Abstract
Membrane contact sites between organelles are critical for the transfer
of biomolecules. Lipid droplets store fatty acids and form contacts
with mitochondria, which regulate fatty acid oxidation and adenosine
triphosphate production. Protein compartmentalization at lipid
droplet-mitochondria contact sites and their effects on biological
processes are poorly described. Using proximity-dependent biotinylation
methods, we identify 71 proteins at lipid droplet-mitochondria contact
sites, including a multimeric complex containing extended synaptotagmin
(ESYT) 1, ESYT2, and VAMP Associated Protein B and C (VAPB). High
resolution imaging confirms localization of this complex at the
interface of lipid droplet-mitochondria-endoplasmic reticulum where it
likely transfers fatty acids to enable β-oxidation. Deletion of ESYT1,
ESYT2 or VAPB limits lipid droplet-derived fatty acid oxidation,
resulting in depletion of tricarboxylic acid cycle metabolites,
remodeling of the cellular lipidome, and induction of lipotoxic stress.
These findings were recapitulated in Esyt1 and Esyt2 deficient mice.
Our study uncovers a fundamental mechanism that is required for lipid
droplet-derived fatty acid oxidation and cellular lipid homeostasis,
with implications for metabolic diseases and survival.
Subject terms: Fatty acids, Fat metabolism, Lipid signalling
__________________________________________________________________
Protein-mediated transport is implicated in trafficking fatty acids at
contact sites of lipid droplets and mitochondria. Here, the authors use
proteomics to catalogue the proteins at this contact site and report a
mechanism of fatty acid transfer that regulates fatty acid oxidation
and lipid homeostasis.
Introduction
Lipid droplets (LDs) and mitochondria are critical regulators of lipid
homeostasis and ATP production in eukaryotic cells^[62]1. LDs are
dynamic organelles that store lipids as metabolic fuels and release
fatty acids in times of energy demand via lipolysis. This is a highly
regulated process involving a complement of proteins that temporarily
transit or permanently reside on the surface of LDs^[63]2,[64]3. The
primary function of mitochondria is to meet the energy demands of cells
by providing ATP through oxidative phosphorylation of metabolic
substrates, including fatty acids. Both organelles are important for
human health, as mutations in genes encoding LD and mitochondrial
proteins are linked to metabolic diseases such as non-alcoholic fatty
liver disease (e.g., PNPLA3, TM6SF2, MBOAT7, HSD17B13)^[65]4–[66]6,
type 2 diabetes (e.g., NDUFC2, COX7A2, HSL, CIDEA/B/C)^[67]7–[68]10,
lipodystrophies (e.g., PLIN1, BSCL1/AGPAT2, BSCL2/seipin, MFN2,
CGI-58)^[69]11–[70]14 and autosomal recessive fatty acid oxidation
disorders (e.g., CPT1A, HADHA, ACADVL)^[71]15–[72]17.
The interaction between LDs and mitochondria are increased during times
of heightened energy demand, such as β3-adrenergic
stimulation^[73]18,[74]19 or exercise^[75]20, and during states of
increased reliance on fatty acid oxidation, such as
starvation^[76]21,[77]22 and cold adaptation^[78]23,[79]24. This has
led to the view that close LD-mitochondria contact is essential for
efficient β-oxidation and protection against the accumulation of
cytosolic fatty acids and cellular lipotoxicity. In agreement with
this, extensive LD and mitochondria contact is evident in metabolically
active tissues including skeletal muscle^[80]25, heart^[81]26,
liver^[82]27 and brown adipose tissue^[83]23,[84]28 and proteins
including PLIN5^[85]23,[86]29, SNAP23^[87]30, FATP4 (ACSVL4)^[88]31,
VPS13C^[89]32, VPS13D^[90]33, and Rab8a^[91]34 appear to mediate
LD-mitochondria interactions.
Although the importance of these organelles and their interaction is
well recognized, we know little about the molecular machinery and
processes by which fatty acids stored within LDs are transferred to
mitochondria for their eventual oxidation. Protein-mediated lipid
transport routes have been implicated in trafficking fatty acids at the
membrane contact site of LDs and mitochondria and these proteins
possess several important features including the capacity to extract
lipids from a donor membrane; to shield lipids from the aqueous
cytosolic environment, typically within their hydrophobic pockets; and
to deliver lipids to a target membrane^[92]35. A recent study indicates
that VPS13D meets these criteria and can facilitate fatty acid transfer
at LD-mitochondria contact sites by mediating ESCRT-dependent
remodeling of LD membranes^[93]33. While the details of proteins
residing in the membrane contact sites of LDs and mitochondria are
beginning to emerge, an unbiased inventory of proteins that reside at,
or are temporarily recruited to these contact sites, is necessary to
better understand this critical pathway of fatty acid trafficking and
lipid metabolism.
Herein, we sought to unravel the mechanism of fatty acid transfer from
their site of storage to their site of oxidation. Starting with
unbiased proximity labeling proteomics in mammalian cells, we
identified a multimeric protein complex containing ESYT1, ESYT2 and
VAPB, which we further showed to localize to tripartite LD,
mitochondria, and endoplasmic reticulum (ER) interaction sites. This
complex appears to facilitate the transfer of fatty acids from LDs to
mitochondria, thereby regulating efficient fatty acid oxidation and the
maintenance of cellular lipid homeostasis.
Results
Proteins residing at lipid droplet-mitochondria contact sites
Lipid transfer between organelles requires spatial organization of
proteins at lipid contact sites^[94]36. To identify and map proteins at
LD-mitochondria contact sites that is known to reach 10 to 30 nm wide,
we used BioID proximity labeling. This method takes advantage of the
ability of BirA* to conjugate biotin onto local proteins^[95]37–[96]39.
We used small and monomer fluorescent proteins mScarlet^[97]40 and
mTurquoise2^[98]41 (unlike tetramaric GFP), and small targeting signal
peptides (10–30 amino acids, Fig. [99]1a, b) to localize BirA* to
either LDs or mitochondria. We also used multiple BirA* tag proteins to
internally validate the interface proteome (Fig. [100]1. a, b, e).
Initially, we engineered the C-terminal portion of BirA* to target LDs
by fusing to the transmembrane domain of methyltransferase like protein
AAM-B (also called METTL7B; AAM[TMD]-RFP-HA-BirA*) (Supplementary
Fig. [101]1a), which was shown previously to specifically target
constructs to LDs, but not to endoplasmic reticulum, as is the case for
full length AAM-B (ER)^[102]42. This was confirmed by Airyscan
super-resolution confocal microscopy staining with BODIPY (staining
LDs), PLIN2 (constitutive LD localized protein), and the ER-specific
protein calnexin. We show prominent ring like structures of BirA on the
surface of LDs, with evidence of very limited staining on the ER
(Supplementary Fig. [103]1c, d). We also targeted BirA* to the
cytosolic side of the outer mitochondrial membrane using the targeting
signal of FIS1 (Supplementary Fig. [104]1b). Mitochondrial localization
of the FIS1[TMD]-CFP-FLAG-BirA* constructs were confirmed by Airyscan
super-resolution confocal microscopy staining with TOMM20
(Supplementary Fig. [105]1e). Protein biotinylation was confirmed at
the surface of organelles, with some diffuse biotin staining, which is
likely indicative of on/off organellar interactions and/or protein
movement within the cell (Supplementary Fig. [106]1f–h). All constructs
were stably expressed in HepG2 cells.
Fig. 1. Identification of LD-mitochondria interface proteins using proximity
proteomic screens.
[107]Fig. 1
[108]Open in a new tab
a BioID targeted to the surface of LDs, containing the transmembrane
domain of AAM-B, RFP (mScarlet), HA, and BirA* (AAM[TMD]-RFP-HA-BirA*).
b BioID targeted to the surface of the outer membrane of mitochondria,
containing the transmembrane domain of FIS1, CFP (mTurquoise2), FLAG,
and BirA* (FIS1[TMD]-CFP-FLAG-BirA*). c Mass spectrometric analysis of
biotinylated proteins purified from HepG2 cells stably expressing
AAM[TMD]-RFP-HA-BirA*chimeras (LD-BioID). Plot compares BirA* to the
negative control in HepG2 cells supplemented with biotin. Colored dots
indicate proteins enriched using the three independent constructs
(a–e). Proteins known to be enriched at LDs (ArtGAP1, SNX, FASN),
peroxisomes (GOSAR1, ABCD3), endoplasmic reticulum (ESYT1, ESYT2, RTN4)
and mitochondria (DNM1L, CSDE1, OCIAD1) are highlighted. d Mass
spectrometric analysis of biotinylated proteins purified from HepG2
cells stably expressing Fis1[TDM]-CFP-FLAG-BirA* (Mito-BioID).
Experiment and data analysis as indicated in (c). e Design of the
Split-BioID approach, containing the transmembrane domain of FIS1, CFP,
FLAG, and half of BirA*(BirA[N]*) targeted to the surface of the outer
membrane of mitochondria (Fis1[TMD]-CFP-FLAG-BirA[N]*). The
transmembrane domain of AAM-B anchors RFP, HA, and the other half of
BirA*(BirA[C]*) targeted to the surface of LDs (AAM[TMD]-RFP-HA-BirA*).
f Airyscan imaging of HepG2 cells stably expressing the Split-BioID.
Merged image showing Fis1[TMD]-CFP-FLAG-BirA[N]* (magenta) and
AAM[TMD]-RFP-HA-BirA[C]*(green). (representative of 4 independent
experiments, scale bar, 2 μm). g Mass spectrometric analysis of
biotinylated proteins purified from HepG2 cells stably expressing
Split-BioID and negative controls (non-BirA*-transfected HepG2 cells).
Colored dots indicate commonly enriched proteins in panels C & D. h
Venn diagram showing the number of proteins enriched using Split-BioID,
LD-BioID and mito-BioID approaches. i Dot-plot analysis of commonly
enriched proteins using Split-BioID, LD-BioID and mito-BioID
approaches. j Validation of proteins identified by BioID using an
organelle coprecipitation assay of HepG2 cell lysate. LD-mitochondria
fraction (LD+Mito), mitochondrial fraction (Mito), and cytosol fraction
(Cyto) were examined by immunoblot. Representative of n = 3 biological
replicates. For c, d, g and i, data analyzed with unpaired two-tailed
Student’s t-test with FDR < 0.05, n = 4 biological replicates. See also
Supplementary Fig. [109]1 and Supplementary Data [110]1. Created in
BioRender. Keenan, S. (2025) [111]https://BioRender.com/n75s805.
Biotinylated proteins were then recovered on streptavidin beads in the
presence of 1% sodium dodecyl sulfate (SDS) and 6 M urea to eliminate
nonbiotinylated protein-protein interactions, which were detected by
mass spectrometry (MS) and quantified. HepG2 cells treated with biotin
in the absence of BirA* were used as negative controls for all
experiments. Enrichment of endogenous biotinylated proteins was not
different between wildtype cells and BirA*-expressing cells
(Supplementary Fig. [112]1i). Using the full length BirA* targeted to
LDs, we identified 974 proteins (Fig. [113]1c), of which 174 and 307
were previously reported using APEX proximity labeling of LDs in Huh7
and U2OS epithelial cells, respectively^[114]43 (Supplementary
Fig. [115]1j). We detected proteins known to reside at LDs (ArtGAP1,
SNX, FASN), peroxisomes (GOSAR1, ABCD3), endoplasmic reticulum (RTN4,
VAPB, ESYT1) and mitochondria (DNM1L, CSDE1, OCIAD1) (Fig. [116]1c).
Using full length BirA* bait targeted to mitochondria we identified 625
proteins that reside near the mitochondria outer membrane
(Fig. [117]1d).
We also employed a ‘split’ version of the promiscuous biotin ligase,
BirA*, where inactive halves of the ligase are fused to two different
bait proteins. When baits interact the halves come together to form an
active BirA* that conjugates biotin onto proteins within 10–30 nm of
the interaction site^[118]37–[119]39,[120]44,[121]45. We targeted the
N-terminal portion of BirA* to the cytosolic side of the outer
mitochondrial membrane by fusing to the transmembrane domain of
FIS1^[122]46 (FIS1[TMD]-CFP-FLAG-BirA*[N]), and the C-terminal portion
of BirA* was targeted to LDs by fusing to the transmembrane domain of
methyltransferase like protein AAM-B (AAM[TMD]-RFP-HA-BirA*[C])
(Fig. [123]1e, f). Hence, when co-expressing these constructs BirA* is
functional only at sites where LDs and mitochondria are in close
proximity. Note that these experiments were conducted before the
development of split-TurboID^[124]47, hence the use of split-BioID.
Using streptavidin conjugated horseradish peroxidase and western
blotting, we confirmed that when expressed alone neither half is
active, whereas co-expression leads to efficient protein biotinylation
(Supplementary Fig. [125]1k). Using this approach, we identified 81
proteins localized to the LD-mitochondria interface in live cells
(Fig. [126]1g). PCA analysis showed that while the proteomes identified
using independent approaches are distinct (Supplementary Fig. [127]1l),
71 proteins were commonly enriched in all three proximity biotinylation
experiments (Fig. [128]1h), many of which have been reported at other
organelle contact sites, including the ER. This was unsurprising given
the close association between LD-mitochondria-ER contact
sites^[129]48,[130]49. Given that three independent BirA* constructs
were used for identification, we define these proteins as
LD-mitochondria interface proteins. A complete list of all identified
proteins can be found in Supplementary Data [131]1.
LD-mitochondria contact proteins exert diverse functions
Pathway enrichment analysis of the 71 high-confidence LD-mitochondria
proteome using Gene Ontology (GO) and Kyoto Encyclopedia of Genes and
Genomes (KEGG)^[132]50,[133]51 databases showed enrichment of proteins
involved in protein processing at the ER, phagosome, endocytosis, and
glycerophospholipid metabolism, indicating diverse and unappreciated
functions at the LD-mitochondria interface (Supplementary Fig. [134]1m,
n). We identified known proteins involved in lipid transport (e.g.,
ESYT1 and ESYT2), membrane docking and priming contacts (e.g., VAPB),
metabolic sensors (e.g., RAB1A and RAB7, and their effector protein
ARHGDIA) and membrane structural proteins, such as MBOAT7
(Fig. [135]1i). Enrichment of these proteins was confirmed at the
LD-mitochondria interface using an organelle coprecipitation
assay^[136]23 and immunoblot analysis (Fig. [137]1j).
ESYT1, ESYT2 and VAPB form homo- and hetero-oligomeric complexes
Among the most highly enriched proteins at the LD-mitochondria
interface, we focused on ESYT1, which was previously reported to be
localized to ER-mitochondria contact sites and to tether the plasma
membrane to the ER to facilitate transfer of phospholipids via a highly
conserved lipid binding SMP-domain (Synaptotagmin-like, Mitochondrial
lipid binding Protein-domain)^[138]52–[139]54. We then set out to
determine whether ESYT1 forms a complex with other proteins at contact
foci of LDs-mitochondria. 3HA-eGFP-ESYT1 was expressed in HepG2 cells
under the PolG promoter to induce low level expression, where it had no
effect on mitochondrial metabolism^[140]55. Possible protein-protein
interactions were determined by HA-immunoprecipitation followed by
label-free mass-spectrometry. As expected, ESYT1 was highly enriched
(Fig. [141]2a, FDR p < 0.05) and efficiently co-isolated with known
interactors RDH11, YIF1B, ARL6IP5, and EXOS6^[142]56 (Supplementary
Fig. [143]2a and Supplementary Data [144]2). ESYT2 and VAPB were also
enriched with ESYT1 immunoprecipitation, both of which were also
identified in our LD-mitochondria interface proteome (Fig. [145]1g, i
and Supplementary Dataset [146]2). ESYT3 can form heterodimers and
heteromultimers with ESYT1 and 2 but was not detected in either the
BioID or immunoprecipitation experiments, likely owing to low
abundance. Heterodimerization of ESYT1 and ESYT2 is documented^[147]57,
however, VAPB interaction with ESYT1 and ESYT2 has not been
investigated (Supplementary Fig. [148]2b, c). To validate these
interactions, HA-tagged ESYT2 and VAPB were expressed in HepG2 cells
and subjected to immunoprecipitation and mass spectrometry. The
presence of ESYT1, ESYT2 and VAPB and other interactors were confirmed
in pulldowns from the independent cell lines (Fig. [149]2b, c,
Supplementary Data [150]2). Consistent with these results,
immunoprecipitation and immunoblot analysis from HepG2 cells confirmed
the interaction of endogenous ESYT1, ESYT2 and VAPB (Fig. [151]2d–f).
Next, we interrogated protein-protein interactions from
co-fractionation experiments performed on tissues from mice^[152]58. In
agreement with our data, ESYT1-ESYT2-VAPB co-fractionated in
high-molecular-weight pools from several tissues, including liver and
muscle where ESYT1 and VAPB had identical profiles (Supplementary
Fig. [153]2d).
Fig. 2. ESYT1, ESYT2 and VAPB form homo- and hetero-oligomeric complexes.
[154]Fig. 2
[155]Open in a new tab
a Volcano plot showing proteins associated with ESYT1. Data are from a
label-free proteomic analysis of anti-HA immunoprecipitates from HepG2
cells stably expressing 3HA-eGFP-ESYT1 versus 3HA-EGFP. p-values of
two-tailed Student’s t-test, n = 4 biological replicates. b, c Volcano
plot showing proteins associated with ESYT2 and VAPB using HepG2 cells
stably expressing 3HA-eGFP-ESYT2 and 3HA-eGFP-VAPB. Experiments and
statistical analysis as indicated in (a). d–f Western blot analysis of
ESYT1, ESYT2 and VAPB interactions following anti-HA immunoprecipitants
from HepG2 cells stably expressing 3HA-eGFP-ESYT1, 3HA-eGFP-ESYT2,
3HA-eGFP-VAPB or empty vector (EV; 3HA-EGFP) and probed for endogenous
proteins. MBOAT9 was used as a negative control. Representative of
n = 3 biological replicates. g, h Lifetime maps (g) of the FLIM data
acquisitions between eGFP-ESYT1, eGFP-ESYT2, and eGFP-VAPB (donor) with
VAPB-mCH (acceptor) pseudo-colored according to the FRET palette
defined in the phasor plot (h) (i.e., teal pixels = 0 % FRET while red
pixels = 23% FRET) that reports hetero protein-protein interaction. i
Hetero protein-protein interaction of ESYT1, ESYT2 and VAPB in HepG2
cells. From left to right columns: n = 21, 15, 16, 17, 18, 19 and 14
cells. j Fraction of heterocomplex between eGFP-ESYT1, eGFP-ESYT2, and
eGFP-VAPB with VAPB-mCH. From left to right columns: n = 16, 13, 13,
15, 15, 18, and 17 cells. k Structure of ESYT1 in complex with ESYT2.
Arrows- hydrophobic side chain residues at the interacting interface
that are identified via in silico saturation mutagenesis and selected
for further experimental validation of heterodimer formation. l
Quantification of the fraction of pixels exhibiting FRET between
eGFP-ESYT2 and wild type and mutant ESYT1-mCH (i.e., hetero
protein-protein interaction) across multiple HepG2 cells, from left to
right columns n = 5, 6, 8, 4, 6, 5, 4, 6 and 6 cells. Data analyzed
using unpaired two-tailed t-tests. *P < 0.05. The box and whisker plots
in 2i, j and l show the minimum, maximum and sample mean. In 2i, 2j and
2 l, *p < 0.05, unpaired t-test, two-sided. See also Supplementary
Fig. [156]2 and Supplementary Data [157]2.
To determine in a living cell whether ESYT1-ESYT2 form a heterocomplex
with VAPB, and investigate the stoichiometry of these three different
subunits, we labeled these proteins with eGFP (eGFP-ESYT1, eGFP-ESYT2,
eGFP-VAPB) or mCherry (mCH; ESYT1-mCH, VAPB-mCH) and employed
fluorescence lifetime imaging microscopy (FLIM) of Förster resonance
energy transfer (FRET) alongside fluorescence fluctuation spectroscopy
(FFS) to spatially map hetero- versus homo-oligomer protein-protein
interactions in HepG2 cells. The phasor approach to FLIM analysis of
(FRET) between eGFP-ESYT1, eGFP-ESYT2 and eGFP-VAPB (donor molecules)
with VAPB-mCH (acceptor molecule) (Fig. [158]2g, h), and eGFP-ESYT2
(donor molecule) with ESYT1-mCh (acceptor molecule) (Supplementary
Fig. [159]2e), revealed that ESYT1 and ESYT2 form hetero- and
homo-complexes in the absence and presence of forskolin stimulation.
Specifically, it was found that the hetero-complex between ESYT1 and
ESYT2 with VAPB is promoted by the protein kinase A (PKA)-activator
forskolin and VAPB self-associates into a dimer or higher order
oligomer independent of this stimulation (Fig. [160]2i). This result
was orthogonally validated by FFS based approaches called cross raster
image correlation spectroscopy (cRICS) (Fig. [161]2j) as well as number
and brightness (NB) analysis (Supplementary Fig. [162]2f, g).
To further assess heterodimer formation of ESYT1 and ESYT2, we next
performed in-silico saturation mutagenesis at the interface of modeled
structures of ESYT1 and ESYT2 (Fig. [163]2k and Supplementary
Fig. [164]2h). Here we assessed missense mutations based on their
predicted change in binding affinity (ΔΔG), which indicates a more
drastic effect on how ESYT1 interacts with ESYT2, as a guide for our
experiments. Mutations of hydrophobic side chains of ESYT1 (F152, L250,
I251) at the interface of ESYT1-ESYT2 to negatively charged amino acids
(Glu and Asp) were ranked among the topmost to impact the formation of
the heterodimer (Fig. [165]2k and Supplementary Fig. [166]2h). The
phasor approach to FLIM analysis of FRET between eGFP-ESYT2 (donor) and
ESYT1-mCH (acceptor) confirmed the presence of ESYT1-ESYT2 heterodimers
and that heterodimer formation was enhanced by PKA activation
(Fig. [167]2l). Consistent with the in-silico model, the phasor
approach to ESYT1/ESYT2 FLIM-FRET analysis showed that double and
triple residue mutations in ESYT1-mCherry (ESYT1^L250E, I251E-mCH and
ESYT1^L250E, I251E, F152E-mCH) significantly reduced heterodimer
formation with eGFP-ESYT2, both in presence and absence of PKA
activation (Fig. [168]2l). Together, these data demonstrate the
presence of ESYT1-ESYT2-VAPB multimeric complexes in mammalian cells
that are regulated by β-adrenergic stimulation. Since β-adrenergic
stimulation and PKA activation increases both lipolysis and
LD-mitochondria contact^[169]18,[170]59, this result collectively
suggests that ESYT1-ESYT2-VAPB complex formation is functionally
important at the LD-mitochondria interface.
ESYT1/2-VAPB complex forms at LD, mitochondria, and ER contact sites
Previously identified proteins mediating LD and mitochondria tethering,
including SNAP23, FATP4 (ACSVL4), VPS13 and VPS13D, and Rab8a are also
localized at the interface of other cellular organellar contact sites,
most notably the ER^[171]60–[172]67. As ESYT1 and ESYT2 are
predominantly localized to the ER where they form contact sites with
the plasma membrane or mitochondria^[173]52,[174]68, and in light of
our identification of ESYT1/2 at the LD-mitochondria interface, we
hypothesized that ESYT1, ESYT2, and VAPB complexes are in fact
localized at a tripartite organellar interface consisting of LDs,
mitochondria and ER. Consistent with this notion, Airyscan imaging and
line scanning analysis showed that eGFP-ESYT1, eGFP-ESYT2 and eGFP-VAPB
localize to the interface of LD-mitochondria-ER (Fig. [175]3a).
Formation of tripartite organellar interface consisting of LDs,
mitochondria and ER was also confirmed in HeLa and Hek293T cells
(Supplementary Fig. [176]3a, b). We confirmed localization of
endogenous ESYT1 and VAPB at the LD-mitochondria interface
(Supplementary Fig. [177]3c). Since ESYT1, ESYT2 and VAPB localization
has been well established in the ER, we next used the phasor approach
to FLIM-FRET analysis to monitor eGFP-ESYT1/eGFP-ESYT2 (donor)
interaction with VAPB-mCherry (acceptor) at LD-mitochondria contact
sites in HepG2 cells before and after forskolin administration. We used
an immunofluorescence mask defined by PLIN2-AF405 for lipid droplet and
TOMM20-AF633 for mitochondrial localization (Fig. [178]3b). The
interaction of ESYT1 or ESYT2 with VAPB was significantly increased at
the LD-mitochondria interface after forskolin treatment, but not under
non-stimulated conditions (as compared to whole cell FRET)
(Fig. [179]3c). This indicates that ESYT1/2-VAPB complex recruitment to
the LD-mitochondria interface is cellular signaling-dependent.
Fig. 3. ESYT1/2-VAPB form a complex at the LD-Mitochondria-ER interface.
[180]Fig. 3
[181]Open in a new tab
a Left panel: Single plane view of Airyscan imaging of HepG2 cells
stably expressing eGFP-ESYT, eGFP-ESYT2 or eGFP-VAPB (blue). HepG2
cells were fixed and stained for LDs with HCS LipidTox red Neutral
Lipid Stain (green), mitochondria with TOMM20 antibody (yellow), and ER
with Calnexin antibody (magenta). White arrows highlight tripartite
interfaces of LD, mitochondria, and ER. A total of 16 cells from four
independent experiments were imaged and a representative of image is
indicated. Large scale bars, 2 μm and inset scale bars 0.5 μm. Right
panel: Line scan analysis of images presented to the left. A and B on
x-axis corresponds to the line scan in panel a inset. b Left panel:
Donor lifetime maps of the FLIM data acquisitions between donor
eGFP-ESYT1, eGFP-ESYT2, and eGFP-VAPB with acceptor VAPB-mCH FRET pair
pseudo-colored according to the palette defined in the phasor plot in
(Fig. [182]2h) spatially map the hetero protein-protein interaction
(red). Right panel: Identification of the LDs and mitochondrial
interface based on PLIN2-AF405 and TOMM20-AF647 immunofluorescence
intensity mask analysis. c Quantification of the fraction of ESYT1/VAPB
(upper panel), ESYT2/VAPB (middle panel), and VAPB/VAPB interaction
inside the LDs and mitochondrial interface defined by PLIN2-AF405 and
TOMM20-AF647 intensity masks (panel B, right) versus whole cell protein
interaction fraction with and without Forksolin stimulation across
multiple HepG2 cells. ESYT1/VAPB -FSK n = 10; ESYT1/VAPB + FSK n = 7;
ESYT2/VAPB -FSK n = 10; ESYT2/VFSK+fork n = 9; VAPB/VAPB -FSK n = 9;
VAPB/VAPB + FSK n = 7 cells. *P < 0.05 paired t-test, one-tailed. See
also Supplementary Fig. [183]3.
Loss of functional ESYT1, ESYT2 or VAPB leads to enlarged lipid droplets
The presence of the ESYT1-ESYT2-VAPB complex at the LD-mitochondria
interface indicated a possible role in lipid metabolism. Accordingly,
we determined the effects of CRISPR-Cas9-mediated depletion of ESYT1,
ESYT2, or VAPB in HeLa cells (Fig. [184]4a). Because ESYT1 and ESYT2
can form both homo- and heterodimers, we also created an ESYT1-ESYT2
double knockout cell line (DKO). Airyscan imaging analysis showed that
ESYT1, ESYT2, VAPB and the DKO deletion increased total cellular LD
volume by 34-86% (Fig. [185]4b). This was due to increased LD size
rather than LD number for ESYT1, ESYT2 and VAPB KO cells. For DKO, LD
size was similar to control cells, but LD number was increased by 43%
(Supplementary Fig. [186]4a, b). These findings were confirmed by
transmission emission microscopy (Supplementary Fig. [187]4c).
Fig. 4. Impaired fatty acid metabolism in ESYT1/2-VAPB-deficient cells.
[188]Fig. 4
[189]Open in a new tab
a Representative western blot showing ESYT1, ESYT2 and VAPB protein in
HeLa cells following CRISPR-Cas9 mediated knockout of ESYT1, ESYT2,
VAPB and ESYT1 and ESYT2. GAPDH, loading control. * non-specific
immunoreactive band. b 3D rendering images of Control and KO cells
stained for LDs with HCS LipidTox Deep Red Neutral Lipid Stain (various
colors) and mitochondria with TOMM20 antibody (red). Prior to staining
ESYT1/2/VAPB^KO and Control HeLa cells were treated with 400 μM fatty
acids (oleic acid and palmitic acid; 2:1) for 4 h. Scale bars: 0.5 μm.
c Quantification of volume occupied by LDs per cell from experiments
shown in panel B. Control n = 35, ESYT1^KO n = 44, ESYT2^KO n = 42,
VAPB^KD n = 34, DKO n = 35 cells. d Lipidomic analysis showing
triacylglycerol content in HeLa cells treated with 400 μM oleic acid
and palmitic acid (2:1) for 4 h. n = 6 biological replicates. e, f HeLa
cells were pulsed for 6 h with radiolabeled fatty acid (^14C oleic
acid) conjugated to 1% BSA and chased for 4 h in low glucose medium
without fatty acids (starvation medium). Data indicates rate of ^14C-
oxidation (LD-derived fatty acid oxidation) and ^14C remaining in
triglyceride following the chase period. Bar graphs represent the
mean ± SEM from 3 (e) and 4 (f) biological replicate experiments. g
BODIPY C[16] in LDs of cells that were pulsed with BODIPY C[16] for
6 h, washed, and incubated in starvation medium for 4 h. n = 6
(VAPB^KD, DKO) or 7 (Control, ESYT1^KO, ESYT2^KO) biological
experiments. h Western blot showing steady state level of
phosphorylated and total ATGL and HSL protein in cells. * denotes
non-specific immunoreactive band. i Flux of ^13C fatty acids into TCA
cycle and non-mitochondria metabolites (n = 4 per group). The
^13C-fatty acid mix contained myristic (0.2%), palmitoleic (9.4%),
palmitic (38.9%), margaric (0.3%), linoleic (10.7%), oleic (26.9%),
elaidic (1.6%), and stearic (1.6%) acid. For (c– i) bar graphs
represent the mean ± SEM, *p < 0.05 vs. Control using one-way ANOVA
with Bonferroni’s multiple comparison test. See also Supplementary Fig.
[190]4 and Supplementary Data [191]3.
Given the differences in LD volume, we performed liquid chromatography
tandem mass spectrometry lipidomics to determine whether components of
the ESYT-VAPB complex impact acylglycerol abundance, which are the
major lipid type in LDs. Cells were incubated in medium containing
500 µM fatty acid mixture (oleate and palmitate, 2:1 molar ratio) to
enhance the detection of low abundance lipid species. Consistent with
the changes in LD volume, acylglycerol abundance was increased in KO
and DKO cell lines compared with control cells, including triglycerides
(29-57%, except VAPB^KO), diglyceride (32-56%) and monoglycerides
(203-283%) (Fig. [192]4d and Supplementary Fig. [193]4d, e). Together,
these data indicate that components of the ESYT-VAPB complex regulate
LD metabolism.
ESYT1, ESYT2 and VAPB deficiency reduce lipid droplet-derived fatty acid
oxidation
To test whether ESYT1, ESYT2 or VAPB regulates the oxidation of fatty
acids derived from LDs, we used ‘pulse-chase’ experiments where cells
were incubated with ^14C-oleate for 6 h to load ^14C-fatty acids into
triglycerides contained in LDs, after which the metabolic fate of ^14C
was traced in the absence of extracellular ^14C-fatty acids. The
oxidation of fatty acids derived from LDs was reduced by 28-47% in
ESTY1, ESYT2 and VAPB null cells, and by 67% in DKO cells
(Fig. [194]4e). The amplified impairment of fatty acid oxidation in the
DKO cells supports the notion that ESYT1 and ESYT2 form both homo- and
heterodimers for fatty acid transfer. Consistent with these results,
the ^14C-fatty acid remaining in triglyceride at the end of experiments
was increased in each of the KO cell lines (Fig. [195]4f). To further
demonstrate the importance of ESYT1-ESYT2-VAPB as a mediator of
LD-derived fatty acid oxidation, we incubated cells with fatty acid
C[16]-BODIPY for 6 h, then assessed LD-BODIPY degradation via its green
fluorescence 4 h later. The C[16]-BODIPY in LDs was increased by 27-47%
in KO cell lines compared with control cells (Fig. [196]4g). The
reduction in LD-derived fatty acid oxidation was unlikely to result
from insufficient triglyceride lipolysis as inferred by similar
contents and phosphorylation of activating serine resides of the
rate-limiting lipolytic enzymes ATGL (Ser^404) and HSL (Ser^660)
(Fig. [197]4h).
We performed orthogonal validation using ^13C-isotopologue profiling in
cells to track fatty acid carbons through the tricarboxylic acid (TCA)
cycle and other non-mitochondria metabolic processes. Isotopic
enrichment of ^13C-fatty acids (16:0, 16:1, 18:0 and 18:1) was not
significantly different in KO and control cells (Supplementary
Fig. [198]4f). Upon entry to the mitochondria, fatty acids undergo
β-oxidation which produces acetyl CoA composed of two ^13C labels that
can be further traced through the TCA cycle. In agreement with the
radiolabel experiments, m + 2 isotopologue enrichment in TCA cycle
metabolites such as citrate, fumarate, and malate were significantly
reduced in single KO cell lines, with this effect further amplified in
the DKO cells (Fig. [199]4i). Rerouting of ^13C through TCA cycle
anaplerotic pathways, including to aspartate and glutamate, was also
reduced in KO cell lines. Isotopologue labeling was near undetectable
in metabolites of carbohydrate and glutamine metabolism, indicating
specificity of this response (Supplementary Fig. [200]4g). Together,
these data demonstrate that each component of the ESYT1-ESYT2-VAPB
complex is necessary for efficient LD-derived fatty acid oxidation in
cells.
We next ruled out the possibility that ‘mitochondrial dysfunction’
could provide a mechanism for reduced fatty acid oxidation in KO cells.
The decrease in LD-derived fatty acid oxidation was not related to
changes in mitochondrial volume, as assessed by TOMM20 staining and the
expression of proteins that regulate oxidative phosphorylation, or
mitochondrial function, as assessed by measuring the rate of glucose
oxidation by radiometric techniques or oxygen consumption rate in live
cells (Supplementary Fig. [201]4h–l). LD-mitochondria contact is
postulated to facilitate efficient fatty acid oxidation. The number of
LDs in close contact with mitochondria (≤30 nm) was not different
between control and ESYT1 and ESYT2 knockout cells, and LD-mitochondria
contact was increased in VAPB and DKO cells, indicating that these
proteins are unlikely to be important for physical tethering of LDs to
mitochondria (Supplementary Fig. [202]4m). We also showed that the
levels of other fatty acid transfer proteins previously identified at
the LD-mitochondria interface^[203]33, including VPS13D, TSG101 and
ESCR-III (Supplementary Fig. [204]4n, o), were increased in ESYT
knockout cells compared with control cells.
SMP domain of ESYT1 and ESYT2 is required for trafficking of fatty acids from
LDs to mitochondria
The SMP domain of ESYT1 and ESYT2 binds glycerophospholipids and
transports these lipids between the ER and plasma
membranes^[205]52,[206]53,[207]69. Having shown localization of ESYT1
and ESYT2 at the LD-mitochondria interface and the necessity of these
proteins for LD-derived fatty acid oxidation, we reasoned that ESYT
proteins could transfer fatty acids between organelles. Computational
structural biology strategies were employed to test this possibility.
Homology modeling was carried out in Schrodinger Maestro to model the
ESYT1/ESYT2 heterodimer based on the experimental structure of ESYT2
asymmetrical dimer (PDB ID: 4P42). Two complexes were modeled, which
differed in the chain within the ESYT2 homodimer substituted by ESYT1
(Fig. [208]5a), which was possible due to homology (39% sequence
identity).
Fig. 5. SMP domain of ESYT1 and ESYT2 are required for fatty acid transfer
from LDs to mitochondria.
[209]Fig. 5
[210]Open in a new tab
(a) Apolar channel (yellow) formed by ESYT1 or ESYT2 SMP domain in
ESYT1/2 dimer complex. b-c Ligand docking analysis showing oleic acid
inside apolar channel of ESYT1 as a lateral (b) and frontal (c) view.
(d) Enrichment of oleic acid in 3HA-EGFP-ESYT2 cells (compared with
empty vector, EV, 3HA-EGFP) following HA immunoprecipitation and mass
spectrometry lipidomic analysis. Cells were incubated in 400 μM oleic
acid for 4 h prior to lysing. *p < 0.05 vs EV by unpaired two-tailed
t-tests. n = 4 biological replicates. Bar graphs represent the
mean ± SEM. e-f A comparison of SMP domain and key residues substituted
by heavy-side chain amino acid of ESYT1 that protrude into the apolar
channel to obstruct fatty acid trafficking. g Airyscan images of HeLa
cells showing lipid droplets in ESYT1^KO cells without or with
re-expression of wild type ESYT or mutations in the SMP domain. Cells
were treated with 400 μM oleic acid and palmitic acid (2:1) for 4 h
prior to staining. LD staining with HCS LipidTox Green (green) and DAPI
(Blue). Scale bar: 3 μm. h Fatty acid oxidation in HeLa cells as
described in G. n = 6 biological replicate experiments. i, j A
comparison of WT SMP domain (i) and a residue substituted by heavy-side
chain amino acid (I353Y; j) of ESYT2 that protrudes into the apolar
channel. k Airyscan images of HeLa cells showing lipid storage in
ESYT2^KO cells in comparison with ESYT2^KO cells with re-expression of
ESYT2^WT-eBFP or a mutation in the SMP domain (ESYT2^I354Y-eBFP). Prior
to staining cells were treated as in g for LD staining with HCS
LipidTox Green Neutral Lipid (green) and nuclei staining with SYTOX
Deep Red (Blue). Scale bar: 5 μm. (l) Fatty acid oxidation in HeLa
ESYT2^KO cells in comparison with rescued cell lines with expression of
wild type or mutant ESYT2 on SMP domain. n = 4 biological replicates.
In (h, i) * P < 0.05 using one-way ANOVA with Bonferroni’s multiple
comparison test. Bar graphs represent the mean ± SEM. See also
Supplementary Fig. [211]5.
Fragment screening analyses highlighted the presence of an apolar
channel within both complexes (Fig. [212]5a), which corresponded to the
apolar channel observed within Chain A of the ESYT2 homodimer. Within
both complexes generated, and similar to the ESYT2 homodimer, the
channel appears to be extended by the other subunit within the complex,
suggesting that passage of fatty acids may occur when ESYT1 and ESYT2
form a larger complex with VAPB. The observed apolar channel was
therefore considered as a point of fatty acid entry for further
transport (Fig. [213]5b, c and Supplementary Fig. [214]5a–l) and was
used as a search space for ligand docking of long chain fatty acids
(palmitic acid, C16:0; oleic acid, C18:1; linoleic acid; C18:2) and
other lipid types contained in LDs (triacylglycerol,
phosphatidylcholine and cholesteryl oleate). In both cases, the fatty
acids, which are released from LDs and are readily available for
transport, docked and fitted within the apolar channel, whereas complex
lipids such as tristearin (triacylglycerol) did not (Fig. [215]5a–c and
Supplementary Fig. [216]5a–t). These results suggested an important
functional role for the apolar channel in fatty acid binding. To
directly test this possibility, we performed a lipidomic analysis of
immunoprecipitated lipids from cells expressing ESYT2-HA or HA (empty
vector). ESYT2 was enriched for oleic and linoleic acid (Fig. [217]5d).
Notably, there was no enrichment of other lipids in ESYT-HA, including
components of LDs such as triacylglycerols, cholesterol esters and
phospholipids.
Further analysis identified residue pairs within the apolar channel of
each model located at proposed lipid entry, midway through, and at the
edge of the channel. Residues were tested for changes in protein
stability upon mutation to larger residues. Within the ESYT1 apolar
channel, mutations S172W, I294Y and S297Y resulted in moderate
increases in stability while the local rigidification predicted
possible impaired fatty acid transport (Fig. [218]5e-f and
Supplementary Fig. [219]5u). Similarly, within the ESYT2 apolar
channel, H231W at the entrance and I354Y located midway across the
channel, conferred mild increases in protein stability which is
predicted to block fatty acid entry or transport across the complex
(Fig. [220]5i, j and Supplementary Fig. [221]5v).
In light of these predictions, we evaluated the requirement of ESYT1
and ESYT2 fatty acid transfer capacity for β-oxidation by creating
point mutations designed to produce aromatic residues that block the
lipid binding pocket in ESYT1 or ESYT2 (Fig. [222]5e, f, i, j). We
expressed either eGFP-ESYT1^WT or mutant eGFP-ESYT1^S172W,
eGFP-ESYT1^I294Y or eGFP-ESYT1^S297Y in ESYT1^KO cells. Compared with
ESYT1^KO, eGFP-ESYT1^WT reduced LD volume (Fig. [223]5g) and increased
fatty acid oxidation (Fig. [224]5h), while expression of the ESYT1
mutants failed to rescue the ESYT1^KO phenotype (Fig. [225]5g, h).
Similarly, expression of ESYT2^WT-eBFP in ESYT2^KO cells decreased LD
volume, but not LD number (Supplementary Fig. [226]5x, y), and
increased fatty acid oxidation when compared to the parental ESYT2^KO
cells (Fig. [227]5l). Expression of ESYT2^I354Y-eBFP had no effect on
these parameters (Fig. [228]5k, l, Supplementary Fig. [229]5x, y).
Together, these data demonstrate a role of ESYT proteins in fatty acid
transport.
ESYT-VAPB complex proteins protect cells from lipotoxicity
Lipotoxicity refers to the detrimental effects of lipid metabolites on
cellular functions in non-adipose tissues and is prevalent in metabolic
diseases such as obesity and type 2 diabetes^[230]70,[231]71.
Lipotoxicity can result from excess fatty acid flux into cells and/or
uncoupling of LD lipolysis and mitochondrial β-oxidation
(Fig. [232]6a)^[233]70,[234]72. To investigate whether components of
the ESYT/VAPB complex are required to prevent cellular lipotoxicity,
HeLa cells were incubated in medium containing 500 µM palmitate
(complexed with albumin), a fatty acid known to induce stress signaling
pathways and cell death^[235]72,[236]73. ESYT1, ESYT2 and VAPB deletion
resulted in marked remodeling of the cellular lipidome including
increases in triglycerides, as well as lipids known to induce cell
stress damage, such as diglycerides, ceramide, sphingomyelin,
cardiolipin and lysophosphatidylcholine (Fig. [237]6b–f). In comparison
to wildtype cells, endoplasmic reticulum stress was increased in all KO
cell lines as indicated by increased phosphorylation of the UPR signal
activators ATF6 (57–89%) and protein kinase R-like ER kinase (PERK,
~40%), and by reduced protein disulfide isomerase (PDI, ~30%) and
binding immunoglobulin protein (BiP, 37–65%) levels (Fig. [238]6g).
Activation of other stress-regulated protein kinases, such as c-Jun
N-terminal kinase (JNK, 50–125%) and extracellular signal-regulated
kinase (ERK, 63-95%), were also increased in KO cells (Fig. [239]6h).
Sensitivity to palmitate-induced lipotoxic stress in KO cells extended
to activation of pyroptosis, which is an inflammatory form of
programmed cell death^[240]74. Specifically, ESYT1, ESYT2 and VAPB
deletion resulted in activation of the canonical inflammasome pathway
as indicated by increased caspase 1 cleavage, an increase in cleaved
gasdermin D (GSDMD) and increased levels of the damage-associated
molecular pattern high mobility group box 1 (HMGB1) (Fig. [241]6i).
Re-expression of ESYT2^WT, but not ESYT2^S297Y, in ESYT2^KO cells
ameliorated JNK phosphorylation and GSDMD cleavage (Supplementary
Fig. [242]5z). Taken together, these data demonstrate that components
of the ESYT-VAPB complex are required to prevent lipotoxicity in cells.
Fig. 6. ESYT1, ESYT2 or VAPB depletion sensitizes cells to palmitic
acid-induced lipotoxicity and cellular stress.
[243]Fig. 6
[244]Open in a new tab
a Schematic representation of fatty acid trafficking at LD,
mitochondria and ER contacts in the presence of protein-mediated fatty
acid transfer (efficient fatty acid trafficking) and with inefficient
fatty acid trafficking mediated by disruption to the ESYT/VAPB complex.
Created in BioRender. Keenan, S. (2025)
[245]https://BioRender.com/n75s805. b–f Lipidomic analysis showing
abundance of lipids associated with lipotoxicity including
diacylglycerol, ceramide, sphingomyelin (SM), cardiolipin, and
lysophosphatidylcholine (LPC). Bar graphs represent the mean ± SEM from
6 biological replicate experiments. *p < 0.05 vs. Control using one-way
ANOVA with Bonferroni’s multiple comparison test. g Representative
Western blot and quantitation of ER stress marker proteins. Bar graphs
represent the mean ± SEM from n = 4 biological replicate experiments of
BIP, ATF5, p-PERK (Thr980), PERK and PDI. ATF5[s]/ATF6[FL] indicates
ratio of protein content of short and full length ATF6. GAPDH was used
as loading control. Data analysed using one-way ANOVA and a
Kruskal-Wallis post hoc test. p < 0.05 vs. Control. (h) Representative
Western blot and quantitation of p-ERK (Thr202/Tyr204) and ERK, p-JNK
(Thr183/Tyr185) and JNK. GAPDH was used as a loading control. Bar
graphs represent the mean ± SEM. (p-JNK/JNK n = 4 and p-Erk1/2 / ERK1/2
n = 5 independent experiments). Data analysed using one-way ANOVA and a
Kruskal-Wallis post hoc test. *p < 0.05 vs. Control. i Representative
Western blot and quantitation of pyroptosis markers with palmitic
acid-induced stress in control and KO cells. In all experiments in this
panel, HeLa cells were treated with palmitate (500 µM) conjugated to 1%
BSA for 4 h. Bar graphs represent the mean ± SEM protein content of
HMGB1 (n = 6 for Control, ESYT2^KO, VAPB^KD and DKO, n = 5 ESYT1^KO),
cleaved caspase 1 (n = 4), ratio of full length and splice variant of
Gasdermin (n = 4) from independent experiments. GAPDH was used as
loading control. Data analysed using one-way ANOVA and a Kruskal-Wallis
post hoc test. *p < 0.05 vs. Control.
ESYT1 and ESYT2 regulate fatty acid oxidation in livers of mice
To determine the physiological relevance of ESYT1 and ESYT2 as a
regulator of fatty acid oxidation in vivo, we studied mice lacking
either Esyt1(Esyt1^KO) or Esyt2 (Esyt1^KO) in hepatocytes. To do this
we employed CRISPR gene-editing and identified guide RNA’s (gRNA)
targeting proximal regions of the mouse Esyt1 or Esyt2 gene. AAV
constructs were generated expressing two gRNA sequences targeting Esyt1
or Esyt2 alongside a Cre-dependent mCherry to report AAV-transduced
cells (AAV-gRNA-FLEX-mCherry, Fig. [246]7a). To target CRISPR-mediated
excision of Esyt1 in hepatocytes, we crossed Alb-Cre with LSL-Cas9-GFP
knock-in mice to generate Alb-Cre::LSL-Cas9-GFP mice, which
specifically expressed Cas9 and GFP in hepatocytes. Hepatocyte-specific
targeting and silencing of Esyt1 and Esyt2 was induced following
administration of AAV-gRNA-FLEX-mCherry into adult
Alb-Cre::LSL-Cas9-GFP mice, which was confirmed by mass spectrometry
proteomics and immunohistochemistry (Fig. [247]7b and Supplementary
Fig. [248]6a). Loss of Esyt2 reduced Esyt1 in hepatocytes, which is
consistent with the notion that protein stability is often contingent
on the presence of interacting partners. Loss of either Esyt1 or Esyt2
resulted in a ~50% reduction in fatty acid oxidation in isolated murine
hepatocytes (Fig. [249]7c). There were no differences between groups
for metabolic processes that influence LD volume including fatty acid
uptake, storage of fatty acids in triglycerides and cholesterol esters,
or de novo lipogenesis (Supplementary Fig. [250]6b, c). These data
provide direct evidence that ESYT1 and ESYT2 are critical for fatty
acid oxidation in vivo.
Fig. 7. ESYT1 and ESYT2 regulate fatty acid oxidation in vivo.
[251]Fig. 7
[252]Open in a new tab
a Schematic outlining strategy for CRISPR-Cas9 mediated deletion of
Esyt1 and Esyt2 in hepatocytes of mice. b Esyt1 and Esyt2 peptide
abundance in liver of mice. Data are mean ± SEM. *P < 0.05 vs. WT using
one-way ANOVA and a Kruskal-Wallis post hoc test. n = 5 Control, n = 3
ESYT1^KO, n = 3 ESYT2^KO (n = 5 ESYT1 peptide; n = 3 ESYT2 peptide),
ESYT1^KO (n = 3 for both ESYT1 and ESYT2 peptide), ESYT2^KO (n = 3 for
both ESYT1 and ESYT2 peptide). c Fatty acid oxidation in precision-cut
liver slices from Control, Esyt1^KO and Esyt2^KO mice. Data are
mean ± SEM. *P < 0.05 vs. WT using one-way ANOVA and a Kruskal-Wallis
post hoc test. n = 8 Control, n = 5 ESYT1^KO, n = 7 ESYT2^KO. d dEsyt2
mRNA in WT and dEsyt^KO and dEsyt^FB-KO Drosophila. Data are
mean ± SEM. *p < 0.05 vs. WT by unpaired two-tailed t-test. n = 3 per
group. e Breeding strategy for the generation of fat body-specific
dEsyt2 inhibition in Drosophila (left) and survival response of WT ^FB
and dEsyt^FB-KO males under starvation (right), n = 100 flies per
genotype. Data analyzed using Log-rank (Mantel-Cox) test. (f) Breeding
strategy for the generation of global dEsyt inhibition in Drosophila
(left) and survival response of WT and dEsyt^KO males under starvation
(right), n = 100 flies per genotype. Data analyzed using Log-rank
(Mantel-Cox) test. See also Supplementary Fig. [253]6. Created in
BioRender. Keenan, S. (2025) [254]https://BioRender.com/n75s805.
To provide further evidence that ESYT proteins are critical regulators
of lipid metabolism in vivo, we silenced dEsyt in Drosophila. The
Drosophila genome encodes one ESYT orthologue^[255]54,[256]75 and
silencing was induced in the whole body (dEsytd^KO) and in the fat body
(dEsytd^FB-KO), which is a liver- and adipose-like tissue that stores
triglyceride. dEsyt silencing was confirmed by qPCR (Fig. [257]7d).
Triglycerides contained in the fat body are broken down by lipases and
the liberated fatty acids provide the main source of energy during
starvation^[258]76. In keeping with an important role of ESYT in
regulating lipid metabolism and survival, deletion of dEsyt in
Drosophila reduced survival in response to starvation (Fig. [259]7e,
f). While altered lipid metabolism can result in changes to starvation
resistance, the same is true for altered glucose metabolism or
perturbations to cellular membrane function. Therefore, we cannot
discount the effects of removing dEsyt on other processes that
influence starvation resistance.
Discussion
Using unbiased proximity proteomics, we profiled the proteins residing
at the interface of LDs and mitochondria. Within this proteome, we
identified a multimeric protein complex consisting of ESYT1 and/or
ESYT2 and VAPB, which facilitates mitochondrial oxidation of fatty
acids derived from LDs and is essential to protect cells from
lipotoxicity. We show that this complex resides at the interface of a
tripartite organellar contact involving the LD, mitochondria and ER,
and the interaction of these proteins is enhanced with β-adrenergic
stimulation, a state characterized by increased lipolysis of
triglycerides in LDs^[260]77. Deletion of ESYT/VAPB complex components
reduces the oxidation of fatty acids derived from triglycerides stored
in LDs, and results in the accumulation of enlarged LDs, which is a
defining feature of cells in organisms with metabolic
disease^[261]77,[262]78. The reduction in fatty acid oxidation most
likely results from loss of the lipid transfer function of ESYT1/2 and
not from alterations in mitochondrial mass or function, proteins that
regulate lipolysis (e.g., ATGL, HSL), or the tethering of LDs to
mitochondria. The observation that fatty acid oxidation is decreased in
the liver of mice with Esyt1 or Esyt2 deficiency reinforces the
importance of this function. These findings constitute a molecular
mechanism for fatty acid transfer in mammalian cells and identify a
function for ESYT in regulating cellular lipid metabolism.
Although complimentary approaches to manipulate ESYT functions and
multiple models revealed consistent findings on ESYT functions, our
study has raised several considerations. Deletion of both ESYT1 and
ESYT2 reduced LD-derived fatty acid oxidation by 67%, indicating that
additional mechanisms regulate fatty acid trafficking between the LD
and mitochondria. This is likely to include other proteins at
LD-mitochondria contact sites, including a fatty acid transport process
involving the coordinated actions of VPS13D and the endosomal sorting
complex required for transport (ESCRT) protein tumor susceptibility 101
(TSG101), which is invoked with starvation^[263]33. While, these
proteins were not identified in our proximity labeling of the
LD-mitochondria proteome, immunoblot analysis showed increased
abundance of these proteins with ESYT deletion. This indicates a
compensatory mechanism to retain fatty acid trafficking between LDs and
mitochondria in the absence of the complete ESYT-VAPB complex. Another
complex consisting of PLIN5 and FATP4^[264]79 was shown to enhance
LD-mitochondria contact and increase fatty acid oxidation, however,
direct evidence of fatty acid transfer by this complex requires
confirmation. Taken together, these findings support a model whereby
various protein complexes co-exist to coordinate membrane tethering
and/or inter-organellar fatty acid transfer for efficient oxidation,
and at least in the cell types examined herein, that the ESYT/VAPB
complex is requisite for efficient fatty acid metabolism and to prevent
cellular lipotoxicity. In addition, fatty acid transfer between
organelles would require the conversion of fatty acid to fatty acyl-CoA
esters^[265]80 for mitochondrial uptake. In this context, we identified
ACSL3 in the split-BioID analysis. This protein has long chain
acyl-coenzyme A synthetase activity and is likely to facilitate this
function.
ESYT and VAPB proteins are localized at contact sites of several
cellular organelles. ESYT has been reported at ER-plasma membrane and
ER-mitochondria contact sites^[266]53,[267]69,[268]81,[269]82, and both
VAPB and ESYT1/2 are enriched in other proximity based proteomic
mapping of LDs^[270]83 and the outer mitochondria membrane^[271]84.
VAPB has docking and tethering functions^[272]85, most notably with the
protein tyrosine phosphatase-interacting protein-51 (PTPIP51) to
regulate ER-mitochondria contacts^[273]85. In these cases, VAPB anchors
to the ER membrane using its C-terminal transmembrane domain while it
interacts with the other proteins or protein complex using its
N-terminal major sperm domain^[274]86. While our data show that ESTY1/2
and VAPB interact (i.e., by HA-affinity purification and mass
spectrometry, immunoprecipitation and immunoblot analysis, and
FLIM-FRET) and places the ESYT/VAPB complex at the interface of
LD-mitochondria-ER, we have an incomplete understanding of how the
components are organized (Fig. [275]8). We suggest that VAPB interacts
with mitochondria, while ESYTs bind the ER and VAPB to interact with
LDs to facilitate fatty acid transfer. Future studies using high
temporal and nanoscale spatial resolution approaches will be required
to provide further insights^[276]48,[277]49,[278]87.
Fig. 8. Proposed Model of ESYT1-ESYT2-VAPB Mediated Fatty Acid Transfer at
the LD-Mitochondria-ER Interface.
[279]Fig. 8
[280]Open in a new tab
Left panel: TEM image of HeLa cells showing an exploratory tripartite
interaction among LDs, mitochondria and the ER. Right panel:
Summarizing model of complex formation of ESYT1, ESYT2 and VAPB at the
interface of these organelles. Created in BioRender. Keenan, S. (2025)
[281]https://BioRender.com/n75s805.
The prevailing view that mitochondria-LD coupling promotes flux of
fatty acids from LDs for mitochondrial β-oxidation has been challenged
by emerging evidence indicating the existence of metabolically distinct
subpopulations of mitochondria that promote opposing functions of fatty
acid oxidation or fatty acid synthesis and/or LD biogenesis. Some
suggest that LD-associated mitochondria support higher rates of fatty
acid oxidation^[282]22,[283]33,[284]79,[285]88,[286]89, while others
indicate that LD-associated mitochondria have low fatty acid oxidation
capacity and support LD expansion by providing pyruvate-derived ATP for
triglyceride synthesis^[287]23,[288]48. In these latter studies, fatty
acids were shown to be selectively trafficked to and oxidized in
mitochondria that were not in contact with LDs^[289]23,[290]48. Our
data using complimentary fatty acid tracing methods show that
expression of wild type ESYT1/2, but not ESYT1/2 with impaired fatty
acid binding capacity, is required for LD-derived fatty acid oxidation,
which supports the premise that LD-associated mitochondria are
important for cellular fatty acid oxidation.
There are several limitations to this study. While many lines of
evidence strongly support a role for ESYTs in lipid transport, direct
biochemical evidence or single-particle imaging for ESYT-mediated fatty
acid transfer remains to be established. Additionally, alternative
mechanisms could explain ESYT/VAPB effects on fatty acid oxidation,
perhaps via phospholipid transfer from LDs to ER, and such
possibilities require evaluation.
Impaired fatty acid metabolism and lipotoxicity are a common feature of
metabolic diseases, including obesity, non-alcoholic fatty liver
disease, diabetes, some cancers, and neurodegenerative
diseases^[291]49,[292]70,[293]71,[294]90–[295]92. ESYT deletion results
in susceptibility to cellular stress and lipotoxicity in cells,
suggesting that impaired ESYT function may contribute to excessive
lipid accumulation in tissues and disease pathogenesis. Further, our
indexing of proteins residing at LD-mitochondria contact sites provides
a valuable resource for further research that could lead to strategies
to mitigate metabolic and other disease states.
Methods
Cell culture
HeLa, HepG2 and HEK293T cells were cultured in high glucose and
GlutaMAX (Thermo Fisher Scientific; 11965092) supplemented with 10%
fetal bovine serum (FBS; Cell Sera; AU-FBS/PG) and 1%
Penicillin-Streptomycin (10,000 U/mL; Thermo Fisher Scientific;
15140122) at 37 °C in a 5% CO[2] incubator. For starvation experiments,
cells were incubated in DMEM low glucose (5.5 mM) (Thermo Fisher
Scientific; 11054020) supplemented with glutamine for the indicated
time.
Molecular cloning
Plasmids generated for this study were assembled using the Gibson
assembly method^[296]93 and Q5® Site-Directed Mutagenesis Kit (E0554)
from New England Biolabs (NEB). DNA was transformed into NEB® 5-alpha
Competent E. coli cells via heat shock at 42 °C for 30 sec, followed by
incubation for 2 min on ice and 1 h at 37 °C in Luria-Bertani (LB) on
an Eppendorf Thermomixer shaker set to 300 rpm. Transformed bacteria
were spread onto LB plates (containing either Ampicillin or Kanamycin)
and incubated at 37 °C overnight. DNA was isolated from the resulting
colonies using QIAprep Spin Miniprep Kit (Qiagen) and sequenced to
verify the insertion of the respective DNA.
Transfections and transductions
All stable cell lines were produced by transducing with lentiviral or
retroviral particles followed by either antibiotic selection or
fluorescence-activated cell sorting. Lentiviral particles were produced
by transfecting HEK293T cells in 6-well plates with 1.5 μg of vector
and 1.0 μg and 0.5 μg of the packaging plasmids (i.e., proportions
3:2:1) using Lipofectamine LTX. The packaging plasmids for lentiviral
transduction were psPAX2 (Addgene #122600) and pMD2.G (Addgene #12259).
The packaging plasmids for retroviral transduction were pUMVC3 and
pCMV-VSV-G plasmids (Addgene #8449 and 8454). The transfection media
was removed after 16 h and virus-containing supernatants was collected
after 48 h and filtered using a 0.45 μM PES filter (Merck). Target
cells were transduced in 6-well plates using 500 μL of viral
supernatant and 500 μL of fresh medium supplemented with 8 μg/mL of
polybrene (Sigma-Aldrich). Selection with the appropriate antibiotic or
fluorescence sorting was initiated after 48 h.
Generation of KO cells
ESYT1^KO, ESYT2^KO and VAPB^KD HeLa cells were generated by the
CRISPR/Cas9 system. The DNA oligonucleotide sequence (Star Method) was
chosen as a guide RNA using CHOPCHOP. Two gRNAs oligos were designed
and separately cloned into All-in-one plasmid system
(pSPCas9(BB)-2A-GFP vector; Addgene #48138). HeLa cells transfected
with the plasmids (GFP + ) were sorted into a 96 well plate using a
FACS ARIA III (BD Biosciences) at a density of 1 cell per well. ESYT1,
ESYT2 or VAPB deficiency was confirmed by immunoblot analysis and
Sanger sequencing. The gRNAs used were: TGTTTTCCCTTACCGGGCGTCGG and
CGCAAAACGCCATGTAGCTGAGG for ESYT1; AGTGGAACGGTGATTCGATTGGG and
GACACGCTTACCTTTGGTACAGG for ESYT2; CAGCACGAGCTCAAATTCCG and
TGAGCTCGTGCTGCGGCTCG for VAPB; and GGCTTCGCGCCGTAGTCTTA for
control/scramble.
Immunoblot analysis
Cells were washed once with ice-cold PBS, centrifuged at 800 × g for
5 min at 4 °C, and lysed in RIPA buffer (50 mM Tris HCL, pH 7.4; 150 mM
NaCl; 0.1% SDS; 0.5% sodium deoxycholate; 1% Triton X-100) supplemented
with protease inhibitor cocktail (cOmplete, Roche) and phosphatase
inhibitor (PhosSTOP, Roche) on ice. Cell lysates were clarified by
centrifugation, and the protein concentration was determined using a
BCA protein assay kit (Pierce, Thermofisher Scientific). After mixing
with 5X Laemmli buffer, samples were subjected to 10% or 12% SDS-PAGE.
Proteins were then transferred to a nitrocellulose membrane (0.45 µm,
#1620264; Bio-Rad) followed by blocking with 5% (wt/vol) skim milk in
TBS-T for 2 h at RT. Incubation with primary antibody was performed
overnight at 4 °C followed by three washes for 5 min each in TBS-T. The
following primary antibodies were used for immunoblotting: ESYT1
(ab118805, Abcam, 1:1,000), ESYT2 (HPA002132, Sigma-Aldrich, 1:1000),
VAPB (ab241298, Abcam, 1:1000), RAB1 (ab302545, Abcam, 1:1,000), RAB7
(ab137029, Abcam, 1:1,000), RAB7 (ab137029, Abcam, 1:1,000), MBOAT7
(ab262944, Abcam, 1:1,000), PLIN2 (ab220738, Abcam, 1:1,000), GAPDH
(ab9482, Abcam, 1:1,000), HMGB1 (ab18256, Abcam, 1:1,000), Gasdermin D
(ab209845, Abcam, 1:1,000), Total OXPHOS (ab110413, Abcam, 1:1,000),
ATGL (2138S, Cell Signaling Technology, 1:1000), HSL (4107 s, Cell
Signaling Technology, 1:1000), BIP (3177S, Cell Signaling Technology,
1:1000), p-PERK (Thr980, 3179S, Cell Signaling Technology, 1:1000),
PERK (3192S, Cell Signaling Technology, 1:1000), PDI (3501S, Cell
Signaling Technology, 1:1000), p-JNK (Thr183/Tyr185, 9251S, Cell
Signaling Technology, 1:1000), JNK (9252S, Cell Signaling Technology,
1:1000), p-ERK1/2 (Thr202/Tyr204, 9101S, Cell Signaling Technology,
1:1000), ERK1/2 (4695S, Cell Signaling Technology, 1:1000), cleaved
caspase-1 (89332S, Cell Signaling Technology, 1:1000), ATF6 (70B1413.,
Enzo Life Sciences, 1:1000) and Streptavidin (SA10001, Thermo Fisher
Scientific). Secondary antibodies were peroxidase-conjugated
anti-rabbit used at a 1:5,000 dilution at RT for 2 h. The bound
antibodies were detected by Clarity Western ECL Reagent (Bio-Rad) and
visualized with Molecular Imager® ChemiDoc™ XRS+ (Bio-Rad). Western
blots were quantified using Image Lab software (ver. 6.10 build 7,
Bio-Rad).
In cell biotin labeling with BioID and Split-BioID and mass spectrometry
analysis
For biotin labeling, we generated stably expressing full length BirA*
(AAM[TMD]-RFP-HA-BirA* or Fis1[TMD]-CFP-FLAG-BirA*) and split BirA*
(AAM[TMD] -RFP-FLAG-BirA[N] and BirA-C-HA-mTurquiose2CFP-Fis1[TMD]),
using pBMN-Z vector (Addgene#1734, #36047), pcDNA3.1- PP1g -Flag -
BirA* (AA2-140) (Addgene#86886), pcDNA3.1 - BirA* (AA141-321) - HA -
NIPP1 (Addgene#86885), pPalmitoyl-mTurquoise2 (Addgene#36209) and
pCytERM_mScarlet_N1(Addgene#85066). We used the first (E140/Q141) of
several described splitting sites to generate the two BirA*
fragments^[297]44,[298]45. Cells were seeded at 50% confluency in a
10 cm petri dish the day prior to experiments. Biotin (10 mM) was
diluted in complete media and added directly to cells to a final
concentration of 50 μM and incubated at 37 °C for 16 h. Non-transfected
HepG2 cells treated with 50 μM biotin for 16 h used as control. For
both immunoblot and mass spectrometry experiments, labeling was stopped
after the indicated time periods by transferring cells on plates to ice
and washing gently with cold DPBS. Cells were dislodged by scrapping in
DPBS containing 1 μM EDTA.
For BioID experiments, cell lysates were obtained by centrifugation and
loaded consecutively on spin columns (Pierce) containing
streptavidin-coated beads prewashed with lysis buffer 1 (Pierce) and
biotinylated proteins were enriched^[299]94. The reduction of cysteine
bonds was mediated by 5 mM Tris(2-carboxyethyl)phosphine (TCEP) for
30 mins at 37 °C and alkylation with 10 mM iodoacetamide. Beads were
then resuspended in digestion buffer containing sequencing grade
modified trypsin (Pierce) at 37 °C overnight. After quenching with 0.2%
TFA, the samples were desalted by C18 reversed-phase spin columns
according to the manufacturer’s instructions (Pierce) and eluted with
0.2% TFA and 80% acetonitrile (ACN). The eluted peptide sample was
dried in a vacuum centrifuge and reconstituted to a final volume of
30 μl in 0.2% TFA and 1% acetonitrile (ACN). Analysis was performed on
a Q-Exactive Plus mass spectrometer (Thermo Fisher Scientific) coupled
to an Ultimate 3000 liquid chromatography system. Peptides were loaded
on to an Acclaim C18 PepMap nano Trap x 2 cm, 100 µm I.D, 5 µm particle
size, 300 Å pore size trap column (Thermo Fisher Scientific) at
15 µl/min for 3 min prior to switching the trap in line with the
analytical column (Acclaim RSLC C18 PepMap Acclaim RSLC nanocolumn
75 μm × 50 cm, PepMap100 C18, 3 μm particle size 100 Å pore size;
ThermoFisher Scientific). A 65 min 250 nl/min non-linear gradient
(buffer A, 0.1% formic acid, 2% ACN and buffer B, 0.1% formic acid, 80%
ACN) was used to separate peptides. Data were collected in positive
mode using Data Dependent Acquisition using m/z 375–1400 as MS scan
range, HCD for MS/MS of the 15 most intense ions with charge ≥ 2. Other
instrument parameters were: MS1 scan at 70,000 resolution (at 200 m/z),
MS maximum injection time 50 ms, AGC target 3E6, MS/MS resolution
17,500, MS/MS AGC target of 5E4, MS/MS maximum injection time 50 ms,
minimum intensity was set at 5E3 and dynamic exclusion was set to 30 s.
Raw files were analyzed using the MaxQuant platform^[300]95 version
1.6.5.0 searching against the UniProt human database containing 20,399
reviewed, canonical entries (January 2019) and a database containing
246 common contaminants. Default search parameters were used with
“Label free quantitation” set to “LFQ”. Trypsin/P cleavage specificity
(cleaves after lysine or arginine, even when proline is present) was
used with a maximum of 2 missed cleavages. Oxidation of methionine and
N-terminal acetylation were specified as variable modifications.
Carbamidomethylation of cysteine was set as a fixed modification. A
search tolerance of 4.5 ppm was used for MS1 and 20 ppm for MS2
matching. False discovery rates (FDR) were determined through the
target-decoy approach set to 1% for both peptides and proteins. Using
the Perseus platform^[301]96 version 1.6.5.0, proteins group LFQ
intensities were log2 transformed. Values listed as being “Only
identified by site,” “Reverse,” or “Contaminants” were removed from the
data set. Experimental groups were assigned to each set of
quadruplicates and the rows filtered to contain at least one group
where the number of valid values was 3. Using the “Subtract row
cluster” function, the dataset was normalized using to the average
abundance of cluster of endogenously biotinylated proteins containing
ACACA, MCCC1, PC, and PCCA identified through hierarchical clustering.
A modified two-sided t-test based on permutation-based FDR
statistics^[302]96 was performed between each experimental group and
the control, where significance was determined by an s0 factor of 1 at
5% FDR. To identify candidate interface proteins, we intersected the
resulting lists from the proteomes of full-length LD-BioID, full-length
mito-BioID, and split-BioID. We did not apply additional layers of
filtering to avoid excluding important proteins involved in broader or
complex processes, especially those that are less studied, have
undefined localizations, or undergo dynamic transitions between
cellular compartments.
Affinity purification of the interacting proteins
HepG2 cells stably expressing empty vector (3HA-eGFP, control),
3HA-eGFP-ESYT1, 3HA-eGFP-ESYT2 or 3HA-eGFP-VAPB were grown on 10 cm
dishes. Cells were lysed in 0.75 ml lysis buffer (25 mM HEPES pH 7.4,
150 mM NaCl, 1 mM EDTA, 10% glycerol, 1% wt/vol
n-dodecyl-β-D-maltoside) on ice for 30 min. The lysates were cleared by
centrifuging at 18,000 × g for 15 min at 4 °C. An equal volume of cell
lysates (1∼2 mg of protein) was mixed with 25 µl EZview™ Red Anti-HA
Agarose beads (#E6679; Sigma-Aldrich). The protein bead mixture was
gently rotated at 4 °C overnight, followed by washing three times with
PBS and centrifuging at 16,000 × g for 3 min at 4 °C. For immunoblot
analysis, immunoprecipitated proteins were eluted with 30 µl
2 × Laemmli sample buffer by vortexing at RT for 5 min then denatured
at 95 °C for 5 min. The resultant samples were subjected to SDS-PAGE
and immunoblotting. For proteomic analysis of the immunoprecipitated
proteins, the cysteine bonds were reduced on beads with 5 mM
Tris(2-carboxyethyl)phosphine (TCEP) for 30 mins at 37 °C followed by
alkylation with 10 mM iodoacetamide. Beads were then resuspended in
digestion buffer containing sequencing grade modified trypsin (Pierce)
at 37 °C overnight. After quenching with 10% TFA, the samples were
desalted by C18 reversed-phase spin columns according to the
manufacturer’s instructions (Pierce). The eluted peptide sample was
dried in a vacuum centrifuge and reconstituted to a final volume of
30 μl in 0.1% TFA and 1% CH[3]CN. Analysis was performed on a
Q-Exactive mass spectrometer (Thermo Fisher Scientific) coupled to
liquid chromatography system. The raw files were first searched by
Maxquant and enriched proteins were analyzed by Perseus platform using
the same search parameter as indicated above.
LD and mitochondria staining
Cells were treated with 500 µM oleate/palmitate (2:1) conjugated with
1% BSA for indicated times followed by 10 min fixation with Image-iT™
Fixative Solution (4% formaldehyde, methanol-free) at RT. After three
washes with PBS, cells were permeabilized with 0.1% Triton X-100 for
10 min and blocked for 1 h at room temperature with 2% BSA in PBS. LDs
were stained with HCS LipidTOX Deep Red Neutral Lipid Stain (1:1,000;
Invitrogen) and mitochondria with anti-TOMM20 (Abcam) in PBS (2% BSA)
for 1 h, followed by three washes in PBS at RT with light avoidance.
Slides were mounted in Vectashield antifade aqueous mounting medium
(Vector Laboratories).
Airyscan imaging
Images were acquired on a Zeiss LSM 880 confocal microscope with an
Airyscan detector (Carl Zeiss). Data were collected using either 63x
oil (1.4 NA) or 40x oil (1.3 NA) objective lens for most experiments.
Samples were excited using laser lines of 405, 458, 488, 561, 594 and
633 nm and the emitted signal was detected on Airyscan detector using
‘SuperResolution’ (SR) mode. Where z-stacks were collected, the
software-recommended optimal slice sizes were used. For all
experiments, images were collected sequentially to minimize bleed
through. To maintain clarity and uniformity throughout the paper, some
images have been pseudocolored.
Deconvolution and 3D rendering
The raw Airyscan images containing 32-phase images were deconvolved
with Huygens Professional Software (v22.10, Scientific Volume Imaging)
using Array Detector deconvolution module. The deconvolved images were
imported to Imaris software (v9.9, Oxford Instruments) and the whole
cell, LD and mitochondria were reconstructed to 3D surfaces. To retain
the structure detail, surfaces were created without applying smoothing.
Using the whole cell mask, individual LDs and mitochondria surfaces
were labeled per cell and the statistics were drawn per cell
accordingly. The surface area, number, and volume were exported from
the Statistics tool. In measuring LD-mito contact site, the distance
between LDs and mitochondria was measured as the shortest distance
between the organelle surfaces border and the distance <30 nm were
considered as touching objects.
Fluorescence lifetime imaging microscopy of Förster resonance energy transfer
All live and fixed cell fluorescence lifetime microscopy (FLIM)
measurements of Förster resonance energy transfer (FRET) were performed
on an Olympus FV3000 laser scanning microscope coupled to a 488 nm
pulsed laser operated at 80 MHz and an ISS A320 FastFLIM box. A 60x
water immersion objective (1.2 NA) was used for all experiments, and
the cells were imaged at 37 °C. Prior to acquisition of FLIM data in
the donor channel (eGFP-ESYT1, eGFP-ESYT2, eGFP-VAPB) for FRET analysis
with the acceptor channel (mCherry-VAPB), multi-channel intensity
images were acquired from each selected HepG2 cell to verify that the
FRET acceptor was present in excess of the donor (i.e., acceptor–donor
ratio > 1), and in the case of fixed cell experiments where
immunofluorescence against PLIN2 (AF405) and TOMM20 (AF647) was
performed, to identify the localization of LD-mitochondria contacts
sites for a masked FRET analysis of FLIM data. This involved sequential
imaging of a two-phase light path in the Olympus FluoView software. The
first phase was set up to image eGFP and mCherry via use of solid-state
laser diodes operating at 488 and 561 nm, respectively, with the
resulting signal being directed through a 405/488/561/633 dichroic
mirror to two internal GaAsP photomultiplier detectors set to collect
500–540 nm and 600–700 nm. The second phase was set up to image AF405
and AF647 via use of solid-state laser diodes operating at 405 and
633 nm, respectively, with the resulting signal being directed through
a 405/488/561/633 dichroic mirror to two internal GaAsP photomultiplier
detectors set to collect 420–460 nm and 600–700 nm. Then in each HepG2
cell selected, a FLIM map of eGFP was imaged within the same field of
view (256 × 256-pixel frame size, 20 µs/pixel, 90 nm/pixel, 20 frame
integration) using the ISS VistaVision software. This involved
excitation of eGFP with an external pulsed 488 nm laser (80 MHz) and
the resulting signal being directed through a 405/488/561/633 dichroic
mirror to an external photomultiplier detector (H7422P-40 of Hamamatsu)
that was fitted with a 520/50 nm bandwidth filter. The donor signal in
each pixel was then subsequently processed by the ISS A320 FastFLIM box
data acquisition card to report the fluorescence lifetime of eGFP. All
FLIM data were pre-calibrated against fluorescein at pH 9 which has a
single exponential lifetime of 4.04 ns.
FLIM-FRET analysis
The fluorescence decay recorded in each pixel of an acquired FLIM image
was quantified by the phasor approach to lifetime analysis^[303]97. As
described in previously published papers^[304]98, this results in each
pixel of a FLIM image giving rise to a single point (phasor) in the
phasor plot, which when used in the reciprocal mode enables each point
in the phasor plot to be mapped to each pixel of the FLIM image. Since
phasors follow simple vector algebra, it is possible to determine the
fractional contribution of two or more independent molecular species
coexisting in the same pixel. For example, in the case of two
independent species, all possible weightings give a phasor distribution
along a linear trajectory that joins the phasors of the individual
species in pure form. While in the case of a FRET experiment, where the
lifetime of the donor molecule is changed upon interaction with an
acceptor molecule, the realization of all possible phasors quenched
with different efficiencies describes a curved FRET trajectory in the
phasor plot that follows the classical definition of FRET
efficiency^[305]98.
In the context of the FLIM-FRET experiments presented in this
manuscript, the phasor coordinates (g and s) of the unquenched donor
(eGFP-ESYT1, eGFP-ESYT2, eGFP-VAPB) were first determined independently
in fixed and live HepG2 cells to enable definition of a baseline from
which a FRET trajectory could be extrapolated and then used to
determine the dynamic range of FRET efficiencies that describe FRET
interaction with an acceptor (mCherry-VAPB). All FLIM-FRET
quantification was performed in the SimFCS software developed at the
LFD.
Fluorescence fluctuation spectroscopy (FFS)
All FFS measurements for Number and Brightness (NB) analysis and cross
RICS were performed on an Olympus FV3000 laser scanning microscope
coupled to an ISS A320 Fast FLIM box for fluorescence fluctuation data
acquisition. For single channel NB FFS measurements, eGFP tagged
plasmids were excited by a solid-state laser diode operating at 488 nm
and the resulting fluorescence signal was directed through a
405/488/561 dichroic mirror to an external photomultiplier detector
(H7422P-40 of Hamamatsu) fitted with an eGFP 500/25 nm bandwidth
filter. For dual channel RICS FFS measurements (that enable cross
RICS), the eGFP and mCherry plasmid combination, were excited by
solid-state laser diodes operating at 488 nm and 561 nm, respectively,
and the resulting signal was directed through a 405/488/561/640
dichroic mirror to two internal GaAsP photomultiplier detectors set to
collect 500–540 nm and 620–720 nm, respectively.
All FFS data acquisitions employed a 60X water immersion objective (1.2
NA) and first involved selecting a 10.6 μm region of interest (ROI)
within a HepG2 cell at 37 °C in 5% CO2. Then a single or simultaneous
two channel frame scan acquisition was acquired (N = 100 frames) in the
selected ROI with a pixel frame size of 256 × 256 (i.e., pixel size
~41 nm) and a pixel dwell time of 12.5 µs. These conditions resulted in
a scanning pattern that was found to be optimal for simultaneous
capture of the apparent brightness and mobility of the different eGFP
and mCherry constructs being characterized by NB and cross RICS
analysis; all of which was performed in the SimFCS software developed
at the Laboratory for Fluorescence Dynamics (LFD).
Number and brightness (NB) analysis
The oligomeric state of the different eGFP-tagged plasmids investigated
(i.e., eGFP-ESYT1, eGFP-ESYT2, and eGFP-VAPB) was extracted and
spatially mapped throughout single channel FFS measurements via a
moment-based brightness analysis that has been described in previously
published papers^[306]99. In brief, within each pixel of an NB FFS
measurement there is an intensity fluctuation
[MATH: F(t) :MATH]
which has: (1) an average intensity
[MATH: Ft :MATH]
(first moment) and (2) variance
[MATH: σ2
:MATH]
(second moment); and the ratio of these two properties describes the
apparent brightness (B) of the molecules that give rise to the
intensity fluctuation. The true molecular brightness (ɛ) of the
fluorescent molecules being measured is related to B by
[MATH: B=ε+1 :MATH]
, where 1 is the brightness contribution of a photon counting detector.
Thus, if we measure the B of monomeric eGFP
(B[monomer] = ɛ[monomer] + 1) under our NB FFS measurement conditions,
then we can determine ɛ[monomer] and extrapolate the expected B of
eGFP-tagged dimers (B[dimer] = (2 x ɛ[monomer]) + 1) or oligomers
(e.g., B[tetramer] = (4 x ɛ[monomer]) + 1), and in turn define
brightness cursors, to extract and spatially map the fraction of pixels
within a NB FFS measurement that contain these different species. These
defined brightness cursors were used to extract the fraction of
eGFP-ESYT1, eGFP-ESYT2, and eGFP-VAPB dimer and oligomer (i.e., number
of pixels assigned B[dimer] or B[oligomer]) within a NB FFS measurement
and quantify the degree of protein homodimer and oligomer fractions
across multiple cells. Artifact due to cell movement or photobleaching
were subtracted from acquired intensity fluctuations via use of a
moving average algorithm and all brightness analysis was carried out in
SimFCS from the Laboratory for Fluorescence Dynamics.
RICS and cross RICS analysis
The fraction of interaction between the eGFP tagged ESYT1, ESYT2 and
VAPB with mCherry tagged VAPB was extracted via application of the
cross RICS functions described in previously published paper to the
dual channel FFS measurements^[307]100. In brief, the fluorescence
intensity recorded within each frame (N = 100) of each channel (i.e.,
CH1 and CH2) was spatially correlated via application of the RICS
function, and spatially cross-correlated between channels (CC) via
application of the cross RICS function, alongside a moving average
algorithm (N = 10 frames) in both instances. Then the recovered RICS
and cross RICS correlation profiles were fit to a 3D diffusion model
and the amplitude versus decay of each fit recorded in the form of a G
value. The ratio of the cross RICS amplitude (i.e., G[CC]) with the
limiting channel RICS amplitude (i.e., G[CH1] or G[CH2]) enabled the
fraction of eGFP-ESYT1, eGFP-ESYT2, and eGFP-VAPB molecules interacting
with VAPB-mCherry molecules to be extracted. All cross RICS analysis
was carried out in SimFCS from the Laboratory for Fluorescence
Dynamics.
Radiolabeled fatty acid tracer studies
To accumulate radiolabeled fatty acids in LDs, cells were treated for
indicated times with 1 μCi/ml of [^14C]-oleic acid (NEC317050C,
PerkinElmer, Waltham, Massachussetts) and 500 μM oleic acid
(Sigma#O1006) conjugated to 1% BSA (wt/vol) in low glucose DMEM (1 g
per liter; Gabico#10567-014). Oxidation measurements were performed by
trapping the released ^14CO[2] in 1 M sodium hydroxide in sealed
system. Incompletely oxidized acid-soluble metabolites containing ^14C
and lipids were extracted from the total cell lysate mixed with 3
volumes of chloroform/methanol (2:1). The upper phase containing
incompletely oxidized fatty acids were added to 2 mL scintillation
cocktail and analyzed by liquid scintillation counting while the bottom
organic phase was collected and dried.
Lipids were resuspended in chloroform/methanol (2:1) containing a
standard mix of glyceryl tripalmitate (1.75 mg/ml; Sigma #T588),
dipalmitin (0.875 mg/ml; Sigma #D2636), oleate (1.7 mg/ml; Sigma
#O1006), and C24 ceramide (1 mg/ml, Sigma #43799). The mixture was
spotted on 60-Å silica gel TLC plates (Whatmann, Partisil LK6D) and
dried for several minutes at RT. Chloroform/methanol/water (65:30:5)
solvent was used for initial separation until the solvent front reached
~30% of the plate. Next, after drying, plates were resolved in a
solvent mixture of hexane/diethyl ether/acetic acid (70:30:1) until the
solvent front reached 1 cm from the top of the plate. Lipid spots were
then visualized by staining with 0.02% dichlorofluorescein (Sigma
#D6665) and radioactive bands corresponding to different lipid
standards were scraped and mixed in 2 mL scintillation cocktail and
counted on a scintillation counter.
Fluorescent fatty acid tracer experiments
HeLa cells were seeded in 12 well plates in complete medium (DMEM
Glutmax with 10% fetal bovine serum). Twenty-four hours after seeding,
the cells were pulsed in low-glucose DMEM containing 1 μM BODIPY
558/568 C16 (Thermo Fisher Scientific, D3821). The cells were then
washed three times with DPBS and chased for the indicated times in
low-glucose medium. Fluorescence quantifications were performed at room
temperature with a CLARIOstar plus fluorimeter (BMG LabTech).
Gas chromatography–mass spectrometry stable isotope labeling analysis
Cells were washed with Milli-Q water at 37 °C then snap-frozen by
covering the plate in liquid nitrogen. Metabolites were extracted on
ice by addition of 600 µl/well of methanol:chloroform (9:1 v/v),
containing the internal standard, scyllo-inositol (16.6 µM). Cells were
scraped and incubated on ice for 10 min. Samples were then centrifuged
(5 min, 6000 g, 4 °C) to pellet precipitated proteins and the
supernatants transferred to fresh Eppendorf tubes.
Cell extracts were transferred to vial inserts and evaporated to
dryness under vacuum. Polar metabolites were derivatized online using
an AOC 6000 autosampler robot (Shimadzu). Samples were first
methoxyaminated by the addition of 25 µL methoxyamine (30 mg/mL in
pyridine, 2 h, 37 °C, 750 rpm), followed by trimethylsilylation with
25 µL N, O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA) + 1%
trimethylchlorosilane (TMCS; 1 h, 37 °C, 750 rpm). Following a 1 µl
splitless injection, metabolite profiles were acquired on a 2030
Shimadzu gas chromatograph and a TQ8050NX triple quadrupole mass
spectrometer (Shimadzu, Japan). The mass spectrometer was tuned
according to the manufacturer’s recommendations using
tris-(perfluorobutyl)-amine (CF43). GC-MS was performed on a 30 m
Agilent DB-5 column with 0.25 mm internal diameter column and 1 µm film
thickness. The injection temperature (inlet) was set at 280 °C, the MS
transfer line at 280 °C and the ion source adjusted to 200 °C. Helium
was used as the carrier gas at a flow rate of 1 mL/min and the mass
spectrometer was set to scan mode with a scan range of 50–500 m/z. The
analysis of TMS samples was performed under the following oven
temperature program; 100 °C start temperature, hold for 4 min, followed
by a 10 °C min^−1 oven temperature ramp to 320 °C with a following
final hold for 11 min.
^13C-derived carbon labeling was determined in key metabolites of the
glycolytic and tricarboxylic acid (TCA) cycle via mass isotopomer peak
shift analysis. Data files were imported into DExSI^[308]101 (V3.05),
which enabled quantitation of mass isotopologues for relevant
^13C-labeled metabolites. Fractional abundance data and mass
isotopologue distributions were produced following correction for
natural background isotopic abundance.
Lipidomic analysis
Lipidomic extraction
Briefly, 1 × 10^6 cells were lysed in water/methanol (1:3.5 v/v) with
10 µl internal standard (Splash Lipidomix (Cat no. 330707) Deuterated
Ceramide Lipidomix (Cat no. 330713), Cholesterol-d7 (Cat no. 700041 P),
Avanti Polar Lipids, USA). Then, 1 mL MTBE was added, and the mixture
was shaken for 30 min at room temperature. Phase separation was induced
by adding 0.25 mL of MS-grade water and centrifuged at 1,000 g for
20 min. The upper (organic) phase was collected and dried in a vacuum
centrifuge. Samples were resuspended in 100 µl chloroform/methanol
(1/9, v/v), centrifuged at 14,000 g for 10 min and transferred to glass
LCMS vial.
Ultrahigh-performance liquid chromatography (UHPLC) and mass spectrometric
(MS) analyses
Samples were analyzed by ultrahigh performance liquid chromatography
(UHPLC) coupled to tandem mass spectrometry (MS/MS) employing a
Vanquish UHPLC linked to an Orbitrap Fusion Lumos mass spectrometer
(Thermo Fisher Scientific, San Jose, CA, USA), with separate runs in
positive and negative ion polarities. Solvent A was 6/4 (v/v)
acetonitrile/water with 5 mM medronic acid and solvent B was 9/1 (v/v)
isopropanol/acetonitrile. Both solvents A and B contained 10 mM
ammonium acetate. 10 uL of each sample was injected into an Acquity
UPLC HSS T3 C18 column (1 × 150 mm, 1.8 µm; Waters, Milford, MA, USA)
at 50 °C at a flow rate of 80 μl/min for 3 min using 3% solvent B.
During separation, the percentage of solvent B was increased from 3% to
70% in 5 min and from 70% to 99% in 16 min. Subsequently, the
percentage of solvent B was maintained at 99% for 3 min. Finally, the
percentage of solvent B was decreased to 3% in 0.1 min and maintained
for 3.9 min.
All MS experiments were performed using an electrospray ionization
source in positive mode at 3.5 kV and negative mode at 3.0 kV
separately for each sample. The flow rates of sheath, auxiliary and
sweep gases were 25 and 5 and 0 arbitrary unit(s), respectively. The
ion transfer tube and vaporizer temperatures were maintained at 300 °C
and 150 °C, respectively, and the ion funnel RF level was set at 50%.
In the positive mode from 3 – 24 min, the top speed data-dependent scan
with a cycle time of 1 s was used. Within each cycle, a full-scan
MS-spectra were acquired firstly in the Orbitrap at a mass resolving
power of 120,000 (at m/z 200) across an m/z range of 300–2000 using
quadrupole isolation, an automatic gain control (AGC) target of 4e5 and
a maximum injection time of 50 milliseconds, followed by higher-energy
collisional dissociation (HCD)-MS/MS at a mass resolving power of
15,000 (at m/z 200), a normalized collision energy (NCE) of 27% at
positive mode and 30% at negative mode, an m/z isolation window of 1, a
maximum injection time of 35 milliseconds and an AGC target of 5e4. For
the improved structural characterization of glycerophosphocholine (PC)
lipid cations, a data-dependent product ion (m/z 184.0733)-triggered
collision-induced dissociation (CID)-MS/MS scan was performed in the
cycle using a q-value of 0.25 and a NCE of 30%, with other settings
being the same as that for HCD-MS/MS. For the improved structural
characterization of triacylglycerol (TG) lipid cations, the fatty
acid + NH[3] neutral loss product ions observed by HCD-MS/MS were used
to trigger the acquisition of the top-3 data-dependent ion trap
CID-MS^[309]3 scans in the cycle using a q-value of 0.25 and a NCE of
30%, with other settings being the same as that for HCD-MS/MS.
For identification and quantification of lipids and statistical
analysis, LC-MS/MS data was searched through MS Dial 4.90. The mass
accuracy settings are 0.005 Da and 0.025 Da for MS1 and MS2. The
minimum peak height is 50000 and mass slice width is 0.05 Da. The
identification score cut off is 80%. Post identification was done with
a text file containing name and m/z of each internal standard. In
positive mode, [M + H]+, [M + NH4]+ and [M + H-H2O]+ were selected as
ion forms. In negative mode, [M-H]- and [M + CH3COO]- were selected as
ion forms. All lipid classes available were selected for the search.
PC, LPC, DG, TG, CE, and SM were identified and quantified at positive
mode while Cer and CL were identified and quantified at negative mode.
The retention time tolerance for alignment is 0.1 min. Lipids with
maximum intensity <5-fold of average intensity in blank were removed.
All other settings were default. All lipid LC-MS features were manually
inspected and re-integrated when needed.
Triglyceride assessment
Triglyceride content was measured using an Infinity Triglycerides kit
(Thermo Scientific #TR22421), following the manufacturer’s protocol.
Briefly, lipids were extracted and dried as described above,
resuspended in ethanol, then transferred into 96 well plates containing
200 μL Infinity Triglyceride reagent. A standard curve was generated
using glyceryl trioleate (Sigma#92860) and the absorbance measured at
500 nm using a CLARIOstar plus fluorimeter (BMG LabTech).
In silico modeling and docking analyses
The ESYT1/ESYT2 complex was modeled using the experimental crystal
structure of the ESYT2 dimer (PDB ID: 4P42)^[310]52, based on a
sequence identity of 39% between homologs. As the ESYT2 dimer was
asymmetric, two models were generated: one having ESYT1 replacing
homodimer chain B (model 1) and the other replacing chain A (model 2).
Models were built using Advanced Homology Modeling in Schrodinger
Maestro v.11.4, after preprocessing of experimental structure 4P42
filled in missing atoms and residues in Prime. Both models were used
for further analyses and lipid docking.
Initial fragment hotspot analysis using CCDC^[311]102 on both models
identified an apolar groove within chain B, which was used along with
SiteMap within Schrodinger to guide the search space for docking oleic
acid, linoleic acid, palmitic acid, triacylglycerol,
phosphatidylcholine and cholesteryl oleate. Lipid ligand files were
initially obtained from PubChem, and subjected to ligand preparation
using LigPrep, which assigns rotations within single bonds to permit
the sampling of accurate ligand poses. Docking was carried out on both
models (ESYT2 in model 1, ESYT1 in model 2) using Schrodinger Prime.
Within the lipid binding cavity, 9 residues within ESYT2 (model 1) and
7 residues within ESYT1 (model 2) were tested for stability changes
upon mutation to larger residues, using mCSM-Stability^[312]103.
Residues were chosen as pairs on opposite sides of the lipid channel
entrance, middle and deeper ends, for their potential to block lipid
entry upon mutation, where mutants were determined according to channel
depth at residue position.
The in silico saturation mutagenesis at the protein interface for both
model structures was carried out via mCSM-PPI2^[313]104, which uses a
threshold of 5 Å distance from any other molecule in a 3D space to
define interface residues for any given complex structure. Each
interface residue is then mutated to each one of the other 19 standard
amino acids, and served as input for a machine learning model that
predicts changes in protein binding affinity.
Oxygen consumption measurements
The oxygen consumption rate (OCR) of HeLa cells (Control and KO cell
lines) was measured using a Seahorse XF24 extracellular flux analyzer
(Seahorse Bioscience). HeLa cells were seeded at 30,000 per well in
24-well XF plates the day before assay. Basal medium supplemented with
1 mM pyruvate, 10 mM glucose and 2 mM glutamine was added to each well
of a Seahorse XF24 cell culture plate and incubated overnight at 37 °C.
Cells were equilibrated for 1 h in XF assay medium in a non-CO[2]
incubator. OCR was monitored according to the manufacturer
recommendations by sequential injections of oligomycin (1 μM), carbonyl
cyanide-4-(trifluoromethoxy)phenylhydrazone (1 μM) and
rotenone/antimycin A (0.5 μM). Cells were harvested at the conclusion
of OCR assessment and data normalized to cell number.
Mouse breeding and husbandry
All experimental protocols were approved by the University of Melbourne
Anatomy & Neuroscience, Pathology, Pharmacology, and Physiology Animal
Ethics Committee (Ethics #21228). To generate Alb-Cre:LSL-Cas9-GFP
mice, hemizygous Alb-Cre mice (Strain #:003574) were bred with
homozygous LSL-Cas9-GFP mice (Strain #:028551). In total, 20 male mice
were used for experiments. All mice were housed at 22 °C and maintained
on a 12 h light, 12 h dark cycle in cages with 2-5 mice per cage. Mice
had ad libitum access to a standard chow diet (Specialty Feeds, WA,
Australia) containing 23% energy from protein, 12% from fat, 59 % from
carbohydrate and water or ad libitum access to a high-fat, high-sucrose
diet (High Fat Rodent Diet SF04-001, 43 % energy from fat, Specialty
Feeds, HFD) starting at 8–10 weeks of age for a total of 4 weeks. After
4 weeks of high fat feeding, mice were killed, and hepatocytes were
isolated by collagenase digestion as previously described^[314]105.
AAV design and production
gRNAs for AAV-gScrambled-FLEX-mCherry
(pAAV-U6>mScramble-GTGTAGTTCGACCATTCGTG)-CAG > LL:rev(mCherry):rev(LL):
WPRE, AAV-Esyt-FLEX-mCherry
(pAAV[-U6>mEsyt1[gRNA-TCCCTACGCGCTCGTCCGTGT]-U6>mEsyt1[gRNA-GGGGCACCAAC
AGTCGGTTA]-CAG > LL:rev(mCherry):rev(LL):WPRE) and
AAV-Esty2-FLEX-mCherry
(pAAV[-U6>mEsyt2[gRNA-ACAGCGCCAGCGCGCGGCAC]-U6>mEsyt2[gRNA-GGTCGCAGGCGC
GCACTCCC]-CAG > LL:rev(mCherry):rev(LL):WPRE) viral vectors were cloned
and packaged into the AAV-DJ steotype at a titre of > 2 × 10^13 GC/ml
(VectorBuilder, Chicago, IL). All AAV’s were injected intravenously at
a concentration of 1 × 10^12 genome copies per mouse in 100 µl.
Fatty acid metabolism in primary hepatocytes
Cell density was determined, and hepatocytes were plated at 500,000
cells per well in a 6-well plate (BD Falcon, #35043) and left to adhere
for 4 h in adherence medium containing M199, Penicillin / Streptomycin
(P/S), BSA 10%, dialysed FBS, dexamethasone (100 nM) and insulin
(100 nM). The medium was replaced with low insulin (1 nmol/L) basal
culture media containing M199, P/S and dexamethasone (100 nM) and
incubated overnight at 37 °C. The next day, hepatocytes were incubated
for 2 h with low-glucose DMEM Gluta-MAX (Life Technologies) containing
2% fatty acid-free BSA, 1 μCi/mL oleic acid (NEC317050UC, PerkinElmer)
and 0.5 mmol/L oleic acid (Sigma-Aldrich). Fatty acid oxidation was
calculated as the sum of complete oxidation to ^14CO[2] and incomplete
oxidation (i.e., ^14C-containing acid-soluble metabolites). Fatty acid
uptake was calculated as the sum of fatty acid oxidation and fatty
acids stored in complex lipids (i.e., ^14C in the organic fraction of
the lysed liver slices). Incorporation of ^14C into triglyceride was
measured by thin-layer chromatography as previously described^[315]106.
Identification of ESYT1 and ESYT2 by targeted mass spectrometry
Hepatocytes from Control, ESYT1^KO and ESYT2^KO mice were collected,
snap frozen and stored at -80°C. Identification of ESYT1 and ESYT2 was
performed using targeted mass spectrometry as previously
described^[316]105. The peptide sequences used to detect ESYT1
(LLAETVAPAVR) and ESYT2 (ALALLEDEEQAVR) were unique to each protein.
Drosophila experiments
Breeding and husbandry
To generate fat-body dEsyt knockout flies (dEsyt2^FB-KO), we crossed
LppGal4, UAS-Cas9.P2 (Bloomington Drosophila Stock Center (BDSC) 67078)
females with dEsyt2 gRNA expressing males (BDSC 83802). To generate
whole-body dEsyt knockout flies (dEsyt^KO), we crossed
Df(3 R)Exel7357/TM6B females (chromosomal deletion that includes Esyt2
and hereafter referred to as Df(Esyt2), BDSC 7948) with
Mi(ET1)Esyt2^MB02922/TM6C (a transposable element insertion that is a
protein null allele^[317]75 and hereafter referred to as Esyt^−, BDSC
23501) males. Of the resulting offspring we selected trans-heterozygote
mutant animals. The controls used were wiUAS-Cas9.P2/+; LppGal4/+
heterozygotes (Fat-body control, WT^FB-KO) or Df(3 R)Ecel7357/Tm6C,Sb
(Whole-body control, WT). All flies were raised on standard
molasses-based food at 25 °C.
For gene expression analysis, whole male adult flies (with heads
removed) were snap frozen and stored at -80°C for later analysis. 10
flies were pooled to create each biological replicate. RNA was
extracted with TRI reagent (Sigma-Aldrich) and reverse transcribed with
iScript Reverse Transcriptase (Bio-Rad). Gene expression was determined
by quantitative RT-PCR (BioRad CFX384 Touch™ Real-Time PCR System)
using SYBR Green PCR Master Mix (QIAGEN). Results were analysed using
BioRad CFX Manager™ software (BioRad). Esyt2 gene expression (Forward;
5’TATCTGGTGGGTTACATGGGC, Reverse; 5’CAGGATTACGTCCTTCTCGGA) was
normalized to Actin-5 (Forward; 5’CGAAGAAGTTGCTGCTCTGG, Reverse;
5’AGAACGATACCGGTGGTACG) as the housekeeper gene. mRNA levels were
determined using the 2^−ΔΔCt method.
Starvation assay
For starvation experiments, male flies <36 h of age were transferred
from normal media to vials containing 2 mL of starvation media (1% agar
in water). Flies were kept at 25 °C and transferred to new vials every
48 h. To determine starvation resistance, 10 starvation vials with 10
flies each were monitored, and the number of dead flies were recorded
every 12 h until all flies were dead.
Figure preparation
Figures were prepared using Microsoft PowerPoint (Microsoft 365 MSO).
Images were edited with Image Lab (for Western blot), Image J
(microscopy images). GraphPad Prism 9 (GraphPad Software) was used to
create graphs and calculate statistical significance.
Statistical and reproducibility
Statistical analyses were performed using GraphPad Prism 9. All data
was assessed for normal distribution using the D’Agostino & Pearson or
Shapiro-Wilk test. For normally distributed data, differences between
groups were assessed using two-tailed unpaired t-tests or one-way
analysis of variance (ANOVA) with Bonferroni multiple comparison tests.
For nonparametric data, differences between groups were assessed using
a Kruskal-Wallis test with Dunn’s multiple comparison tests. Starvation
resistance was analysed using a log-rank (Mantel Cox) test.
Survivorship curves show the percentage of flies remaining alive as a
function of time in hours. Differences were considered significant at
p < 0.05. The number of technical replicates and/or biological
replicates are described in each figure legend. No statistical method
was used to predetermine sample size and no data were excluded from the
analyses. The experiments were not randomized and the investigators
were in most cases not blinded to allocation during experiments and
outcome assessment.
Materials availability
All unique/stable reagents generated in this study are available from
the lead contact with a completed materials transfer agreement.
Reporting summary
Further information on research design is available in the [318]Nature
Portfolio Reporting Summary linked to this article.
Supplementary information
[319]Supplementary Information^ (3.9MB, pdf)
[320]41467_2025_57405_MOESM2_ESM.docx^ (12.7KB, docx)
Description of Additional Supplementary Information
[321]Supplementary Data 1^ (442.5KB, xlsx)
[322]Supplementary Data 2^ (3.7MB, xlsx)
[323]Supplementary Data 3^ (69.4KB, xlsx)
[324]Reporting Summary^ (84.9KB, pdf)
[325]Transparent Peer Review file^ (381.8KB, pdf)
Source data
[326]Source Data^ (6.2MB, xlsx)
Acknowledgements