Biochem. J. (2009) 419, 1–13 (Printed in Great Britain)
1
doi:10.1042/BJ20081950
REVIEW ARTICLE
Phosphatidylinositol 3,5-bisphosphate and Fab1p/PIKfyve underPPIn
endo-lysosome function
Stephen K. DOVE1 , Kangzhen DONG, Takafumi KOBAYASHI, Fay K. WILLIAMS and Robert H. MICHELL
School of Biosciences, University of Birmingham, Birmingham B15 2TT, U.K.
PtdIns(3,5)P2 is one of the seven regulatory PPIn (polyphosphoinositides) that are ubiquitous in eukaryotes. It controls membrane
trafficking at multiple points in the endosomal/lysosomal system
and consequently regulates the size, shape and acidity of at least
one endo-lysosomal compartment. PtdIns(3,5)P2 appears to exert
this control via multiple effector proteins, with each effector
specific for a subset of the various PtdIns(3,5)P2 -dependent
processes. Some putative PtdIns(3,5)P2 effectors have been
identified, including Atg18p-related PROPPIN [β-propeller(s)
that bind PPIn] proteins and the epsin-like proteins Ent3p and
Ent5p, whereas others remain to be defined. One of the principal
functions of PtdIns(3,5)P2 is to regulate the fission/fragmentation
of endo-lysosomal sub-compartments. PtdIns(3,5)P2 is required
for vesicle formation during protein trafficking between endolysosomes and also for fragmentation of endo-lysosomes into
smaller compartments. In yeast, hyperosmotic stress accelerates
the latter process. In the present review we highlight and discuss
recent studies that reveal the role of the HOPS–CORVET complex
and the vacuolar H+ -ATPase in the process of endo-lysosome
fission, and speculate on connections between these machineries
and the Fab1p pathway. We also discuss new evidence linking
PtdIns(3,5)P2 and PtdIns5P to the regulation of exocytosis.
INTRODUCTION
As indicated in Figure 2, PtdIns3P has its own functions
and effectors. It is involved in at least three distinct membrane
trafficking processes: in post-Golgi sorting of lysosomal enzymes,
in autophagy and also in the retromer pathway [2] (and see
Figure 3). This latter process recycles membrane-bound receptors
for soluble cargos, for example the M6R (mannose-6-phosphate
receptor), and sorts them from endosomes to the TGN (transGolgi network) [10]. PtdIns3P is made at multiple sites in the
cell, including endosomes, pre-autophagic structures and the TGN
[2,11]. In contrast, PtdIns(3,5)P2 is probably generated on the
vacuole (endosome/lysosome in higher eukaryotes; see Appendix
1), although the lack of a suitable probe for this lipid has
prevented an exact definition of its whereabouts in cells [12].
Indeed PtdIns(3,5)P2 synthesis in other compartments is strongly
implied by some studies [13,14]. Like PtdIns3P, PtdIns(3,5)P2
also appears to have multiple functions in cells (see Figure 4 for
one hypothesis capable of explaining this).
PIPkIIIs (Type III PtdInsP kinases) are the enzymes that
catalyse 5-phosphorylation of PtdIns3P to PtdIns(3,5)P2 and are
found in almost all eukaryotic cells [1]. These large enzymes,
ranging from ∼ 1500 kDa to 2850 kDa, are all similarly organized,
comprising a PtdIns3P binding FYVE domain located near the
N-terminus, followed by a regulatory CCT-like chaperone domain
which is connected to the lipid kinase domain via a unique
sequence rich in cysteine and histidine residues [1]. The FYVE
domain is dispensable for Fab1p (where Fab is formation of aploid
and binucleate cells) activity since expression of a FYVE-deleted
mutant Fab1p in cells recovers all of the phenotypes associated
The days when all of the information on PtdIns(3,5)P2 could be
summarized in a concise work are past, so this review emphasizes
the latest developments. Several excellent recent reviews focus
on topics that are not highlighted in the present review
[1–5].
IDENTIFICATION AND BIOSYNTHESIS
The existence of PtdIns(3,5)P2 was suggested in 1989 [6], but it
was not formally identified until almost a decade later [7,8]. This
lipid is present in all eukaryotic cells that have been examined to
date and appears to control several membrane trafficking events,
as well as one arm of the cellular response to hyperosmotic stress
[1]. Like the other eukaryotic regulatory glycerolipids, known as
PPIn (PolyPhosphoInositides), PtdIns(3,5)P2 is synthesized from
PtdIns [7,8] (Figure 1).
PtdIns(3,5)P2 biosynthesis from PtdIns is achieved by two
sequential PPIn kinases, with each enzyme specific for a particular
hydroxy group of the inositol ring (Figure 2) [1]. The first step in
PtdIns(3,5)P2 biosynthesis is catalysed by the PtdIns 3-kinase
Vps34p (mVps34 in animals; where Vps is vacuolar protein
sorting), an endosome-resident enzyme that phosphorylates the
3-hydroxy group of the inositol ring of PtdIns, but not other PPIn
[9]. Vps34p-related Type III PI3Ks (phosphoinositide 3-kinases)
are found in almost all eukaryotes [1], and all members of this
subfamily appear to display the same substrate specificity [2].
Key words: endosome, Fab1p/PIKfyve, phosphatidyl 3,5bisphosphate, phosphoinositide, stress.
Abbreviations used: ALP, alkaline phosphatase; AP, adaptor protein; CMT, Charcot-Marie-Tooth; Cvt, cytoplasm-to-vacuole trafficking; Fab, formation
of aploid and binucleate cells; GEF, guanine-nucleotide-exchange factor; IRAP, insulin-regulated aminopeptidase; M6R, mannose-6-phosphate receptor;
MTM, myotubularin; MTMR, MTM-related; MVB, multi-vesicular body; PIPkIII, Type III PtdInsP kinase; PKB, protein kinase B; PLCδ, phospholipase Cδ;
PPIn, polyphosphoinositides; PROPPIN, β-propeller(s) that bind PPIn; PVE, pre-vacuolar endosome; RS-ALP, retention sequence-ALP; SGK1, serumand glucocorticoid-induced protein kinase 1; siRNA, small interfering RNA; SNARE, soluble N -ethylmaleimide-sensitive fusion protein-attachment protein
receptor; SNX, sorting nexin; SPR, surface plasmon resonance; TGN, trans -Golgi network; V-ATPase, vacuolar ATPase; Vps, vacuolar protein sorting.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2009 Biochemical Society
2
Figure 1
S. K. Dove and others
Structure of PtdIns and PtdIns(3,5)P 2
PtdIns is an acidic diacylglycerolipid that comprises between 5–20 % of total phospholipid in
eukaryotic membranes [112]. The fatty acids of PtdIns are highly unsaturated at the sn -2 position
of glycerol in many organisms and the same is consequently true of all PPIn [113]. The unique
headgroup of PtdIns, is myo -inositol: a cyclohexane-like molecule with six hydrogen atoms
replaced by hydroxy groups. When linked via the D-1 hydroxy group to a glycerolipid backbone,
all other hydroxy groups on inositol become stereochemically unique. PPIn are synthesized
via PPIn kinases, but no enzyme has yet been found that will phosphorylate the D-2 position.
D-6 phosphorylation appears to be reserved for the formation of GPI-anchored proteins [7].
Hence all seven known PPIn are derived from combinations of D-3 and/or D-4 and/or D-5
phosphorylation [113].
Figure 2 PPIn metabolism in eukaryotes and PtdIns(3,5)P 2 synthesis and
degradation in S.cerevisiae
PPIn are generated in invariant sequences from PtdIns by three disparate families of PPIn
kinases: the Type I PtdIns kinases, the Type II PtdIns kinases and the PtdInsP kinases [114].
These families are very distinct from each other, exhibiting little homology between members of
different families. Steps in the synthesis and degradation of ‘housekeeping’ PPIn are often (but not
always) associated with transitions between organelles because PPIn kinases and phosphatases
that are part of a particular synthetic or degradative sequence are sometimes resident in different
compartments [114]. The kinases that produce PtdIns(3,5)P 2 are Vps34p and Fab1p and appear
to reside on the endosome and vacuole respectively, although their localizations do overlap to
some degree [113]. Thus the observation that many PPIn are synthesized/degraded via an
invariant sequence of phosphorylations/dephosphorylations is probably a consequence both of
the specificity of PPIn kinases for a particular hydroxy group on the inositol ring, and because
of the cell biology underlying these processes [113].
c The Authors Journal compilation c 2009 Biochemical Society
Figure 3
Pathways of membrane trafficking in yeast
The organelles of the yeast S. cerevisiae are superficially similar to those in animal cells.
However, as explained in Appendix 1, this similarity is deceptive, e.g. the PGE (post-Golgi
endosome) probably does not equate with the mammalian early endosome and the PVE
probably does not equate with the mammalian late endosome. This Figure highlights the
established trafficking pathways (solid arrows), as well as more speculative connections that
are likely to exist but for which evidence is less substantial (broken arrows). The Fab1p
pathway mediates the retrograde vacuole-to-PVE pathway (green arrow), as well as the sorting
of some proteins into intraluminal vesicles at the MVB. The Cvt pathway begins with the
envelopment of octameric proaminopeptidase (yellow spheres) by membrane (of unknown
origin) and proceeds to deliver vesicles to the vacuole. During starvation a mechanistically
related process (autophagy) delivers cytoplasm and organelles to the vacuole for amino-acid
recycling. Atg18p is required for both autophagy and the Cvt pathway, whereas Atg21p
is needed only for the latter process. Neither process requires PtdIns(3,5)P 2 , but both
processes appear to require PtdIns3P . An animated version of this Figure can be found at
http://www.BiochemJ.org/bj/419/0001/bj4190001add.htm.
with loss of Fab1p [15]. However, the enzyme is no longer
tightly localized to the vacuole. The CCT domain is known to
be regulatory because of the isolation of several constitutively
hyperactive mutants of Fab1p that result in amino-acid changes
in this region [16,17]. In addition, at least one regulator of Fab1p
also binds in this area (see below).
PIPkIIIs are exemplified by Fab1p in Saccharomyces cerevisiae
and PIKfyve in mammals [1,18,19]. Both are active PtdIns3P
5-kinases in vitro [20] and in vivo [20,21], although PIKfyve
has also been reported to phosphorylate PtdIns to PtdIns5P [22].
We find this to be a very minor activity in yeast and it remains
an open question as to whether PtdIns5P represents a bona fide
biological product or a non-biological artefact of in vitro assays
[23]. PIKfyve also acts as a protein kinase: it is reported both
to be autophosphorylated and to phosphorylate at least one other
protein, the Rab9 effector p40 (see below) [24].
In yeasts, Fab1p-like PtdIns3P 5-kinases provide the sole
route of PtdIns(3,5)P2 synthesis: deletion of the FAB1 gene (in
S. cerevisiae) or the STE12 gene (Schizosaccharomyces pombe)
results in a complete absence of PtdIns(3,5)P2 [20,21,25].
However, it is possible that PIKfyve might not provide the
only route to PtdIns(3,5)P2 in mammals. In a recent study of
NIH 3T3 cells, a PIKfyve inhibitor (YM201636) diminished the
32
P-radiolabelled peak representing PtdIns(3,5)P2 by only ∼ 85 %
[26]. This raises the possibility of a second route to PtdIns(3,5)P2
in animals, possibly via PtdIns3P phosphorylation by Type I
PtdIns4P 5-kinases [27]. This route does not appear to function
in S. cerevisiae [23], despite the presence therein of a Type I
PtdIns4P 5-kinase (Mss4p). It remains to be determined whether
this ‘YM201636-insensitive pool of PtdIns(3,5)P2 ’ is genuine
or represents some other co-chromatographing 32 P-radiolabelled
compound. Whatever the answer, it is certain that PIKfyve makes
most of the PtdIns(3,5)P2 in mammals. Moreover, PIKfyve can
PtdIns(3,5)P 2 and Fab1p/PIKfyve
Figure 4
3
PPIn as foci for formation of membrane-bound complexes
PPIn may direct formation of membrane-bound complexes in a similar way to how Rab-GTPases function [113,114]. In the absence of PPIn, membrane proteins (green and yellow) and soluble
proteins (L1, L2 and purple) are unable to interact because protein–protein affinities are too low to sustain the formation of a complex. When a cargo ligand (L), binds to cargo receptors (yellow), then
PPIn are generated (dark blue spheres) via PPIn kinases (omitted for clarity). The PPIn effector proteins (L1 and L2) can then interact with membranes/membrane proteins and bring them together
to form a complex. Bridging proteins, that bind the effectors and membrane proteins, are then also recruited (purple). Once the complex has served its purpose, PPIn can be hydrolysed and the
complex disassembled because it is no longer stable without PPIn [114,115]. This model requires that all interactions between components be of low affinity so that several components are required
for any one protein to enter the complex. The above may impact on compartment identity, i.e. how the cell knows that the Golgi is the Golgi [115]. Membrane proteins are not likely to be compartment
markers because almost all are inserted into the endoplasmic reticulum and must traffic to reach their correct site of residence, and hence are mislocalized at least some of the time [15,32]. It is more
likely that protein complexes, such as the one above, mark compartments so that mislocalization of any one component does not result in the inappropriate assembly of the complex [32]. This may
also explain how a single species of PPIn, e.g. PtdIns(3,5)P 2 , is able to regulate several disparate functions in the same cells at the same time; because effectors are coincidence detectors and require
both membrane and protein partners to carry out their functions, any individual effector is only recruited to a protein complex if both its preferred PPIn and its preferred protein ligand are present in
the same membrane or microdomain [15]. So PPIn can regulate many different protein complexes by regulating several effectors that all bind the same lipid but have different protein partners [15].
functionally replace S. cerevisiae Fab1p in fab1 yeast; indeed
it can even recover hyperosmotic-stress-stimulated PtdIns(3,5)P2
synthesis, contrary to what we originally reported [23].
PtdIns(3,5)P2 is of low abundance in both yeast and animal
cells. In unstressed mammalian cells, it typically constitutes
0.04–0.08 % of total inositol phospholipid [7,8]: steady-state
levels of PtdIns(3,5)P2 are approx. one-tenth to one-third of the
concentrations of PtdIns3P and it is many-fold less abundant than
either PtdIns4P or PtdIns(4,5)P2 . In response to hyperosmotic
stress, the concentration of PtdIns(3,5)P2 in yeast becomes
elevated transiently: up to 20-fold in both S. cerevisiae and
S. pombe [7]. More modest increases have been observed in plant
cells and also in at least one differentiated animal cell line, namely
3T3 L1 adipocytes [28,29].
Figure 5
NORMAL PtdIns(3,5)P 2 SYNTHESIS ALSO REQUIRES Vac7p,
Vac14p AND Fig4p
Three other proteins, Vac7p, Vac14p and Fig4p, are required
if Fab1p is to sustain both basal and hyperosmotically
stimulated PtdIns(3,5)P2 synthesis in S. cerevisiae [16,17,30,31].
Inactivation of any of these three proteins causes deficient
PtdIns(3,5)P2 synthesis, with the cells displaying most or all of
the defects characteristic of fab1 cells (see below). In addition,
one of the effectors of PtdIns(3,5)P2 , Atg18p, also modulates
Fab1p activity, but this time in a negative fashion (see below and
Figure 5).
BLAST searches indicate that Vac7p is present in ascomycete
fungi and maybe their close relatives, but not in other eukaryotes
[30]. Vac7p is an integral vacuolar membrane protein that is
predicted to possess a single transmembrane domain; detergent
is needed to extract it from vacuole membranes [30]. Its Cterminal segment, which extends into the interior of the yeast
vacuole, is highly conserved. In contrast, much of the rest of the
protein, including its large N-terminal cytosolic domain, shows
low sequence complexity and considerable sequence variation,
even among close relatives of S. cerevisiae, suggesting that it
is not evolutionarily constrained. Vac7p is also required for
Atg18p, a PtdIns(3,5)P2 effector, to associate with the vacuole
membrane (see below) [15]. Atg18p homologues are found in
almost all eukaryotes [1,32], so it might be expected that nonfungal eukaryotes will possess some as yet unrecognized protein
that functionally substitutes for Vac7p.
Atg18p mediates feedback inhibition of Fab1p
The PtdIns(3,5)P 2 effector Atg18p seems to feedback regulate the kinase, Fab1p, that produces
this lipid. The following is a model based on data from [116]. (1) Atg18p is not present on the
vacuole and so Fab1p is active, and produces PtdIns(3,5)P 2 (blue spheres) [116]. (2) Atg18p
is recruited to the vacuole membrane via binding to PtdIns(3,5)P 2 and inhibits Fab1p, possibly
via competition with Vac7p [116]. (3) PtdIns(3,5)P 2 is hydrolysed (by Fab1p-associated Fig4p)
and Atg18p leaves the membrane. Fab1p becomes active once released from Atg18p-mediated
inhibition and the cycle begins again [117]. A similar mechanism could account for control
of synthesis of other PPIn and explain how their mass levels are so tightly regulated, together
with the fact that PPIn kinases often seem to be associated with lipid phosphatases, e.g. the
MTM–Vps34p pairing or the Fab1p–Fig4p complex.
Vac14p and its murine homologue mVac14 (also known as
ArPIKfyve) are large peripheral membrane proteins that appear to
be composed largely of HEAT-like domains [17,31,33]. Vac14p
and mVac14 deletion cause, respectively, an ∼ 90 % decrease
in PtdIns(3,5)P2 synthesis in S. cerevisiae [17,30,31] and at
least a 50 % reduction in knockout mice [34]. Some Vac14p is
mislocalized in fab1 yeast, suggesting that Fab1p is needed
for efficient association of Vac14p with the vacuole membrane
[35]. Indeed recent work has shown that this interaction between
Fab1p and Vac14p is mediated by a sequence, T1017 ILLR1021 ,
that is present in the CCT-like domain of Fab1p [36]. PIKfyve
and mVac14, can also be co-immunoprecipitated from animal
cells [37], and this association is probably mediated by the
same interaction sites. Vac14p also physically interacts with
the PtdIns(3,5)P2 -specific 5-phosphatase, Fig4p: Vac14p and
Fig4p co-immunoprecipitate, suggesting that they co-exist in
a membrane-associated complex, and they have also been
c The Authors Journal compilation c 2009 Biochemical Society
4
S. K. Dove and others
functionally linked by yeast two-hybrid analyses [17,31,35].
Vac14p is partially mislocalized in a fig4 mutant, much as
is it is a fab1 mutant, suggesting that Fab1p and Fig4p are
both required to keep Vac14p on the vacuole membrane [35].
Indeed recent data has shown that Fab1p, Vac14p and Fig4p are
present in a complex that probably contains at least two copies of
Vac14p [36]. It seems paradoxical that Vac14p should associate
with both a PtdIns3P 5-kinase and a PtdIns(3,5)P2 -directed
5-phosphatase, so it is possible that there is some spatial or
temporal discrimination between these Vac14p functions (but see
below) [35].
The mVac14 protein also keeps other interesting company, for
unexplained reasons: it binds to the PDZ domain of nNOS [neural
isoform of NOS (nitric oxide synthase)] in vertebrates [38], as well
as associating with the Tax protein produced by the retrovirus
HTLV-1 (human T-cell lymphotrophic virus-1) [39].
Fig4p is a Sac1p-like PPln phosphatase that removes the
5-phosphate of PtdIns(3,5)P2 to produce PtdIns3P (see Figure 2)
[16]. In the one published study of recombinant Fig4p activity,
PtdIns(3,5)P2 was the only PPln attacked by Fig4p [35]. This
observation is surprising given the similarity that the catalytic
site of Fig4p bears to the relatively non-specific Sac1p PPIn
phosphatase [40]. Fig4p displays a classical CX5 R catalytic motif
and an extensive C-terminal coiled-coil domain that mediates
interaction with Vac14p: deletion of Vac14p or removal of
this C-terminal segment renders Fig4p completely cytosolic, as
does deletion of FAB1 [35]. Surprisingly, inactivation of this
PtdIns(3,5)P2 5-phosphatase, in yeast or in animal cells, causes
a profound defect in PtdIns(3,5)P2 synthesis [17,41–43]. Recent
work has unravelled this apparent conundrum by demonstrating
that Fig4p is required as a scaffold protein which is necessary
for Fab1p and Vac14p to interact [36]. Hence Fab1p-like lipid
kinases appear, like Vps34p, to associate with their cognate
lipid phosphatases.
MYOTUBULARINS DEGRADE PtdIns(3,5)P 2
The MTMs (myotubularins) are a family of evolutionarily related
‘dual-specificity protein phosphatase-like’ PPIn phosphatases,
comprising MTM1 and the MTMR (MTM-related) proteins
(MTMR1–MTMR13 and MTMR14/JUMPY) [44–46]. Their
catalytic sites include CX5 R motifs akin to those in Fig4p and
the PtdIns(3,4,5)P3 3-phosphatase PTEN (phosphatase and tensin
homologue deleted on chromosome 10) [45]. They were initially
considered to be PtdIns3P phosphatases, but were then shown
to 3-dephosphorylate PtdIns(3,5)P2 to the novel PPln PtdIns5P,
a molecule which may itself have signalling functions [47–50].
One recent hypothesis suggests that active MTMs naturally target
PtdIns3P, but that the inactive versions can heterodimerize with
the active phosphatases and direct them to PtdIns(3,5)P2 [51].
Mutations in members of the MTM family are causal in the
genetically transmitted progressive human diseases X-linked myotubular myopathy (MTM1 or Jumpy respectively, in X-linked
and autosomal forms of the disease) and CMT (Charcot-MarieTooth) disease (MTMR2 or MTMR13, in two CMT subtypes)
[46]. X-linked myotubular myopathy is characterized by aberrant
muscle cells with central nuclei like those present in foetal
myotubes. CMT disease involves faulty nerve myelination, resulting in poor conduction and neurodegeneration [46]. Moreover,
fig4−/− mice display a condition similar to human CMT disease
[43], and FIG4 is inactivated in some humans with a form of this
disease known as CMT4J [42,52].
The molecular mechanism causing these defects is still not
clear, but appears to result from altered myelination [46].
In fibroblasts cultured from mouse models of these diseases,
c The Authors Journal compilation c 2009 Biochemical Society
inactivation of MTMs leads to an increase in PtdIns3P and
PtdIns(3,5)P2 , but to a reduction in PtdIns5P [46], whereas PtdIns(3,5)P2 levels are much reduced in fig4−/− mice [43]. Exactly
how the dysregulation of PtdIns(3,5)P2 and/or PtdIns3P and/or
PtdIns5P metabolism and of functions controlled by these PPIn
is causal for X-linked myotubular myopathy and the CMTs
remains to be determined from detailed functional studies of
the affected tissues [46]. One recent hypothesis proposes that
PtdIns5P controls exocytosis, and hence plasma membrane area
[51]. Since the neurological problems in the CMT-related diseases
seem to result from aberrant out-foldings of the plasma membrane,
this hypothesis clearly has merit. It might also explain the effect of
PIKfyve on the trafficking to the cell surface of certain permeases
and ion channels (see below).
PUTATIVE EFFECTORS OF PtdIns(3,5)P 2 INCLUDE PROPPINS
[β-PROPELLER(S) THAT BIND PPIn] AND EPSINS
The search for effector proteins that bind to PtdIns(3,5)P2 and
mediate its effects on various cellular processes has identified
several candidates [1].
Perhaps the most convincing are the seven bladed β-propeller
PROPPIN proteins exemplified by Atg18p (also known as Svp1p,
Aut10p, Nmr1p or Cvt18p) [32,53,54]. These proteins were first
associated with PtdIns(3,5)P2 because deletion of ATG18 results
in an enlargement of the yeast vacuole that largely phenocopies
the defect seen in fab1 cells [32]. PROPPINs all include the
PPIn-binding sequence motif Glu/Gln-ψ-Arg-Arg-Gly (where ψ
represents a hydrophobic residue), and are conserved throughout
all eukaryotes, with the exception of some pathogenic microorganisms [1]. Most organisms have multiple PROPPINs: three
in S. cerevisiae (Atg18p, Atg21p, also known as Hsv1p, and
Hsv2p), and four (WIPI-1 to WIPI-4) in mammals [1,32,55–57].
Atg18p has a role in autophagy that appears to be separate from
its role in the Fab1p/PtdIns(3,5)P2 pathway; autophagy does not
appear to require PtdIns(3,5)P2 , at least in yeast (see Appendix
2) [54,58] (see also Box 1 of the Supplementary Information to
[1]). Atg21p is also somehow involved in the autophagy-related
Cvt (cytoplasm-to-vacuole trafficking) pathway that delivers the
protease aminopeptidase I from the cytosol to the vacuole lumen
[59–62] (see Figure 3), and Hsv2p was previously identified as a
factor required for micronucleophagy [32,63].
The PPIn-binding properties of PROPPINs suggest that
they are genuine PtdIns(3,5)P2 effectors [32]. They bind
PtdIns(3,5)P2 with high affinity (K d ∼ 500 nM) and selectivity,
and most of the PROPPINs tested display no appreciable binding
to other PPln when assayed using quantitative approaches
such as PPIn competition assays and SPR (surface plasmon
resonance; Biacore) analysis of PROPPIN binding to PPIn-doped
phospholipid monolayers [32,64,65]. The affinities of Atg18p
and Hsv2p for PtdIns(3,5)P2 are similar to the affinity of the PH
(pleckstrin homology) domain of PLCδ (phospholipase Cδ) for
PtdIns(4,5)P2 , which allows PLCδ to interact robustly with the
plasma membrane [32]. However, PtdIns(3,5)P2 is many-fold
less abundant in unstressed cells than PtdIns(4,5)P2 , so it seems
likely that additional protein–protein interactions might be
needed to reinforce the recruitment of PROPPINs to membranes,
in accordance with the principles alluded to in Figure 4. Vac7p or
a Vac7p-associated protein seems to fulfil this function in yeast
[36].
By contrast, studies using ‘lipid blot’ assays reported an apparent preference of yeast Atg21p and human WIPI-1 for binding
PtdIns3P [61,62]. The physiological meaning of this is not clear,
since quantitative methods such as SPR and lipid-competition
assays have suggested that the PtdIns(3,5)P2 affinities of almost
PtdIns(3,5)P 2 and Fab1p/PIKfyve
all PROPPINs [except for Dm3, a Drosophila melanogaster
PROPPIN with equal affinities for PtdIns3P and PtdIns(3,5)P2 ]
far outweighs their modest affinity for PtdIns3P [65].
Atg21p and Hsv2p appear to be mislocalized from membranes
in vps34, but not fab1 cells, suggesting that PtdIns3P may
indeed contribute to their localization in vivo [61,62]. In fact
a small pool of Atg18p, Atg21p and Hsv2p all localize to
endosomes as well as the vacuole, and this endosomal localization
is dependent upon PtdIns3P generated by the portion of Vps34p
associated with Vps38p: the so-called ‘complex II’ form of
this enzyme [11]. Since the affinities of Atg18p, Atg21p and
Hsv2p are much too low for PtdIns3P to be the key determinant
of their association with membranes, why should a lack of
PtdIns3P mislocalize these proteins? It now seems likely that
some modification of the lipid-binding specificity of PROPPINs
can occur in vivo, although no mechanism is currently defined.
Alternatively, they may associate with protein partners that
themselves depend upon PtdIns3P for their localization.
The functions of Atg18p-like proteins were illuminated by a
study that examined a version of Atg18p that was membranelocalized by virtue of fusion to a transmembrane domain derived
from the vacuolar ALP (alkaline phosphatase), Pho8p [32]. This
Atg18–Pho8p fusion rescued the atg18 strain in terms of the defects associated with the Fab1p pathway, but failed to rescue the
autophagy defect of atg18 cells [66,67]. This confirms what we
originally suggested; namely that the PtdIns(3,5)P2 binding and
autophagic functions of Atg18p are distinct and separable [68]. It
seems logical to suppose that the same could be true of Atg21p and
Hsv2p (and indeed all other PROPPINs); that they participate in
PtdIns(3,5)P2 -dependent and -independent processes. This raises
the question of why PROPPINs should continue to support two
such apparently unrelated processes throughout evolution? One
hypothesis we favour is that PROPPINs have evolved to carryout membrane and protein recycling and that they participate in
this process in a number of different pathways. In the case of
Atg18p, one of these recycling steps is PtdIns(3,5)P2 -dependent
but the other, supporting autophagy, is not and may instead rely
upon PtdIns3P. Hence PPIn may act just to localize the recycling
machinery to particular sites in the cell, with different PPIn
specific for each compartment.
The two epsin-like yeast proteins Ent3p and Ent5p represent a
second class of PtdIns(3,5)P2 effectors [68]. Epsins, first identified
as proteins required for endocytosis, include PPIn-binding
ENTH or ANTH domains [66,67]. The first described epsins
bound PtdIns(4,5)P2 , but epsinR (the mammalian homologue
of Ent3p/Ent5p) appears to bind PtdIns4P [69,70]. By contrast,
Ent3p and Ent5p have both been reported to bind PtdIns(3,5)P2
selectively in vitro and to require Fab1p for vacuolar localization
in vivo [13], but this is put in some doubt by the fact that
others have failed to detect this PtdIns(3,5)P2 binding either
in vitro or in vivo [65]. As discussed below, however, both Ent3p
and Ent5p appear to be implicated in AP1 (adaptor protein 1)dependent trafficking of the chitin synthase Chs3p [13], a
process now known to be dependent on Fab1p and PtdIns(3,5)P2
[14,18,20,21,25,43,71–74]. It is therefore possible that, despite
displaying a lack of selectivity for PtdIns(3,5)P2 as a sole
ligand, Ent3p/Ent5p may be involved in some of the functions
of PtdIns(3,5)P2 [21].
LOSS OF PtdIns(3,5)P 2 SYNTHESIS LEADS TO DEFECTS
IN DISPARATE PROCESSES
The consequences of ablating PtdIns(3,5)P2 synthesis have now
been extensively studied [genetically, by siRNA (small interfering
5
RNA), or by expression of ‘dominant-negative’ Fab1p or PIKfyve
variants] in several model organisms, including S. cerevisiae,
S. pombe, Caenorhabditis elegans, D. melanogaster and various
mammalian cells in culture [1]. Notably, yeast cells in which
the wild-type FAB1 gene has been replaced with a kinase-dead
version of Fab1p (fab1D2134R ) replicate most of the defects seen in
S. cerevisiae fab1 mutants [18,20,21,75], so it seems that the
key event in these mutants is a failure of PtdIns(3,5)P2 synthesis
rather than absence of Fab1p.
These studies implicate PtdIns(3,5)P2 in several cellular
processes, centred around the homoeostasis of acidic organelles in
the endosomal/lysosomal system [17,30]. These, to be discussed
individually, include: (i) control of the size and shape of
endosomes and/or lysosomes [32,76]; (ii) acidification and
inheritance of these organelles [14,71]; (iii) retrieval of membrane
and proteins from the vacuole/lysosome to the endosome
[7,29,77–79]; (iv) ubiquitin-dependent sorting of some cargo
proteins into MVBs (multi-vesicular bodies) [1]; and (v) an illdefined role in stress signalling, most likely involving a still
undefined stress signalling pathway [80–82], stress-provoked
changes in one or more of the above processes, or some
combination of these [83,84].
In addition to these obviously endosomal/lysosomal defects,
loss of PtdIns(3,5)P2 is reported to restrict exocytosis and the
trafficking of some membrane transporters to the cell surface
in animal cells [34,85,86] and of the chitin synthase Chs3p in
S. cerevisiae [1], and there is evidence that M6R recycling
may be perturbed in cells in which the PtdIns(3,5)P2 supply
is compromised [18,42]. The ability of a single lipid signal
to regulate all of these processes is probably possible due to
mechanisms like those outlined in Figure 4.
CONTROL OF ENDOSOME/LYSOSOME SIZE AND TRAFFICKING
Deletion of FAB1, VAC14 or FIG4, or their fungal or metazoan
homologues, produces cells in which the cytoplasm becomes
extensively vacuolated. One or more membrane-delineated
compartments, often of ill-defined origin or identity, become
greatly enlarged [18]. In some cases, enlargement of the aberrant
compartment(s) can perturb the normal inheritance of other organelles [71,72]. In the S. cerevisiae fab1 mutant the enlarged
organelle is so large that it can preclude partitioning of the
two nuclei between mother and daughter cells during mitosis
[75,87,88].
The engorged yeast compartment is undoubtedly the vacuole,
since vacuole markers decorate its surface and it remains distinct
from the adjacent and much smaller endosomal structures,
markers for which (e.g. Snf7p) are not mislocalized to the vacuole
membrane (S. K. Dove, unpublished work). However, its identity
in animal cells remains a matter of debate, partly because many
of the analyses involve examination of terminal phenotypes that
are the results of long-standing gene deletion, so it is difficult
to identify the processes that gave rise to the enlarged vacuolar
structures. Most studies have concluded that late endosomes
or lysosomes, or both, contribute to the vacuolated structures
[89], but some have failed to detect markers of any defined
compartment(s) on these aberrant organelles [26]. By contrast,
a recent study that followed the emergence of the vacuolated
condition during siRNA suppression of PIKfyve expression in
HeLa cells suggests that early endosomes are the first organelles
to be affected [26]. Work has just started using YM201636, a
novel inhibitor of PIKfyve, with which cells could be switched
between the normal and vacuolated states [26]. The enlarged
vacuolar structures in inhibitor-treated cells carried markers for
both late endosomes and lysosomes [76]. Such examinations of
c The Authors Journal compilation c 2009 Biochemical Society
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S. K. Dove and others
the progressive inhibition of PtdIns(3,5)P2 synthesis at early times
should permit better definition of the primary PtdIns(3,5)P2 dependent events. One caveat is that the YM201636-treated
cells retain a little ‘PtdIns(3,5)P2 ’, as discussed earlier, so some
PtdIns(3,5)P2 -dependent events might persist in the presence of
the inhibitor.
MOLECULAR BASIS OF PtdIns(3,5)P 2 -DEPENDENT MEMBRANE
TRAFFICKING: A VACUOLE-TO-ENDOSOME RECYCLING PATHWAY
It seems likely that at least two distinct molecular defects
contribute to the ‘swollen vacuole’ phenotype of fab1 yeast and
PIKfyve-inhibited animal cells. The first defect was uncovered
by following trafficking of RS-ALP (retention sequence-ALP), a
chimaeric membrane protein that traffics to the vacuole from the
Golgi directly by the AP-3 pathway, by-passing all endosomes
[76]. RS-ALP then recycles out of the vacuole compartment
[90] by a retrograde trafficking (retrieval) pathway that takes it
through the PVE (pre-vacuolar endosome; see the green arrow on
Figure 3), and mutants that prevent exit from the PVE can trap RSALP there [76]. The SNARE (soluble N-ethylmaleimide-sensitive
fusion protein-attachment protein receptor) Pep12p is required for
fusion of vacuole-derived vesicles with the PVE, demonstrating
that this pathway proceeds via transport intermediates. Vti1p,
another SNARE, undergoes recycling by the same route [76].
Recycling of RS-ALP from the vacuole requires the Fab1p
activator Vac7p [76], and fab1 cells are also defective for RSALP recycling [32]. This implies that the retrieval of membrane
proteins from the vacuole is PtdIns(3,5)P2 -dependent and that, in
PtdIns(3,5)P2 -deficient cells, a failure of membrane recycling out
of the vacuole to the PVE might provide a mechanism by which the
vacuole could accumulate membrane and become enlarged [32].
The PROPPIN Atg18p/Svp1p is a PtdIns(3,5)P2 effector that
mediates the effects of PtdIns(3,5)P2 on this retrieval pathway
[32]; RS-ALP recycling out of the vacuole to the PVE is blocked
in atg18 cells. However, the PtdIns(3,5)P2 concentration in
unstressed atg18 cells is more than 10-fold higher than in wildtype cells, indicating that feedback inhibition by PtdIns(3,5)P2 ligated Atg18p would normally limit PtdIns(3,5)P2 synthesis at
the vacuole (as schematically depicted in Figure 5) [15,32]. Other
PtdIns(3,5)P2 -dependent events remain normal in atg18 cells,
confirming that PtdIns(3,5)P2 must mediate its diverse effects
through multiple effectors [15,32].
This inhibitory control of Fab1p by Atg18p that had been
localized to the vacuole by PtdIns(3,5)P2 was unexpected,
but clearly it helps to maintain normal PtdIns(3,5)P2 levels
[15]. Whether other PPIn effectors might also influence local
concentrations of their PPIn ligands is not known, but it does
seem an elegant possible means of control of steady-state PPIn
levels in cells.
We originally suggested that PtdIns(3,5)P2 might allosterically
regulate the effect of Atg18p on PtdIns(3,5)P2 synthesis, but it
now seems likely that the key role of PtdIns(3,5)P2 is simply
to bring Atg18p to the vacuole membrane where Fab1p resides
[15]. This conclusion emerges from the properties of an Atg18p–
Pho8p construct that lacks the FRRG motif and does not bind
PtdIns(3,5)P2 , but is vacuole-localized by the transmembrane
domain of Pho8p [15]. This Atg18–Pho8p chimaera is at the
vacuole even in the absence of PtdIns(3,5)P2 . PtdIns(3,5)P2
levels are normal in atg18 cells expressing this chimaera [32],
suggesting that Atg18p and Fab1p only need to be on the same
membrane for Atg18p to restrict the activity of the lipid kinase.
This Atg18–Pho8p chimaera also corrects vacuole enlargement
in atg18 cells and in vac14 cells [despite their PtdIns(3,5)P2
deficit] [32], but not in fab1 or vac7 cells. This suggests that
c The Authors Journal compilation c 2009 Biochemical Society
correcting one membrane trafficking defect can overcome a lack
of Atg18p or of Vac14p, but that a second, and still undefined,
functional defect contributes to vacuole enlargement in fab1 or
vac7 cells. This idea is supported by the fact that the vacuole
defects in fab1 cells are more severe than atg18 cells [91].
For example, raising the temperature to 37 ◦ C exaggerates the
fab1 vacuole defect, but not that in atg18 cells (S. K. Dove,
unpublished work).
Moreover, fab1 cells often display an ‘open figure eight
morphology’, in which the vacuole straddles the junction between
the mother and daughter cell, but in some strain backgrounds
atg18 cells do not (S. K. Dove, unpublished work). We
originally attributed this difference to the fact that the Cvt pathway
(see Figure 3), one of three main routes that deliver proteases to
the lumen of the S. cerevisiae vacuole, is blocked in atg18,
but not fab1, cells. We hypothesized that the slowing of the
Cvt pathway would mean that atg18 vacuoles would be less
enlarged than fab1 vacuoles. However, we constructed fab1
cells in which the Cvt pathway was simultaneously inactivated
and they still had larger vacuoles than atg18 cells and more
frequently displayed the ‘open figure eight’ morphology (S.K.
Dove, unpublished work).
These results all suggest that a second PtdIns(3,5)P2 -dependent
process, requiring Fab1p and Vac7p but not Atg18p, achieves
some of the membrane exit from the vacuole, and that this process
contributes to the vacuole enlargement in fab1 and vac7 cells.
Obvious participants might have been the two other Atg18prelated yeast PROPPINs, Hsv2p and Atg21p in this process.
However, single and double mutants of the genes encoding these
showed no vacuolar defects, even though they bind PtdIns(3,5)P2
avidly [17]. Indeed, they have no known PtdIns(3,5)P2 -dependent
functions as yet, although deletion of either partially reduces
vacuole fragmentation in response to hyperosmotic stress (S. K.
Dove, unpublished work).
CONTROL OF VACUOLE MORPHOLOGY BY THE RELATED HOPS
AND CORVET COMPLEXES
In exponentially growing cells, the S. cerevisiae vacuole normally
exists as 3–6 distinct, but tethered, sub-compartments [91].
This morphology changes during growth so that at high-glucose
concentrations, the vacuole is more subdivided, but at higher cell
densities it becomes progressively larger and less fragmented (see
Figure 6). Vacuole morphology can also be perturbed both by
hyperosmotic stress, which further sub-compartmentalizes the
vacuole into many smaller organelles [92,93], and by hypoosmotic stress, which triggers the sub-compartments to fuse
into one large vacuole (see Figure 6) [93]. The machinery
for homotypic vacuole fusion is relatively well characterized
[94], whereas that for fission was undefined until very recently
[96]. However, a new recognition that the two processes use
common components begins to provide an understanding of the
mechanisms underlying vacuole fusion/fission.
Proteins encoded by the class C VPS genes, which are
conserved in most eukaryotes, make up the HOPS tethering
complex, which controls homotypic vacuole–vacuole fusion
[92]. In the original VPS mutant screens, class C genes were
those whose deletion or mutation led to failure to produce a
distinct vacuole: the vesicles from the four or more pathways
feeding into the vacuole failed to fuse resulting in the formation
of a number of tiny aberrant organelles [92]. Several class C Vps
proteins (Vps11p, Vps16p, Vps18p and Vps33p) form the core of
the HOPS complex [95]. Fusion of vacuolar elements mediated
by the HOPS complex is regulated by the Rab7-like Ypt7p.
Vam6p/Vps39p, which binds preferentially to GTP-ligated
PtdIns(3,5)P 2 and Fab1p/PIKfyve
Figure 6 CORVET–HOPS, the V-ATPase and the Fab1p pathway co-operate
to regulate endo-lysosomal fusion/fission status
The yeast vacuole normally exists as a series of discrete, but tethered, compartments whose
size and shape depend upon growth status, osmolarity and activity of the V-ATPase, the
HOPS–CORVET equilibrium and the activity of the Fab1p pathway. This diagram illustrates
the interplay between these pathways and emphasizes that the interconversion of HOPS into
CORVET has profound implications for the fusion–fission equilibrium. In the centre, the vacuole
is depicted as it appears in exponentially growing cells. Starvation and/or hypo-osmotic stress
can cause vacuolar fusion mediated by HOPS, shifting the vacuole to the single enlarged
organelle shown on the left. Alternatively, hyperosmotic stress or an artificial hyperactivation of
the CORVET or Fab1p pathways can result in extreme fragmentation of the vacuole, as shown
on the right-hand side of the Figure. The casein kinase Yck3p, apparently restrains fusion after
fission has occurred by phosphorylating a number of targets; Vps41p, a subunit of HOPS is one
of these targets.
Ypt7p, appears to be its effector, and the activity of Ypt7p is
controlled by the GEF (guanine-nucleotide-exchange factor)
Vam6p [95].
It now emerges that HOPS can interconvert between a tethering
complex for fusion between vacuole sub-compartments and a
slightly different CORVET complex that catalyses either vacuole
fission into sub-compartments or vacuole–endosome fusion, or
both [95]. It does so by exchanging Ypt7p and its partners for
Vps21p (yeast Rab5) plus its effector (Vps8p) and GEF (Vps3p)
[90]. The physiological process that influences this shift from
fusion to fission is unknown, but overexpression of Vps3p can
drive a striking fission of the vacuole [63] that is not unlike the
fragmentation that occurs during hyperosmotic shock.
It will be important for future studies to examine whether the
CORVET complex might link PtdIns(3,5)P2 /Fab1p and vacuole
sub-compartmentation. The CORVET regulator Vps21p controls
entry of proteins into the Pep12p-positive PVE that is adjacent
to the vacuole (see Figure 3) [96], so it is clearly possible
that PtdIns(3,5)P2 influences vacuolar fission and maybe also
retrograde vesicle trafficking from the vacuole to the PVE
trafficking in some fashion, promoting HOPS–CORVET subunit
exchange. A link between the Fab1p and CORVET machineries is
certainly suggested by the finding that deletion of some CORVET
and/or HOPS subunits can cause mislocalization of Atg18p,
Atg21p and Hsv2p (Figure 7) [97].
PtdIns(3,5)P 2 -DEPENDENT VACUOLE ACIDIFICATION IS REQUIRED
FOR VACUOLE FISSION
The V (vacuolar)-ATPase was recently revealed as another
component of the vacuolar fission machinery [96]. V-ATPase is a
large H+ -pumping protein complex that drives the acidification
of endosomes and lysosomes/vacuoles [96]. Related to the
H+ -gradient-driven ATP synthase of bacteria and bioenergetic
Figure 7
7
PtdIns(3,5)P 2 -dependent events in control of vacuole size
The major defect associated with a loss of PtdIns(3,5)P 2 synthesis is a hypertrophy and fusion
of membrane compartments of endosomal/lysosomal origin. Two separate Fab1p-dependent
events are likely to be the cause of this. The first is shown in (a), where vesicles bud off of the
yeast vacuole to transport membrane proteins, such as Vti1p, to the pre-vacuolar endosome.
This process relies upon PtdIns(3,5)P 2 because it is required to localize Atg18p to the vacuole
surface. Hence Fab1p, Vac7p, Vac14p and Fig4p are also required because they are needed to
maintain basal PtdIns(3,5)P 2 synthesis at normal levels. The second PtdIns(3,5)P 2 -dependent
process is vacuole–vacuole fission illustrated in (b). A bud appears on a vacuole and then grows
as membrane flows into it, although it is not clear if this occurs via vesicular intermediates
or directly via the bud–vacuole junction. The bud then undergoes scission. This process
appears to require PtdIns(3,5)P 2 , Atg18p (partially), the CORVET complex and a functioning
H+ V-ATPase as well as the associated H+ gradient. Are (a) and (b) actually the same process,
just with scission of the growing ‘vesicle’ happening much later in (b) than (a), or are they
mechanistically distinct?
organelles, it is composed of a membrane-embedded V0
subcomplex and an associated peripheral V1 subcomplex [96].
The V-ATPase has been implicated in vacuole–vacuole fusion
for some time, although in a manner that does not require a transmembrane H+ gradient. Previous work has shown that vacuole fission in yeast cells requires both the V-ATPase and vacuole acidification [20,21,30,31]. Inhibition of V-ATPase with bafilomycin
prevents vacuole fission, as does inactivation of any V-ATPase
subunit [21,30]. These treatments also prevent the rapid fragmentation of the vacuole in response to hyperosmotic stress.
This may also explain why, at high cell densities or in the
presence of low glucose, the vacuole tends to fuse to form one
large compartment; a low concentration of glucose causes the
V1 sector to dissociate from the V0 component and this prevents
fission while maintaining fusion, because only the former process
requires the H+ gradient (see Figure 6).
It has long been known that PtdIns(3,5)P2 is required for
vacuolar acidification because fab1, vac7 and vac14 cells all
have neutral vacuoles, a defect that appears to occur independently
of vacuole enlargement [71,72]. Several studies have seen
no difference between the abundances of V-ATPase on the
vacuoles of wild-type and fab1 yeast, so the acidification
defect in cells that cannot make PtdIns(3,5)P2 is not simply
due to mislocalization of the V-ATPase [34,43]. This defective
acidification phenotype also occurs in C. elegans and in
D. melanogaster cells where the Fab1p pathway is defective:
Lysotracker failed to stain the endo-lysosomes or lysosomes
in worms or flies encoding a partially functional PtdIns3P
5-kinase [21]. The PtdIns(3,5)P2 dependence of endo-lysosome
acidification therefore appears to be widespread, but it has
not been reported in knockout mice, probably because the
PtdIns(3,5)P2 complements in the vac14−/− and fig4−/− mice were
only modestly compromised (∼ 50 % of wild-type) [17,31]. Since
c The Authors Journal compilation c 2009 Biochemical Society
8
S. K. Dove and others
less PtdIns(3,5)P2 is required to support vacuole acidification
than to preserve normal vacuole morphology and to localize
Atg18p to the vacuole [1], it is not surprising that vacuolar
acidification defects have not been observed in these mutant
organisms. Thus some mutants, such as yeast vac14 cells
expressing a single copy of FAB1, can have substantially enlarged
vacuoles that are still acidic [98].
Since vacuolar acidification is essential, like Atg18p localization, for vacuolar fission, it is perhaps not surprising that
PtdIns(3,5)P2 regulates both processes [98]. How PtdIns(3,5)P2
regulates vacuole acidification is presently unknown, but is likely
to involve undefined effector proteins that somehow activate
the V-ATPase either directly or indirectly.
ROLE OF PtdIns(3,5)P 2 IN STRESS SIGNALLING/ADAPTATION
Another process that probably involves CORVET, the V-ATPase
and PtdIns(3,5)P2 is stress signalling to the vacuole, especially
in response to hyperosmotic stress. Stress signalling occurs in
response to sudden changes in the extracellular environment
and allows a cell to respond with concomitant adjustments in
metabolism and physiology [7,77]. Many of the responses to
stress are well characterized and include the induction of heat
shock and other chaperone-like proteins as well as up-regulation
of genes, such as catalase, by the general stress-response pathway.
The sensors of the various stresses and the signalling pathways
they initiate are much less well understood [7].
The levels of PtdIns(3,5)P2 in yeast, plant and some animal
cells rise in response to heat, hyperosmotic and alkaline stresses
in a manner that suggests that this lipid may be a regulator
of stress-driven cell responses [16,35]. In the case of hyperosmotic
stress, the rise is transient, with the duration and magnitude of the
accumulation of PtdIns(3,5)P2 proportional to the degree of
hypertonicity. Modest stresses (e.g. 0.4 M NaCl) provoke small
PtdIns(3,5)P2 accumulations that reverse within ∼ 10 min, but
more intense stresses (such as 0.9 M NaCl) provoke much larger
accumulations of PtdIns(3,5)P2 that take up to 40 min to revert to
pre-stress values ([99] and S. K. Dove, unpublished work).
How stress-induced increases in PtdIns(3,5)P2 occur is not
known: they could be caused by increased Fab1p activity or
decreased PtdIns(3,5)P2 5-phosphatase activity, for example of
Fig4p [99]. Vac14p, Vac7p and Fig4p are all required for the full
stress-induced accumulation of PtdIns(3,5)P2 [17], but this is not
informative because these proteins all seem to be required for full
Fab1p activity [96].
PtdIns(3,5)P 2 -DEPENDENT VACUOLE FISSION ACCOMPANIES
STRESSES
A consequence of transient PtdIns(3,5)P2 accumulation in
response to stress is a rapid Fab1p/PtdIns(3,5)P2 -dependent
fragmentation/fission of the vacuole (see Figure 6) [100]. This
also requires Atg18p, although a small amount of fragmentation
is still observed in its absence or in the absence of the CORVET
subunit Vps3p (F. Williams and S. K. Dove, unpublished work),
or of the V-ATPase [100].
The vacuole remains ‘locked’ in the hyperfragmented state for
at least 1 h even after the burst of PtdIns(3,5)P2 has passed [100],
for incompletely understood reasons. This process requires the
palmitoylated vacuolar casein kinase Yck3p, which traffics to
the vacuole via the AP-3 pathway: in yck3 cells the fragmented
vacuoles reassemble after only a few minutes [17]. The HOPS
complex Ypt7p effector Vps41p is phosphorylated by Yck3p
in response to hyperosmotic stress [101], and this might act to
prevent HOPS-mediated vacuole reassembly, so long as Yck3p is
c The Authors Journal compilation c 2009 Biochemical Society
present and active. Alternatively other Yck3p substrates might be
involved.
It has been widely assumed that vacuolar fragmentation in
response to hyperosmotic stress, which greatly decreases the
volume contained in the multiple vacuole sub-compartments, is
somehow adaptive and helps restore the tonicity of the cytosol
[102]. This is clearly possible, as considerable water remains
in the vacuole after even acute hyperosmotic stress [82]. Indeed,
the increase in vacuolar surface area is accompanied by a
considerable decrease in vacuolar volume and we can safely
assume that the excess vacuolar water is indeed extruded into the
cytosol. Sedimentation of yeast at high g forces can also activate
PtdIns(3,5)P2 accumulation, with concomitant fragmentation of
the vacuole (S. K. Dove, unpublished work). In a possibly
related phenomenon, hyperbaric stress to 40–60 MPa, provokes
a transient increase in vacuolar acidity [80,81]. It is tempting
to conclude that hyperosmotic and hyperbaric stresses trigger
a PtdIns(3,5)P2 -dependent increase in V-ATPase activity that
somehow drives vacuolar fission.
PtdIns(3,5)P 2 AND CONTROL OF EXOCYTOSIS
We now turn to other cell activities that appear to be influenced by PtdIns(3,5)P2 , but for which there are as yet no
known mechanisms. First, there seems to be a link between
PtdIns(3,5)P2 and the regulation of exocytosis in several contexts:
(i) addition of IRAP (insulin-regulated aminopeptidase) and
GLUT4 (glucose transporter 4) to the plasma membrane of
adipocytes [103]; (ii) delivery of multiple transporters to the
plasma membrane in response to activation of the protein kinase
SGK1 (serum- and glucocorticoid-induced protein kinase 1) [51];
and (iii) neurosecretion [82]. However, the relationship between
PtdIns(3,5)P2 formation and these process remains unclear,
although the hypothesis that PtdIns5P might regulate exocytosis
seems a plausible explanation [82]. The first observation was
that PIKfyve could be phosphorylated by PKB (protein kinase
B; also known as Akt) in response to insulin, apparently at a
phosphorylation site (Ser318 ) midway between the FYVE domain
and the DEP domain that is only present in animal PIPkIIIs
[103], but this is now in some doubt (see below). PKB-mediated
phosphorylation apparently enhanced PIKfyve kinase activity,
albeit quite modestly, but overexpression of a mutant PIKfyve
with an S318A mutation, unexpectedly led to enhanced delivery
of IRAP to the plasma membrane [103]. It has also been reported
that greatly overexpressing PIKfyve modestly inhibits exocytosis
from neurosecretory cells [80,81]. Since PtdIns3P is essential for
priming in neurosecretory cells, it is perhaps not surprising that
PIKfyve overexpression should turn out to be inhibitory as it will
effectively reduce PtdIns3P levels [80].
Reports of the effects of PIKfyve phosphorylation by SGK1
are more convincing [80]. SGK1 is a serine/threonine-directed
protein kinase that is activated in response to many different
types of stress (including osmotic stress), leading to increased
cell-surface residency of several ion channels and transporters,
presumably because of increased exocytosis from intracellular
pools [80]. In vitro, Ser318 of PIKfyve is phosphorylated by
SGK1 rather than PKB [104], and it seems that phosphorylation
of this site is required for the stimulatory effect of SGK1 on
the exocytosis of at least three membrane channels: the creatine
transporter SLC6A8 [81], the potassium channel KCNQ1 [105],
and the Na+ /glucose co-transporter SLGT-1 [84]. The SGK1
effect on these transporters is likely to involve PtdIns(3,5)P2
and/or PtdIns5P production, but the identity of the effector
mediating this response remains to be determined. Exactly how
PtdIns(3,5)P 2 and Fab1p/PIKfyve
the Ser318 phosphorylation influences PIKfyve activity is also
unknown.
PtdIns(3,5)P 2 , AP-1 AND Ent3p/Ent5p MAY CONTROL Chs3p
TRAFFICKING TO THE S. CEREVISIAE CELL SURFACE
Chs3p, a chitin synthase that makes yeast cell wall chitin,
undergoes regulated secretion to the mother-bud neck in
S. cerevisiae, where it strengthens this growing area of the cell
wall. This transport may be from an intracellular compartment
known as the ‘chitosome’ [106], possibly a specialized organelle
unique to yeasts, to the cell surface [84], but other studies have
suggested that Chs3p shuttles between the TGN and the Snc1ppositive post-Golgi endosome (see Figure 3) [69,70]. Chs3p
sequestration and secretion requires the hetero-tetrameric AP-1
adaptin complex which complexes protein cargoes to the surface
of clathrin-coated vesicles at the TGN [13]. Several studies now
link PtdIns(3,5)P2 and the putative PtdIns(3,5)P2 effectors Ent3p
and Ent5p to the regulation of Chs3p trafficking to and from
the cell surface [13]. From these studies, it emerges that at least
some AP-1 functions in yeast require PtdIns(3,5)P2 , but with
PtdIns(3,5)P2 participating downstream of AP1: deletion of any
AP-1 subunit decreases PtdIns(3,5)P2 synthesis [13]. It seems
that AP-1 is another regulator of Fab1p activity, and that this control is needed for AP1 to work properly [70]. In accordance
with this idea, fab1 cells have a defect in Chs3p recycling and
intracellular retention similar to that in AP-1 mutants [69]. Ent3p
and Ent5p have also been implicated in intracellular retention
of Chs3p [85,89]. These two epsin-like cargo adaptors have
previously been shown to bind clathrin, AP-1 and Gga2p [10]
and, as discussed above, are relatively non-specific PPIn binders.
PtdIns(3,5)P 2 AND RETROMER
PtdIns(3,5)P2 may also influence the recycling of M6R from the
early/late endosome to the TGN [24]. M6R binds precursors of
soluble lysosomal proteases, such as cathepsin D, and carries them
to late endosomes, whence they are transported to the lysosome.
In a Rab9-dependent process catalysed by the retromer complex,
the M6R is then rapidly recycled back to the TGN to pick up more
cargo [24]. Retromer comprises mVps35p (which binds cargo),
the SNXs (sorting nexins) 1/2 and 5/6 (which appear to form the
vesicle coat and to drive membrane deformation), and mVps29p
and mVps26p (which mediate the attachment of mVps35p to the
coat) [107].
The first connection between this process and Fab1p/PIKfyve
was the observation that PIKfyve can associate with and phosphorylate the Rab9 effector, p40 [89]. The effect of this phosphorylation was to promote p40 attachment to membranes and in
some way enhance endosome-to-TGN recycling of cargoes [34].
This result was borne out by a study which identified PIKfyve and
Rab9 as two of several proteins required for endosome-to-TGN
trafficking of a viral Gag protein [26]. Moreover, mammalian
HeLa cells treated with siRNAs to knockdown PIKfyve levels
display a defect in the recycling of M6R and sortillin from
endosomes to the TGN [57], as do vac14−/− mice [85].
Unexpectedly, however, treatment of NIH 3T3 cells with the
PIKfyve inhibitor YM201636 did not reproduce this phenotype:
their M6R trafficking was normal [65]. The authors of the inhibitor
study therefore ascribed the PIKfyve-dependent endosome-toTGN defect to an indirect effect of preventing other aspects of
membrane trafficking that require PtdIns(3,5)P2 . Given the variety
of trafficking routes that have been described, and their variations
between cell types, clear conclusions will have to await further
work, as will the identification of PtdIns(3,5)P2 effectors that
9
are involved. Two candidates have been suggested: WIPI-1,
an Atg18p-like PROPPIN [86], and SNX1 [89]. The PPInbinding specificities of both of these proteins are contested, with
evidence that they may also bind the more abundant PtdIns3P. The
case of WIPI-1 is open: all other PROPPINs bind PtdIns(3,5)P2
preferentially and we have no reason to expect WIPI-1 to behave
differently [108]. SNX1 contains a classical PX (Phox homology)
domain and binds PtdIns3P with a considerably higher affinity
than PtdIns(3,5)P2 [94].
One suggestion that has appeared in several places is that
defective endosome-to-TGN trafficking is likely to be responsible
for the vacuole enlargement in cells devoid of PtdIns(3,5)P2
[109]. This is unlikely, at least in yeast, for the following reason:
trafficking of Vps10p, the classical sortillin homologue, is well
mapped because it is the protein used originally to isolate the
retromer complex [1] and all the VPS proteins from yeast [1];
Vps10p recycles between the Pep12p-positive PVC endosome and
the TGN [110]. When we examined the localization of Vps10p in
fab1 cells, we saw no difference from its localization in wildtype cells (F. Williams and S. K. Dove, unpublished work). In
addition, retromer mutants have different phenotypes to those
defective in PtdIns(3,5)P2 biosynthesis [111]. Thus, although
PIKfyve may assist retromer in higher organisms, it is still
unclear if this depends upon lipid kinase activity and so upon
PtdIns(3,5)P2 .
CONCLUSIONS
Our view of the world of PtdIns(3,5)P2 , a rare but lively
phosphoinositide, continues to expand, and key patterns are
emerging. First, it is likely that PtdIns(3,5)P2 regulates more or
less the same core processes across the eukaryote super-kingdom,
although we badly need evidence from groups of organisms that
branched deeper in the evolutionary tree than fungi and metazoans
[1]. Secondly, the PROPPINs represent an evolutionarily conserved group of PtdIns(3,5)P2 effectors that may mediate both
PtdIns(3,5)P2 -dependent and -independent retrograde trafficking
at many sites in the cell, one of which is essential for autophagy.
Thirdly, outlines of mechanisms of PtdIns(3,5)P2 -dependent
retrograde trafficking from vacuole/lysosome to endosome and of
vacuole fission have emerged, and detail is now needed. Fourthly,
links between PtdIns(3,5)P2 and other cell machinery, including
endo-lysosomal pH control by V-ATPase and CORVET-regulated
processes are becoming apparent. Although there are still many
more hints and challenges than firm facts, we hope readers of the
present review will be as excited by the future prospects as we are.
ACKNOWLEDGEMENTS
We would like to thank Mark Lemmon, Peter Parker, Frank Cooke, Paul Whitley, Harald
Stenmark, Sylvie Urbe and Mike Clague for helpful and stimulating discussions over the
past decade. S. K. D. and R. H. M. would also like to thank past and present members of
the Phosphoinositide Laboratory for their contributions. We would also like to apologize
to those authors whose work has been omitted for reasons of space.
FUNDING
This work was supported by the Royal Society; the Biotechnology and Biological Sciences
Research Council [grant number BB/D522197/1]; the Wellcome Trust [grant number
WT076480MA]; the Adrian Brown postgraduate scholarship fund; and the Dorothy Hodgkin
award for postgraduate students. S. K. D. is a Royal Society University Research Fellow.
Appendix 1: Will the real yeast lysosome please step forward?
A significant impediment to understanding the role of Fab1p
and PtdIns(3,5)P2 in higher organisms is the fuzzy issue of
compartment identity in yeast; it is not always clear which yeast
c The Authors Journal compilation c 2009 Biochemical Society
10
S. K. Dove and others
compartment corresponds to the mammalian lysosome or to the
early, late or recycling endosomes [118]. Part of the problem arises
because these organelles in yeast are often defined on the basis of
genetic rather than morphological evidence.
The Snc1p (t-SNARE)-positive compartment that underlies
the plasma membrane in S. cerevisiae seems well placed to
be either an early or recycling endosome. This appears to be
confirmed by the observation that FM4-64 and other dyes that are
endocytosed seem to enter this compartment at early time points
[119]. Yet there appears to be no direct trafficking pathway from
this compartment to the cell surface; all cargo requiring transport
to the plasma membrane must first pass through the yeast Golgi
(see Figure 3). This has resulted in some individuals dubbing
this compartment the ‘post-Golgi endosome’ or PGE to avoid
confusion with mammalian early endosomes, and we shall do the
same [120].
In contrast, the Pep12p (t-SNARE)-positive compartments
adjacent to the vacuole do appear to connect bidirectionally with
the plasma membrane, at least via genetic evidence; the uracil
permease Fur4p can recycle back to the plasma membrane from
the Pep12p-positive compartment, seemingly without passing
through either the Snc1p-positive compartment or the Golgi (see
Figure 3) [121]. Yet the Pep12p-positive compartment is the site
of formation of MVBs, a function normally associated with late
endosomes in higher eukaryotes. Further obscuring this issue is
the fact that the Rab GTPase, Vps21p, controlling entry into the
Pep12p-positive compartment most closely resembles Rab5,
the Rab controlling the early endosome in animals [122]. An
additional layer of complexity is added by observations suggesting
that the Pep12p compartment and the MVB compartment are
not the same organelle, but that the former matures into the
latter [118]. On balance, the evidence appears to imply that
this compartment is, in fact an early endosome that can mature
or acquire late-endosome characteristics. Again, the Pep12ppositive compartment has been named the PVC (pre-vacuolar
compartment) or PVE (pre-vacuolar endosome) in an attempt to
sidestep this issue and avoid confusion and in the present review
we adopt this practise (see Figure 3).
The most enigmatic of all yeast organelles is the vacuole.
This compartment is very large, acidic (pH 6.0) and is a major
nutrient store. The vacuole has commonly been assumed to be
the yeast lysosome and functionally it may perform this role as
it is the site of protein and organelle degradation and contains
many proteolytic enzyme activities [123]. However, the vacuole
is vastly greater in volume than the endosome in yeast and the
pH is only as acidic as a mammalian endosome, as lysosomes
generally have pHs that are below 5.5. Most intriguing of all is the
fact that the vacuolar Rab is Ypt7p, a protein that is homologous
with mammalian Rab7. In eukaryotes, Rab7 controls entry to,
and is localized on, late endosomes. It seems very likely that the
yeast vacuole is in fact a modified late endosome or endosome–
lysosome hybrid and has no direct equivalent in animal cells.
Might this mean that the MVB machinery could catalyse the
inward invagination of the vacuole membrane as it does at
the PVE? We have in fact observed this phenomenon under certain
conditions (S. K. Dove, unpublished work).
Appendix 2: PtdIns(3,5)P 2 and autophagy
PtdIns(3,5)P2 , Fab1p and PROPPINs seem, in some fashion,
to be associated with autophagy, despite the fact that in
S. cerevisiae, fab1 cells have no gross defect in this process;
fab1 cells do display a kinetic defect in the rate of maturation of
Pho860p, a marker of autophagy, but it is not as profound as the
defect in atg18 cells (S. K. Dove, unpublished work). Fab1p
c The Authors Journal compilation c 2009 Biochemical Society
in D. melanogaster is indeed required for correct autophagy,
highlighting the fact that the morphological description of
autophagy in yeast and in higher animals appears to be different.
In yeast the double membrane-bound autophagosome appears
to fuse directly with the vacuole whereas, in animal cells, an
amphisome is first formed that then matures into an autolysosome
via fusion with the late endosome. Fab1p appears to be required
for this latter maturation effect because amphisomes accumulate
in the cytoplasm of fab1-deleted D. melanogaster cells. Since the
D. melanogaster study was performed on purely morphological
criteria it is difficult to know whether protein degradation is
inhibited in fab1 fly cells. Whether these differences between
autophagy in yeast and animals are real or are artefacts of the
disparate way the two processes have traditionally been studied
(by electron microscopy in animals and by yeast genetics in S.
cerevisiae) remains to be determined; yet if the vacuole is indeed
a late endosome, not a lysosome (see Appendix 1), perhaps yeast
genuinely do have a ‘shorter’ more truncated pathway that has
become PtdIns(3,5)P2 independent?
If this is the case, why do PROPPINs appear to have
PtdIns(3,5)P2 -dependent and -independent functions in many
organisms? Might this family of proteins mediate the same process
(retrograde trafficking) at a number of distinct sites inside the
cell? In this model, only one of these sites requires PtdIns(3,5)P2
to localize the PROPPIN, whereas others are independent of
this lipid and require other (protein or lipid) factors to recruit
the PROPPIN-associated recycling machinery. One of these
PtdIns(3,5)P2 -independent retrograde events may therefore be
required to sustain autophagy and the Cvt pathway.
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Received 25 September 2008/18 December 2008; accepted 22 December 2008
Published on the Internet 13 March 2009, doi:10.1042/BJ20081950
c The Authors Journal compilation c 2009 Biochemical Society