shifted closer to one another, and Vps8Dp accumulated rapidly
at their meeting site just when they came into contact (Fig. 5E and 5F).

Discussion

Genes encoding CORVET subunits underwent a marked expansion in the
lineage leading to Tetrahymena, leading to the six distinct complexes
in\emph{T. thermophila} called 8A-8F based on their unique Vps8 subunit
(Klinger et al., 2013, Sparvoli et al., 2018, Sparvoli et al., 2020).
These complexes are distinct from one another in their localization and
their subunit makeup (Sparvoli et al., 2020). They are accordingly
likely to be individually specialized for distinct functions. We
confirmed this previously for the 8A complex, which we found to be
essential for the formation of lysosome-related organelles called
mucocysts (Sparvoli et al., 2018).

In this manuscript we report analysis of the Vps8 subunit of the 8D
CORVET. Vps8Dp was previously shown to be strongly associated with the
osmoregulatory CVC (Sparvoli et al., 2020, Cheng et al., 2023). Because
the gene was found to be essential for cell viability and therefore
difficult to analyze via gene knockout (Sparvoli et al., 2018), in this
manuscript we instead used a knockdown approach based on induced
expression of a gene-specific RNA hairpin (Howard-Till and Yao, 2006).
One striking finding was that even relatively minor depletion of Vps8Dp
produced cells that were highly sensitive to osmotic shock (Fig. 1).
More informatively, following hairpin induction the cells showed
time-dependent changes in the morphology and functioning of the CVC,
affecting both the spongiome and bladder.

Within the cohorts of knockdown cells in each experiment there was
significant phenotypic heterogeneity, making it difficult to rigorously
establish the order in which different defects were manifest.
Nonetheless, our results suggest that spongiome structure may be more
sensitive to \emph{VPS8D} dosage compared with the bladder structure,
since at early timepoints following hairpin induction only the spongiome
was reduced in size relative to the wildtype, a response that became
more pronounced at longer time points (Fig. 2J). Interestingly, this
shrinkage of the spongiome is very similar to the response seen when
cells are briefly exposed to hypoosmotic shock (Cheng et al., 2023). The
bladder size in contrast did not detectibly change, but contractions
grew less frequent (Fig. 2G). Since the bladder fills with water
transported from the spongiome, the slowed contraction rate may reflect
less efficient filling from a partially disabled water-collecting
reticulum. The bladder in knockdown cells at these early time points
also showed reduced labeling by Dop1p, with a corresponding increase in
diffuse Dop1p signal in the cytoplasm (Fig. 2A and 2B). The mechanistic
role of Dop1p is not yet understood, but in its absence cells show only
infrequent contraction of an enormously expanded bladder (Cheng et al.,
2016, Cheng et al., 2023).

At later time points following hairpin induction, neither the bladder
nor spongiome was detectable, indicating that \emph{VPS8D} is essential
for the maintenance of this organelle (Fig. 3). Proteins that in
wildtype cells localized strongly to the CVC were instead found in
dispersed cytoplasmic puncta. This process may be reversible, based on
our observation that some cells in knockdown cultures recover when
hairpin expression is de-induced. Studying the mechanisms underlying
such recovery may help to shed light on trafficking pathways involved in
CVC biosynthesis, which are poorly understood in any lineage and
particularly in Ciliates. The disappearance of both bladder and
spongiome compartments after prolonged \emph{VPS8D} knockdown is similar
to a phenotype reported in \emph{D. discoideum} that is linked to a
protein in the BEACH family, called LvsA (Gerald et al., 2002).
Interestingly, pulldown experiments in the Apicomplexan \emph{Toxoplasma
gondii} suggest physical interactions between CORVET/HOPS and a
BEACH-domain containing protein, but in the context of secretory
organelle biogenesis (Morlon-Guyot et al., 2018).

The established role of CORVETs, based on studies in yeast and animals,
is in tethering and fusion of membrane compartments (Spang, 2016, van
der Beek et al., 2019). Our imaging data are consistent with such a role
for the Vps8Dp-containing CORVET at the CVC. For our imaging, we
exploited conditions in which the CVC bladder, rather than existing as a
single large vacuole, is replaced by a set of smaller vacuoles that can
still be recognized due to their labeling by Dop1p as well as Vps8Dp. We
found that Dop1p has a relatively uniform distribution at the periphery
of such vacuoles, while in contrast Vps8Dp is concentrated at sites
where neighboring vacuoles come into close contact (Fig. 5). In some
cases, these were seen to be sites of subsequent fusion. A similar
phenomenon has been reported of specific protein and lipid accumulation
at the interface between vacuoles undergoing homotypic fusion in yeast,
termed the vertex ring domain, and it includes the HOPS complex (Wang et
al., 2003, Fratti et al., 2004). Interestingly, we witnessed the same
highly polarized Vps8Dp distribution on small Vps8Dp-decorated vacuoles
that were extruded through breaches in the plasma membrane of perforated
cells. In one case, we witnessed rapid Vps8Dp redistribution at the
perimeters of two initially widely-spaced vacuoles when they came into
contact, leading to accumulation at the junction (Fig. 5E). This may be
explained if the 8D-CORVET complexes on each surface are bound to
integral membrane proteins undergoing rapid 2-dimensional diffusion, but
any complex that simultaneously engages with integral membrane proteins
on an adjacent membrane diffuses more slowly. Based on analysis of the
vertex ring complex in yeast, this partitioning is also likely to
involve lipid subdomains (Fratti et al., 2004).

Our images from osmotically-stressed cells are consistent with a role
for Vps8Dp in homotypic tethering and fusion at the CVC, and therefore
support the paradigm for CORVET and HOPS established in other
eukaryotes. However, no Vps8Dp-labeled vertex ring domains are apparent
in cells grown under non-stress conditions, and moreover there is no
step clearly involving homotypic fusion in the
\emph{Tetrahymena}contractile cycle (Cheng et al., 2023). In Paramecium,
electrophysiological data support a model in which the junctions between
the bladder and emanating spongiome arms undergo fission and subsequent
fusion with each contractile cycle (Tominaga et al., 1998b). Whether
similar fission/fusion cycles occur in Tetrahymena, where the structure
of the bladder/spongiome junction appears quite different, is unknown
(Elliott and Bak, 1964). The role of Vps8Dp is unlikely to be restricted
to either homotypic fusion between bladder-derived vesicles or
spongiome-bladder fusion, since Vps8Dp localizes to puncta throughout
the spongiome (Cheng et al., 2023). Spongiome tubules show dynamic
extension and branching (Cheng et al., 2023), and one possibility is
that the Vps8Dp-containing CORVET complex facilitates homotypic fusion
between branches in a way that is required to maintain a cohesive
reticulum. The retraction of the spongiome at early time points
following \emph{VPS8D} knockdown may be consistent with this scenario.

However, another broad possibility is that Vps8Dp has activities
independent of a conventional CORVET complex, as a monomer or a smaller
sub-complex, for which examples exist in other eukaryotes for CORVET and
HOPS (Asensio et al., 2013, Lorincz et al., 2016). In pulldowns, Vps8Dp
is associated with the 5 other expected subunits of CORVET (Sparvoli et
al., 2020). However, some details suggest it may be an outlier among
the\emph{Tetrahymena} CORVET complexes. First, it is the most
biochemically differentiated, including unique subunit variants for
Vps8, Vps16, Vps18, and Vps33 (Sparvoli et al., 2020). Secondly, Vps8Dp
is roughly 200 amino acids longer than the other Vps8 paralogs in
Tetrahymena, with multiple insertions suggesting the potential for
additional interactions and activities. Lastly, Vps8Dp in pulldowns
appeared by silver staining to be significantly more abundant than the
other subunits with which it associates; significantly, no similar
imbalance was present for the Vps8 subunits of the other five CORVETs
(Sparvoli et al., 2020). Since the affinity tag for these pulldowns was
attached to Vps8Dp, the greater relative abundance of Vps8Dp could be
explained if that particular CORVET were comparatively unstable.
Alternatively or in addition, it could reflect a super-stoichiometric
pool of Vps8Dp in cells. On this basis, we speculate that Vps8Dp could
potentially be playing distinct roles at the bladder vs. spongiome, as
part of different complexes.

\textbf{Materials and Methods}

\textbf{Cell strains and culture conditions}

\emph{Tetrahymena thermophila} strains used in this work are listed in
Table S1 with culture conditions as previously described (Gorovsky et
al., 1975). Cells were grown in SPP medium (2\% proteose peptone (GIBCO,
211684), 0.1\% yeast extract (BD, 212750), 0.2\% dextrose (ACROS,
41095-5000) and 0.003\% EDTA, ferric-sodium salt) supplemented with 100
\selectlanguage{greek}μ\selectlanguage{english}g ml\textsuperscript{-1} Normocin\textsuperscript{TM} (InvivoGen), an
antimicrobial reagent. Cells were grown at 30\selectlanguage{ngerman}°C with agitation at
100\selectlanguage{english} rpm. Cell densities were measured using a spectrophotometer~(Thermo
Spectronic Unicam), where~OD\textsubscript{550}~=~1~corresponds
to~\textasciitilde{} 1~\emph{x}
~10\textsuperscript{6}~cells~ml\textsuperscript{-1} (David L. Spector,
1998)~or a Z1 Coulter Counter (Beckman Coulter Inc., Indianapolis,
Indiana). Cultures were generally used during the log phase of cell
growth (cell density 2-4 \(\times\)10\textsuperscript{5} cells
ml\textsuperscript{-1}). Reagents were from Sigma-Aldrich unless
otherwise noted.

\textbf{Biolistic Transformation}

50 ml~\emph{Tetrahymena} ~cultures were grown to 500,000 cells
ml\textsuperscript{-1} and starved for 10--16 h in 10\selectlanguage{english} mM Tris-HCl
buffer, pH 7.4, at 30degC with agitation. Biolistic transformations were
performed as described previously (Sparvoli et al., 2018, Cassidy-Hanley
et al., 1997). Transformants were identified after 3 days selection in
paromomycin sulfate (120 \selectlanguage{greek}μ\selectlanguage{english}g ml\textsuperscript{-1} with 1 \selectlanguage{greek}μ\selectlanguage{english}g
ml\textsuperscript{-1} of CdCl\textsubscript{2}), and then serially
passaged 5\(\times\)/week for \textasciitilde{}4 weeks in
decreasing concentrations of CdCl\textsubscript{2}~and increasing
concentrations of paromomycin sulfate.

\textbf{Generation of \emph{VPS8D} knockdown strains}

To generate \emph{VPS8D} knockdown strains, a construct was designed as
shown in Fig. 1B, by first PCR amplifying \textasciitilde{}500 bp of
genomic \emph{VPS8D} gene coding sequence with two sets of primers, one
set engineered with ApaI and XhoI restriction sites and another set with
PmeI and SmaI restriction sites. These two fragments were cloned into
PCRII-I3 so that they were inverted relative to one another. The rDNA
backbone vector pIBF (Howard-Till and Yao, 2006) was digested with with
ApaI and PmeI to release the BFP fragment. In parallel, the
\emph{VPS8D}hairpin cassette was excised from the PCRII-I3 backbone with
ApaI and PmeI, and the \textasciitilde{}1kb insert then gel purified and
ligated into the site of the excised BFP fragment in the pIBF backbone.
Approximately 20\selectlanguage{greek}µ\selectlanguage{english}g of this plasmid was used per biolistic
transformation. In addition to the rDNA-based \emph{VPS8D} hairpin
strains, we also engineered a construct to target the
\emph{VPS8D}hairpin cassette to the macronuclear \emph{RPL29} locus. In
this case, the\emph{VPS8D} hairpin cassette was cloned into
pBSICC-rpl29CyR (Yao and Yao, 1991) for biolistic transformation
followed by selection for cycloheximide resistance. In both \emph{VPS8D}
hairpin constructs, hairpin expression is under the control of the
cadmium-inducible\emph{MTT1} promoter (Shang et al., 2002).

\emph{Assaying VPS8D} \textbf{knockdown phenotypes}

To measure survival rates after hypoosmotic shock, wildtype
and\emph{VPS8D} -KD strains (3 of each, with independent isolates of the
mutant strain) were grown to log phase in 10 ml SPP. Cell densities were
measured and equalized, and 0.5ml aliquots were diluted 1:100 in 50 ml
SPP medium with or without 1 \selectlanguage{greek}µ\selectlanguage{english}g/ml CdCl\textsubscript{2}. Cells were
grown for 16 hours at 30\selectlanguage{ngerman}°C with agitation. Culture densities were
measured, and then cells were pelleted by centrifugation in a clinical
centrifuge for 1 minute and resuspended gently in 10 mM Tris-HCl buffer,
pH 7.4. After an overnight incubation, culture densities were once again
measured to calculate survival rates. For experiments monitoring changes
in CVC markers, the \emph{VPS8D} -KD strains endogenously expressing
Dop1p-mNeon, Vma4p-mNeon or Scr7p-mNeon were grown as above in
non-inducing or inducing conditions. Cells were withdrawn and fixed for
imaging after 3, 5 and 8 (Fig. 2) or 16h (Fig. 3).

\textbf{Acute osmotic challenge}

Cells expressing tagged proteins were grown to log growth phase, then
pelleted by centrifugation in a clinical centrifuge for 1 min and gently
resuspended in 22°C 10 mM Tris-HCl buffer, pH 7.4 (hypoosmotic) or 10 mM
sorbitol (hyperosmotic). The cells were then immediately immobilized for
live imaging.

\textbf{Endogenous expression of FLAG-tagged Vps8Dp and mNeon-tagged
genes in \emph{VPS8D} knockdown strains}

pVPS8D-FLAG-ZZ-Neo4, pDOP1-2mNeon-6myc-Neo4, pVMA4-2mNeon-6myc-Neo4 and
pSCR7-2mNeon-6myc-Neo4 (Cheng et al., 2023) were introduced by biolistic
transformation into strains in which the \emph{VPS8D} hairpin expression
cassette was already integrated at the \emph{RPL29} locus, as described
above.

\textbf{Cell immobilization for live imaging}

We used two methods to immobilize cells for live imaging. For both
methods, we prepared a single slide at a time and viewed it immediately.
The first method relied on the pressure exerted on small volume samples
under a cover slip. Cells were first concentrated by centrifugation to
2-5 x 10\textsuperscript{6} cells ml\textsuperscript{-1}. 6 \selectlanguage{greek}μ\selectlanguage{english}l of sample
was applied to the slide, and immediately overlayed with a 22 X 22 mm
cover slip. The resulting pressure frequently results in cell
immobilization, which could be verified by monitoring ciliary beating on
the cell surface at the same time that cell viability could be monitored
by observing the periodic contraction of the contractile vacuole, both
in the DIC channel. For the second method, cells were pelleted and
resuspended in a thermoreversible gel on ice, whose viscosity increases
as it warms to room temperature. Here, cells at the same high density
were mixed well with CyGEL\textsuperscript{TM} (ab109204, ABCAM) (the
mix ratio optimized for each experiment) and immediately mounted with a
cover slip. This approach results in many cells being effectively
immobilized in the thin gel. For both methods, we carefully watched for
proliferation of large cytosolic vesicles. Growing cells have large food
vacuoles derived from the oral apparatus. No new food vacuoles form in
immobilized cells, so any substantial increase in the number of such
vesicles is likely to represent autophagosome formation as part of a
stress response. We rejected any cells showing such an increase.

\textbf{Cell fixation for imaging}

Cells were fixed in a final concentration of 4\% paraformaldehyde in 10
mM Tris-HCl buffer, pH 7.4 (stock: 16\% paraformaldehyde solution in
distilled water, EM grade, 15710, Electron Microscopy Science) for 10-30
minutes at room temperature. They were then washed repeatedly in PBS to
reduce the background autofluorescence, before staining with DAPI
(4',6-diamidino-2-phenylindole) at final concentration 50 ng
ml\textsuperscript{-1} for 10 minutes for nuclear staining, and/or
staining with antibodies for immunofluorescence. Images were captured by
using a Carl Zeiss Microscope stand Axio Observer 7 system or Marianas
Yokogawa-type spinning disc inverted confocal microscope.

\textbf{Fluorescence Recovery after Photobleaching (FRAP)}

Cells were immobilized as described above. FRAP experiments were
performed using a Marianas Yokogawa-type spinning disc inverted confocal
microscope with a 100x/NA1.45 oil (Alpha Plan-Fluar) objective. The
system featured fast shutter speeds and channel switching for high-speed
imaging and a vector high-speed point scanner for bleaching, controlled
using Slidebook software. For each experiment, we first identified a
well-immobilized cell and verified the fluorescence signal with a
\textasciitilde{}100-500 ms exposure. We then used the time-lapse mode
to capture images with the fastest camera speed for a pre-photobleaching
record and then used the screen selection tool and real-time imaging to
draw a ROI (region of interest) for photobleaching.

\textbf{Image analysis}

The image processing package FIJI was used for image processing and
analysis (Schindelin et al., 2012). Processing of raw images included:
image cropping and rotation; bleach correction; background subtraction;
adjustment of brightness/contrast; color switching; selection of images
from z-stacks; intensity threshold adjustments; measurements of
parameters including areas, mean gray values, and integrated densities.

To quantify Dop1p-mNeon fluorescence intensity in the CV bladder vs
whole cell, all images were taken using the same image capturing
settings. The raw images were used to select the ROIs of the area of
Dop1p-labeled bladder and whole cell. The mean fluorescence intensities
of these ROIs were then measured. To quantify Vma4p-mNeon localized
spongiome distribution, the raw images were used to measure the area of
Vma4p-labeled spongiome and whole cell. For FRAP analysis, the
time-lapse images were separately adjusted by bleach correction for the
pre-photobleaching series and after-photobleaching series. The intensity
thresholds were adjusted for background subtraction, and intensity
changes over time were measured in ROIs. Intensities were plotted using
GraphPad Software Prism.

\textbf{Immunofluorescence analysis}

Cells (2\selectlanguage{ngerman}×10\textsuperscript{5}) were fixed with 4\% paraformaldehyde for
30 minutes at room temperature, and immunolabeled as previously
described (Briguglio et al., 2013, Bowman and Turkewitz, 2001). Briefly,
cells were first incubated for blocking with 5\% BSA (bovine serum
albumin, BP1600-1 Fisher Scientific) in PBT (0.3\% Triton X-100 (ACROS)
in PBS) for one hour at room temperature and then incubated with the
primary antibodies in the same blocking buffer. Anti-FLAG M2 antibody
staining was at 1:1000 (Monoclonal ANTI-FLAG M2 antibody, Sigma F1804)
for overnight at 4°C, followed by three washes in 0.3\% Triton X-100 in
PBS. The secondary antibodies were Alexa Fluor 488-conjugated anti-mouse
IgG, incubated for one hour at room temperature (1:1000, Alexa Fluor 488
Goat anti-Mouse IgG, Invitrogen A-11001), and similarly washed. Cell
nuclei were stained with 4\selectlanguage{english}',6-diamidino-2-phenylindole (DAPI) (50 ng
ml\textsuperscript{-1}). Sample was mounted on microscopic slides for
imaging.

\textbf{Acknowledgements}

At the University of Chicago, we thank Vytas Bindokas and Christine
Labno for help at the Integrated Light Microscopy Core Facility. In the
Turkewitz laboratory, we thank Patrick Jiang and Maya Waarts for helpful
discussions.

\textbf{Competing interests}

The authors declare no competing or financial interests.

\textbf{Funding}

Work in APT's laboratory was supported by the National Institutes of
Health (NIH) (GM105783) and the National Science Foundation (MCB
1937326).

Figure legends

Figure 1:

\emph{VPS8D} \textbf{plays an important role in cellular resistance to
osmotic stress.}

A, left: CVC anatomy in \emph{T. thermophila} . The cartoon shows
previously established features of the Tetrahymena CVC including one
central bladder near the cell posterior, the surrounding
tubulo-vesicular spongiome network, and two pores connecting the bladder
to the plasma membrane. Dop1p (blue) primarily labels the bladder, and
Vma4p (green) exclusively labels the spongiome. Also shown are the oral
apparatus (OA) in the anterior end of cell, and both the micronucleus
(MIC) and macronucleus (MAC) near the cell midpoint. The vertical rows
represent the cytoskeletal tracks called primary meridians, along which
the ciliary basal bodies are spaced. For clarity, the cilia themselves
are only shown at the cell edges.

A, right: As previously reported, Vps8Dp localizes to the CVC as well as
to cytosolic puncta. Growing cells expressing Vps8Dp endogenously tagged
with mCherry were fixed and stained with DAPI. Images were taken with a
Marianas spinning disc confocal microscope.

B: \emph{VPS8D} RNAi knockdown strategy via hairpin expression. Cell
lines transformed with the hairpin construct, under the control of the
inducible MTT1 promoter, are called \emph{VPS8D} -KD.

C: \emph{VPS8D} expression is reduced in cells expressing
the\emph{VPS8D} hairpin. The accumulation of Vps8Dp-FLAG, in cells
expressing the tagged protein from the endogenous locus, was measured by
anti-FLAG immunostaining. The labels ``-Cd'' and ``+Cd'' indicate the
presence or absence of cadmium, which induces \emph{VPS8D} hairpin
expression. Images were taken with a Zeiss Axio Observer 7 system. Each
image was generated from10 z-sections across the CVC using the Z project
tool in FIJI software.

D: Quantification of Vps8Dp-FLAG accumulation without/with
hairpin-induced knockdown. 10 images for each condition (+Cd and -Cd)
were used to quantify the intensity of CVC-localized Vps8Dp
immunofluorescence. The data were plotted to a box-and-whiskers plot
with statistical analysis via two-tailed~\emph{t} -test by GraphPad
Software Prism. The boxes display median and interquartile ranges, and
the whiskers represent the minimum to maximum value of data. The
difference between +Cd and -Cd is significant, \emph{P} -value
\textless{}0.001.

E: \emph{VPS8D} knockdown (\emph{VPS8D} -KD) cells fail to restore
normal cell shape after hypoosmotic-induced swelling. DIC images from
wild type (WT) and \emph{VPS8D} -KD cells are shown. Images were taken
with a Zeiss Axio Observer 7 system.

F: \emph{VPS8D} knockdown cells do not survive hypoosmotic stress. Wild
type (WT) and \emph{VPS8D} -KD cells were incubated with (+Cd) or
without cadmium (-Cd), and then given a hypoosmotic challenge. After
16h, cell viability was measured (see details in Materials and Methods).
The data were plotted using GraphPad Software Prism, using a
two-tailed~\emph{t} -test. Individual data points and the mean+-s.d.
percentage of survival are shown. The \emph{P} -value for the difference
between WT and \emph{VPS8D} -KD is \textless{}0.0001.

G: Differences in the responses after hypoosmotic shock between wild
type and \emph{VPS8D} -KD cells.

Figure 2:

\emph{VPS8D} \textbf{is required for features of the bladder and
spongiome.}

A: Dop1p localization becomes more dispersed upon \emph{VPS8D}knockdown.
Growing cells (WT or \emph{VPS8D-} KD) expressing Dop1p-mNeon at the
\emph{DOP1} locus were imaged to assess Dop1p localization. WT (left
panel); induced \emph{VPS8D} knockdown (right panel). Images were taken
with a Zeiss Axio Observer 7 system.

B: Dop1p localization to the bladder decreases upon
\emph{VPS8D}knockdown. \emph{VPS8D} -KD cells expressing Dop1p-mNeon
were incubated without (-Cd) or with (+Cd) cadmium for 3,5 or 8 hours
and then fixed for imaging. 20 images for each sample were analyzed to
quantify Dop1p fluorescence at the bladder and for the whole cell (Fig.
S2). The data were plotted using GraphPad Software Prism and analyzed
using a two-tailed~\emph{t} -test. The ratios of the mean+-s.d.
intensities of Dop1p fluorescence at the bladders relative to the whole
cells from each sample are shown. The difference between +Cd and -Cd
after 8-hour treatment is significant, \emph{P} -value
\textless{}0.0001.

C-G: Dop1p exchange kinetics are slowed upon \emph{VPS8D} knockdown.

C. Wild type cell expressing Dop1p-mNeon analyzed by FRAP. Growing cells
were immobilized and live imaged with a Marianas spinning disc confocal
microscope with a FRAP tool. Three images were extracted from a video
(Movie 1) showing a photobleaching event and recovery. Left panel: cell
at 11s, immediately before photobleaching. Middle panel: cell at t=12s
immediately after photobleaching a region containing part of the CV
bladder. The bleached area appears as a dark square area outlined with a
red dotted box. Right panel: cell at t=2 min 1s.

D: The recovery after photobleaching data were analyzed by FIJI image
process software (see details in Materials and Methods). The
fluorescence intensity in the photobleached area along with the time was
plotted using GraphPad Software Prism.

E: \emph{VPS8D} knockdown cell expressing Dop1p-mNeon analyzed by FRAP.
Three images were extracted from a video (Movie 2) showing a
photobleaching event and recovery, with panels as in Fig. 2C but with
the bleaching event at t=0:26 and recovery at t=5:33.

F: Recovery after photobleaching, plotted as in Fig. 2D.

G: Dop1p recovery after photobleaching is slowed in
\emph{VPS8D}knockdown cell. Three FRAP experiments obtained from wild
type and\emph{VPS8D} knockdown cells were analyzed and calculated to
obtain the mean+-s.d. recovery rate after photobleaching. The data were
plotted using GraphPad Software Prism and using a two-tailed~\emph{t}
-test. The difference between wild type and \emph{VPS8D} knockdown is
significant,\emph{P} -value \textless{}0.05.

H: Contractile cycle period is increased in \emph{VPS8D} knockdown
cells. The contractile cycle period of wild type cell and
\emph{VPS8D}knockdown cell were measured. 50 cycles from wild type cells
and 22 cycles from \emph{VPS8D} knockdown cells were measured to obtain
the contractile cycle period and the data were plotted using GraphPad
Software Prism and using a two-tailed~\emph{t} -test. The difference
between wild type and \emph{VPS8D} knockdown is significant,\emph{P}
-value \textless{}0.0001.

I: Vma4p localization in WT and \emph{VPS8D} knockdown cells. Growing
cells expressing Vma4p-mNeon were imaged to show Vma4p localization in
WT (left panel) and \emph{VPS8D} knockdown cells (right panel). Images
were taken with a Zeiss Axio Observer 7 system.

J: The spongiome area is reduced in \emph{VPS8D} knockdown
cell.\emph{VPS8D} -KD growing cells expressing Vma4p-mNeon were treated
with cadmium (+Cd) for 3,5 and 8 hours or without cadmium (-Cd) and then
were fixed for imaging. 20 images for each sample were analyzed to
quantify the area of the Vma4p-labelled spongiome and the whole cell
(Fig. S3). The data were plotted using GraphPad Software Prism and using
a two-tailed~\emph{t} -test. The ratio of the mean+-s.d. volume of
spongiome to whole cell from each sample is shown. The difference
between +Cd and -Cd at 5- and 8-hours treatment point are significant,
\emph{P} -value \textless{}0.0001.

Figure 3:

\textbf{Prolonged \emph{VPS8D} hairpin induction leads to cytoplasmic
dispersal of Dop1p, Vma4p and Scr7p.}

A: Severe mislocalization of Dop1p upon \emph{VPS8D} knockdown. Growing
cells expressing \emph{DOP1} -mNeon were fixed and the nuclei stained
with DAPI. DIC and fluorescent images of the wild type and
\emph{VPS8D}knockdown cells are shown in the left and right panels,
respectively.

B: Severe mislocalization of Vma4p upon \emph{VPS8D} knockdown. Growing
cells expressing \emph{VMA4} -mNeon were fixed and the nuclei stained
with DAPI. DIC and fluorescent images of the wild type and
\emph{VPS8D}knockdown cells are shown in the left and right panels,
respectively.

C: Severe mislocalization of Scr7p upon \emph{VPS8D} knockdown. Growing
cells expressing \emph{SCR7} -mNeon were fixed and the nuclei stained
with DAPI. Fluorescent images of wild type and \emph{VPS8D} knockdown
cells are shown in the left and right halves, respectively, with focal
planes selected from Z-stacks to illustrate the cell surface (left) and
cell interior (right). Images were taken with a Zeiss Axio Observer 7
system.

Figure 4:

\textbf{Live imaging of Dop1p reveals altered CVC morphology
upon\emph{VPS8D} knockdown.}

A: Live imaging of the CVC contractile cycle in a wildtype cell
expressing Dop1p-mNeon. The focal plane represents a cross-sectional
view of the CVC, with 0.5 sec frame intervals. Seven successive images
extracted from a video (Movie 3) are shown. Cells from growing cultures
were immobilized using CyGEL, as described in Materials and Methods.
Videos were captured using a Marianas spinning disc confocal microscope.

B: Live imaging of the CVC in \emph{VPS8D} knockdown cell expressing
Dop1p-mNeon. The cell selected contains two large bladders that undergo
asynchronous contractions. Imaging conditions were the same as in Fig.
4A. The seven successive time-lapse images were extracted from Movie 4.

C: Live imaging of the CVC in \emph{VPS8D} knockdown cell expressing
Dop1p-mNeon. The cell selected contains multiple Dop1p-labeled vesicles
dispersed in the cell posterior, concentrated in the region where the
single stereotypical CVC bladder is typically located. Two of them
contract over the course of the video. Imaging conditions were the same
as in Fig. 4A, with 1 sec frame intervals. The six successive time-lapse
images were extracted from Movie 5.

Figure 5:

\textbf{Vps8Dp concentrates at junctions between adjacent vacuoles}

A: Live imaging of numerous large Dop1p-labeled vacuoles in cells
expressing Dop1p-mNeon, following \emph{hypo} -osmotic shock. The cell
shown was imaged to capture a cross-sectional view, in a time-lapse
video with 4 sec frame intervals (Movie 6). Four successive images
extracted from the video are shown. Growing cell cultures were
hypo-osmotically challenged with 10\selectlanguage{english} mM Tris-HCl buffer, pH 7.4 and
immediately immobilized for imaging using CyGEL, as described in
Materials and Methods. Videos were captured using a Zeiss Axio Observer
7 system.

B: Live imaging of the numerous Dop1p-labeled vacuoles
following\emph{hyper} -osmotic shock in cells expressing Dop1p-mNeon.
The cell shown was imaged to capture a cross-sectional view, in a
time-lapse video with 0.15 sec frame intervals (Movie 7). Four
successive images extracted from the video are shown. Growing cell
cultures were hyper-osmotically challenged with 10\selectlanguage{english} mM sorbitol and
immediately immobilized for imaging using CyGEL, as described in
Materials and Methods. Videos were captured using a Marianas spinning
disc confocal microscope.

C: Live imaging of numerous large Vps8Dp-labeled vacuoles
following\emph{hypo} -osmotic shock in cells expressing Vps8Dp-mNeon.
The cell shown was imaged to capture a cross-sectional view, in a
time-lapse video with 0.5 sec frame intervals (Movie 8). Four successive
images extracted from the video are shown. Cells were treated as
described in Fig. 5A. Videos were captured using a Marianas spinning
disc confocal microscope.

D-E: Imaging of extracellular Vps8Dp-labeled vacuoles that leaked from
ruptured cells expressing Vps8Dp-mNeon.

D. Vps8Dp localizes densely at contact site between adjacent vacuoles,
which in some cases are sites of subsequent membrane fusion (arrowhead).
Four successive images extracted from Movie 9 are shown.

E: Vps8Dp re-distributes at the time that two vacuoles come into
contact. The two large vacuoles shown were separated by
\textasciitilde{}1\selectlanguage{greek}µ \selectlanguage{english}at the beginning of the video (t=0) (Movie 10). When
they moved into contact starting at t=10s, Vps8Dp accumulated at the
junction (boxed). A second example of this phenomenon is shown in Movie
11.

F. Plot of fluorescence intensity change with time for the boxed area
shown in Fig. 5E, in Movie 10. The data were plotted using GraphPad
Software Prism. The three red arrows correspond to the images shown in
panel E.

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