The Janus-Faced Role of Lipid Droplets in Aging: Insights from the Cellular Perspective

Abstract

It is widely accepted that nine hallmarks—including mitochondrial dysfunction, epigenetic alterations, and loss of proteostasis—exist that describe the cellular aging process. Adding to this, a well-described cell organelle in the metabolic context, namely, lipid droplets, also accumulates with increasing age, which can be regarded as a further aging-associated process. Independently of their essential role as fat stores, lipid droplets are also able to control cell integrity by mitigating lipotoxic and proteotoxic insults. As we will show in this review, numerous longevity interventions (such as mTOR inhibition) also lead to strong accumulation of lipid droplets in Saccharomyces cerevisiae, Caenorhabditis elegans, Drosophila melanogaster, and mammalian cells, just to name a few examples. In mammals, due to the variety of different cell types and tissues, the role of lipid droplets during the aging process is much more complex. Using selected diseases associated with aging, such as Alzheimer’s disease, Parkinson’s disease, type II diabetes, and cardiovascular disease, we show that lipid droplets are “Janus”-faced. In an early phase of the disease, lipid droplets mitigate the toxicity of lipid peroxidation and protein aggregates, but in a later phase of the disease, a strong accumulation of lipid droplets can cause problems for cells and tissues.

Full text

Citation: Bresgen, N.; Kovacs, M.;
Lahnsteiner, A.; Felder, T.K.;
Rinnerthaler, M. The Janus-Faced
Role of Lipid Droplets in Aging:
Insights from the Cellular
Perspective. Biomolecules 2023,13,
912. https://doi.org/10.3390/
biom13060912
Academic Editor: Gabriella D’Orazi
Received: 13 March 2023
Revised: 22 May 2023
Accepted: 29 May 2023
Published: 30 May 2023
Copyright: © 2023 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
biomolecules
Review
The Janus-Faced Role of Lipid Droplets in Aging: Insights from
the Cellular Perspective
Nikolaus Bresgen 1, Melanie Kovacs 1, Angelika Lahnsteiner 1, Thomas Klaus Felder 2, *
and Mark Rinnerthaler 1, *
1Department of Biosciences and Medical Biology, Paris-Lodron University Salzburg, 5020 Salzburg, Austria;
nikolaus.bresgen@plus.ac.at (N.B.)
2Department of Laboratory Medicine, Paracelsus Medical University, 5020 Salzburg, Austria
*Correspondence: t.felder@salk.at (T.K.F.); mark.rinnerthaler@plus.ac.at (M.R.)
Abstract:
It is widely accepted that nine hallmarks—including mitochondrial dysfunction, epigenetic
alterations, and loss of proteostasis—exist that describe the cellular aging process. Adding to this, a
well-described cell organelle in the metabolic context, namely, lipid droplets, also accumulates with
increasing age, which can be regarded as a further aging-associated process. Independently of their
essential role as fat stores, lipid droplets are also able to control cell integrity by mitigating lipotoxic
and proteotoxic insults. As we will show in this review, numerous longevity interventions (such
as mTOR inhibition) also lead to strong accumulation of lipid droplets in Saccharomyces cerevisiae,
Caenorhabditis elegans,Drosophila melanogaster, and mammalian cells, just to name a few examples.
In mammals, due to the variety of different cell types and tissues, the role of lipid droplets during
the aging process is much more complex. Using selected diseases associated with aging, such as
Alzheimer’s disease, Parkinson’s disease, type II diabetes, and cardiovascular disease, we show that
lipid droplets are “Janus”-faced. In an early phase of the disease, lipid droplets mitigate the toxicity
of lipid peroxidation and protein aggregates, but in a later phase of the disease, a strong accumulation
of lipid droplets can cause problems for cells and tissues.
Keywords:
LDs; autophagy; mitochondria; protein aggregates; lipid peroxides; misfolded proteins;
mTOR; IIS; lifespan; aging
1. Introduction
Lipid droplets (LDs) are evolutionary conserved structures that were mentioned for the
first time by Van Leeuwenhoek in 1674, but their reassessment as autonomous organelles
with important key roles in lipid and energy metabolism occurred many years later [
1
,
2
].
LDs originate from the endoplasmic reticulum (ER). In the first step, neutral lipids are
synthesized at the ER and are redirected into the bilayer, leading to an aggregation of
the highly motile lipids. Morphologically, the accumulation of neutral lipids in the ER
bilayer resembles a lens-like structure. Growth in this lens initiates bilayer deformation
and the budding-off of LDs to the cytoplasm [
3
]. Due to this special mode of formation,
LDs are surrounded by a lipid monolayer and are filled with neutral lipids, especially
triacyclglycerols (TAGs) and sterols. Therefore, LDs are mainly considered fat-storage
organelles with high relevance to lipid-metabolism homeostasis. However, in recent years,
evidence has accumulated that LDs are also capable of mediating cytoprotective properties
by either acting as a “buffer” for toxic lipids [
4
–
6
] or serving the cellular clearance of
damaged and misfolded proteins [
7
–
11
] (Figure 1). There is also growing evidence that
LDs are involved in the binding and detoxification of xenobiotics; however, this will not be
discussed in detail within this review, which focuses on the aspect of aging. By studying
protein composition in LDs in different organisms such as bacteria, plants, insects, yeast,
and mammals, hundreds of different LD-surface-associated proteins have been identified.
Biomolecules 2023,13, 912. https://doi.org/10.3390/biom13060912 https://www.mdpi.com/journal/biomolecules

Biomolecules 2023,13, 912 2 of 56
Although the LD proteome shows qualitative and quantitative variations among different
cell types, a typical mammalian LD contains 100–150 different proteins [
12
]. Surface
proteins are important for regulating LD homeostasis and enable the specific contract
with other cell organelles. Major LD-associated proteins in mammals belong to the PAT
protein family, also known as perilipin 1–5 (PLIN1-5) [
13
,
14
], adipocyte differentiation-
related protein (ADRP) [
15
], and tail-interacting protein of 47 kDa (TIP47) [
14
,
15
]. Several
different LD-resident proteins contributing to lipid biogenesis and degradation, as well as
membrane trafficking and signaling, have been well reviewed [
16
]. Furthermore, it is well
established that LDs form defined contacts with several other cellular organelles such as
the ER, peroxisomes, lysosomes, and mitochondria (reviewed in [
17
]). Intriguingly, LDs
may also sequester proteins involved in genetic control such as histones [18].
Biomolecules 2023, 13, 912 2 of 58
proteins have been identified. Although the LD proteome shows qualitative and
quantitative variations among different cell types, a typical mammalian LD contains 100–
150 different proteins [12]. Surface proteins are important for regulating LD homeostasis
and enable the specific contract with other cell organelles. Major LD-associated proteins
in mammals belong to the PAT protein family, also known as perilipin 1–5 (PLIN1-5)
[13,14], adipocyte differentiation-related protein (ADRP) [15], and tail-interacting protein
of 47 kDa (TIP47) [14,15]. Several different LD-resident proteins contributing to lipid
biogenesis and degradation, as well as membrane trafficking and signaling, have been
well reviewed [16]. Furthermore, it is well established that LDs form defined contacts with
several other cellular organelles such as the ER, peroxisomes, lysosomes, and
mitochondria (reviewed in [17]). Intriguingly, LDs may also sequester proteins involved
in genetic control such as histones [18].
Figure 1. LDs as a cellular “buffer organelle”. LDs serve as an intermediate cytosolic lipid buffer
and assist the cell in detoxifying lipids, misfolded proteins, and protein aggregates present in the
cytosol, ER, and mitochondria. Furthermore, LDs are also involved in adaption to cellular stress by
modulating transcriptional control.
Here, we review the existing evidence for a distinct role of LDs in eukaryotic aging
as explicitly reflected by the accumulation of LDs at terminal life periods [19–21]. Focusing
on the “physiologic triad”—metabolic regulation, stress response, and aging—but also
covering the evolutionary context, we decided to provide an up-to date, detailed review
of a multitude of aging-related aspects of LD biology investigated in the classical
biological model system (Saccharomyces cerevisiae, Caenorhabditis elegans, and Drosophila
melanogaster) and also associated with age-related human disease. This aempt, by its
nature, is complex, and discussing the fascinating, multifaceted role of lipid droplets in
the multiple contexts of aging deserves an extended approach. As will be outlined, there
is evidence for a Janus-faced role of LDs, their cellular accumulation counteracting stress-
associated, disease-provoking forces. Thus, this is beneficial to aging, but conversely
accelerates disease progression at advanced stages, which promotes the aging process. As
we show in the course of this review, there is a close interplay between cellular pathways
that regulate aging processes on the one hand, and on the other hand also affect the
biogenesis of LDs and run like a thread through evolution (see Figure 2).
Figure 1.
LDs as a cellular “buffer organelle”. LDs serve as an intermediate cytosolic lipid buffer
and assist the cell in detoxifying lipids, misfolded proteins, and protein aggregates present in the
cytosol, ER, and mitochondria. Furthermore, LDs are also involved in adaption to cellular stress by
modulating transcriptional control.
Here, we review the existing evidence for a distinct role of LDs in eukaryotic aging as
explicitly reflected by the accumulation of LDs at terminal life periods [
19
–
21
]. Focusing
on the “physiologic triad”—metabolic regulation, stress response, and aging—but also
covering the evolutionary context, we decided to provide an up-to date, detailed review of
a multitude of aging-related aspects of LD biology investigated in the classical biological
model system (Saccharomyces cerevisiae,Caenorhabditis elegans, and Drosophila melanogaster)
and also associated with age-related human disease. This attempt, by its nature, is complex,
and discussing the fascinating, multifaceted role of lipid droplets in the multiple contexts
of aging deserves an extended approach. As will be outlined, there is evidence for a
Janus-faced role of LDs, their cellular accumulation counteracting stress-associated, disease-
provoking forces. Thus, this is beneficial to aging, but conversely accelerates disease
progression at advanced stages, which promotes the aging process. As we show in the
course of this review, there is a close interplay between cellular pathways that regulate

Biomolecules 2023,13, 912 3 of 56
aging processes on the one hand, and on the other hand also affect the biogenesis of LDs
and run like a thread through evolution (see Figure 2).
Biomolecules 2023, 13, 912 3 of 58
Figure 2. The longevity–LD interaction network. In most model organisms relevant for human aging
research, inhibition of (1) TGF-β signaling, (2) mTOR signaling, (3) insulin/IGF-1 signaling and
caloric restriction (CR) promotes both longevity and LD formation. Doed lines indicate the absence
of the respective pathways in yeast cells.
2. Lipid Droplets in Saccharomyces cerevisiae
The baker’s yeast Saccharomyces cerevisiae is a valuable tool for aging research, as
many aging- and disease-associated pathways such as DNA repair mechanisms,
lipostasis, proteostasis, oxidative stress responses, regulated cell death, nutrient signaling,
autophagy, and regulation of the cell cycle are evolutionarily conserved to a high degree
[22]. Based on sequence similarity, about 30% of the yeast genome is conserved in the
human genome [23]. Important for aging research is the fact that when adequate and
sufficient nutrients are provided, S. cerevisiae cells grow exponentially via asymmetric
budding of daughter cells from bigger mother cells [24].
2.1. Replicative and Chronologic Lifespan
In general, in yeast cells, two forms of aging mechanisms can be distinguished,
namely, replicative and chronological aging. For both, cell death terminates the lifespan,
caused by the intrinsic, mitochondrial outer membrane permeabilization (MOMP)-based
activation of programmed cell death (PCD)/apoptosis emerging from increased reactive
oxygen species (ROS) production and genomic instability, which provokes damage to the
cellular proteome, lipidome, and organelles such as mitochondria [25]. Upon nutritional
stress (i.e., exhaustion of nutrients), yeast cells stop dividing and enter a stationary phase
which allows survival up to several weeks depending on strain type and culture
conditions [22]. This survival period in the stationary phase is termed the chronological
lifespan [26] and has to be distinguished from the yeast replicative lifespan, which is
measured by the number of daughter cells that can be formed from a mother cell before it
stops dividing [27]. The average lifespan in the yeast background BY4741 (the most used
genetic background, generally considered as the wild type) lasts 25 generations. Aged
(mother) cells are larger, and reveal a slowing down of the cell cycle and a declined protein
synthesis. Each daughter cell that is formed leaves a bud scar on the mother cell surface
that can be observed microscopically by calcofluor-white staining [28]. It is believed that
Figure 2.
The longevity–LD interaction network. In most model organisms relevant for human aging
research, inhibition of (1) TGF-
β
signaling, (2) mTOR signaling, (3) insulin/IGF-1 signaling and
caloric restriction (CR) promotes both longevity and LD formation. Dotted lines indicate the absence
of the respective pathways in yeast cells.
2. Lipid Droplets in Saccharomyces cerevisiae
The baker’s yeast Saccharomyces cerevisiae is a valuable tool for aging research, as
many aging- and disease-associated pathways such as DNA repair mechanisms, lipostasis,
proteostasis, oxidative stress responses, regulated cell death, nutrient signaling, autophagy,
and regulation of the cell cycle are evolutionarily conserved to a high degree [
22
]. Based on
sequence similarity, about 30% of the yeast genome is conserved in the human genome [
23
].
Important for aging research is the fact that when adequate and sufficient nutrients are
provided, S. cerevisiae cells grow exponentially via asymmetric budding of daughter cells
from bigger mother cells [24].
2.1. Replicative and Chronologic Lifespan
In general, in yeast cells, two forms of aging mechanisms can be distinguished, namely,
replicative and chronological aging. For both, cell death terminates the lifespan, caused by
the intrinsic, mitochondrial outer membrane permeabilization (MOMP)-based activation
of programmed cell death (PCD)/apoptosis emerging from increased reactive oxygen
species (ROS) production and genomic instability, which provokes damage to the cellular
proteome, lipidome, and organelles such as mitochondria [
25
]. Upon nutritional stress
(i.e., exhaustion of nutrients), yeast cells stop dividing and enter a stationary phase which
allows survival up to several weeks depending on strain type and culture conditions [
22
].
This survival period in the stationary phase is termed the chronological lifespan [
26
] and
has to be distinguished from the yeast replicative lifespan, which is measured by the
number of daughter cells that can be formed from a mother cell before it stops dividing [
27
].

Biomolecules 2023,13, 912 4 of 56
The average lifespan in the yeast background BY4741 (the most used genetic background,
generally considered as the wild type) lasts 25 generations. Aged (mother) cells are larger,
and reveal a slowing down of the cell cycle and a declined protein synthesis. Each daughter
cell that is formed leaves a bud scar on the mother cell surface that can be observed
microscopically by calcofluor-white staining [
28
]. It is believed that damaged proteins
and organelles (e.g., mitochondria) are specifically retained by the mother cells, which
explains the “rejuvenation” of daughter cells resulting from asymmetric segregation [
25
,
29
].
Representing a mitosis-based lifespan definition, replicative aging in yeast cells mimics the
limited mitotic capacity of non-transformed proliferating mammalian cells types, including
undifferentiated stem cells as defined first by the Hayflick limit [
30
]. On the other hand, many
phenotypical characteristic described for the chronological aging of yeast cells residing
in the stationary phase share similarity with the phenotype of aged, post-mitotic cells in
higher eukaryotes mainly comprising the class of terminally differentiated cell types such
as cells of the central nervous system [31].
In some respects, several properties of S. cerevisiae render this fungal cell system a
preferred aging model advantageous to human
in vitro
cell culture models. For instance,
large numbers of cells can be monitored in comparatively short time periods under
in vivo
conditions in yeast. Of note, contrasting the well-conserved intracellular aging mechanisms
common to both, yeast cells fail to display intercellular effects seen in multicellular organ-
isms, such as inflammatory or systemic responses (e.g., regulated by hormones and/or the
immune system), as well as other mechanisms involved in cell–cell communication [
22
].
However, it should not be overseen that,
in vitro
cell cultures, as for instance derived from
mammalian tissues, are devoid of systemic, physiologic “cross-talks and feedback loops” if
used as primary cell lines, and co-culturing with other cell types will reflect only part of the
systemic complexity directing individual cell fate
in vivo
, in particular under the aspect of
aging. Moreover, interpretation of experimental findings based on immortalized eukaryotic
cell lines, self-evidently, is complicated due to the fundamentally altered growth control.
Both replicative and chronological lifespan in yeast can be extended by caloric re-
striction, which can be obtained by lowering glucose availability in the culture media
(e.g., from 2% to 0.5%) [
32
]. In the absence of caloric restriction, chronologically aged yeast
cells accumulate ethanol produced by glucose fermentation [
32
]. It is speculated that this
counteracts the expression of
β
-oxidation regulatory enzymes Fox1p, Fox2p, and Fox3p
(peroxisomal fatty acid
β
-oxidation core enzymes) leading to a decline in peroxisomal
oxidation of LD-derived non-esterified “free” fatty acids that are synthesized in the ER
and are stored in LDs [
33
,
34
]. In turn, non-oxidized free fatty acids will accumulate in
LDs under normal nutritional conditions (i.e., 2% glucose) which promotes an inhibitory
feedback loop on the ER-based synthesis of triacyclglycerols (TAG) [
33
]. It is hypothesized
that lipid dynamic remodeling of this kind can shorten lifespan in chronologically aged
yeast cells grown without caloric restriction (i.e., in the presence of 2% glucose) by three
different mechanisms: (i) via necrotic cell death ensuing from the peroxisomal failure to
oxidize free fatty acids, (ii) apoptosis stimulated by the accumulation of diacylglycerol and
free fatty acids in the ER (“lipoapoptosis”), or (iii) diacylglycerol initiated protein kinase
C-dependent signaling [33].
This accounts for a pivotal role of lipid dynamics in yeast aging, which is further
supported by the finding that LD biogenesis in yeast is elevated in the course of replicative
and chronological aging as well as under stress conditions [
19
,
35
,
36
]. Of special rele-
vance, Beas et al. reported that overexpression of the BNA2 gene encoding indoleamine
2,3-dioxygenase (BNA2 is the yeast homolog of mammalian IDO1) leads to a 40% reduction
in LD accumulation during replicative aging, which identifies BNA2 as an important regu-
lator of LD abundance [
22
]. Bna2p catalyzes the first step of NAD
+
synthesis converting
tryptophan to formyl-kynurenine; hence, this finding reveals a connection between the
NAD
+
/kynurenine pathway and LD formation in the course of aging. It is proposed that
the glycolytic flux in aging yeast cells is directed towards neutral lipid synthesis and LD gen-
eration, but Bna2p overexpression diverts the glycolytic flux from pyruvate and acetyl-CoA

Biomolecules 2023,13, 912 5 of 56
to the shikimate pathway (responsible for the synthesis of the amino acids phenylalanine,
tyrosine, and tryptophan) and as a result lowers LD accumulation in the aged cells. Impor-
tantly, this investigation reveals that this kind of Bna2p-mediated “metabolic rewiring” in
aged yeast cells is not directly associated with longevity. Moreover, the findings indicate
that LD accumulation does not cause lifespan shortening, but, conversely, exerts protection
of aged cells under stress conditions, which might provide a selective growth advantage
under variable environmental conditions [36].
2.2. Lipid Droplets and Stress Adaptation
This concept is supported by another study that substantiates the role of LDs as key
players in cellular stress adaption. The yeast cell growth rate declines when phosphatidyl-
choline biosynthesis is deficient, which changes the cellular phospholipid content and
causes ER stress, alterations in ER morphology, and enhanced LD formation. In this case,
an excess of phospholipids is converted to TAG by the acyltransferases Lro1p and Dga1p,
which is immediately sequestered by LDs. This LD-generating process allows yeast cells
to rebalance the pool of freely available phospholipids as an indispensable prerequisite
for organelle morphology retrieval and cell growth [
9
]. Besides this pathway of ER-based
regulation of lipid homeostasis yielding LD formation in yeast, ER stress arising from
lipid imbalance is also at risk of activating the unfolded protein response (UPR). In most
model organisms, it is shown that the UPR protects cells from the detrimental effects of
proteotoxicity and is of great importance for the aging process [
37
]. Therefore, it is not
surprising that all interventions that increase the activity of the UPR clearly extend the
replicative lifespan of yeast cells [38].
The UPR provides cellular maintenance by specific handling of accumulated misfolded
protein as well as facing lipid bilayer stress in the ER. Besides ER expansion, UPR signaling
comprises the activity of a number of UPR-related gene products which direct the response
either towards re-established homeostasis or, if not adequately facing a prolonged stress
condition, participate in apoptosis onset (for a review see [
39
]). Essential to a successful
outcome is the proper elimination of the ER stressor. A misfolded protein that initially
accumulates inside the ER is translocated to the cytosol, where it is polyubiquitinylated by
ubiquitin-conjugating enzymes residing at the cytosolic ER surface, the polyubiquitination
serving as tag for proteasomal degradation [
40
]. However, lipid bilayer stress may also
stimulate UPR in the ER (UPR
ER
) [
41
,
42
] which converges with the UPR triggered by
the misfolded protein at the central UPR effector Ire1p (inositol-requiring enzyme 1) [
43
].
Interestingly, in mouse hepatocytes, ER stress stimulates Ire-1 and downstream targets
such as DGAT2 (diacylglycerol-acyltransferase 2) [
44
], with DGAT2 (as well as DGAT1)
being essential to LD biosynthesis [
45
]. Referring to this and findings demonstrating ROS-
triggered LD biogenesis and antioxidant properties of LDs in Drosophila [
46
], Walther et al.
suggested that the Ire1p/DGAT2-stimulated LD formation could antagonize phospholipid
oxidation via LD-mediated ROS scavenging [
47
]. This also underlines the importance of
LDs for the aging process as the accumulation of ROS is one of the most prominent features
at the terminal lifespan [48].
Moreover, linking LD formation to UPR-dependent responses in yeast, it was shown
that ER-derived LDs can be associated with polyubiquitinylated proteins and also can
be enriched in Kar2p, an ER chaperone involved in protein folding [
9
]. This led to the
conclusion that un-/misfolded proteins accumulating in the ER are cleared from this
compartment via LD formation, the released LDs being degraded terminally in the yeast
vacuole by a process resembling microautophagy, termed microlipophagy. It has to be
emphasized that this process differs from starvation-induced macroautophagy, since it
does not involve the ATG-dependent initiation of (macro)autophagosomes, but instead
requires ESCRT components (endosomal sorting complexes required for transport) and
the ER-stress response factor Esm1 (ER stress-induced microlipophagy protein 1) [
9
,
10
].
Both stimulation of autophagy and ESCRT components extend the chronological lifespan
of yeast cells [
49
]. A further study also clearly links LDs with the removal of aggregates

Biomolecules 2023,13, 912 6 of 56
consisting of misfolded proteins. Moldavski et al. showed that so-called inclusion bodies
(IBs) are functionally and spatially linked to LDs [
8
]. Upon stress induction, unfolded or
misfolded proteins, which cannot be cleared by the quality control machinery (e.g., due
to quality control system overload or failure) aggregate and form inclusion bodies. In
an extensive screening approach, Moldavski and co-workers identified thirteen proteins
that are crucial for an efficient and rapid IB clearance. Interestingly one of these proteins,
namely, Iml2p, strongly associates with LDs via interaction with the LD-resident proteins
Pet10p and Pdr16p. It should be noted that Pet10p is the yeast perilipin, which is the
only perilipin discovered so far in S. cerevisiae [
50
]. This interaction especially happens
during cell stress, when Iml2 is exclusively located in inclusion bodies. Under such stress
conditions, a physical tethering between LDs and IBs can be monitored, the physical
binding of LDs to IBs allowing aggregate clearance. Iml2 is essential to this clearance
process, which is considered to be mediated by a soluble sterol derivate effusing from LDs
via interaction with Iml2 [
8
]. These findings highlight the role of LD-dependent protein
aggregate clearance during aging, which is still poorly studied considering the substantial
influence of cellular aging on both protein misfolding and protein toxicity [
51
]. Besides
Pet10p and Pdr16p, another LD-resident protein, Ubx2p, could be involved in protein
homeostasis [
52
,
53
]. This UBX-domain-containing protein resides in the ER but relocates to
LDs upon their formation. UBX2 deletion leads to abnormal cellular numbers of LDs of
reduced size and TAG content [
54
]. At the same time, this protein is also involved in protein
homeostasis, in that Ubx2p recruits Cdc48p and both interact to support ER-associated
protein degradation [55].
2.3. Lipid Droplets: Guardians of Mitochondrial Integrity
In line with these findings, our research also indicates a linkage between LD formation
and the removal of un-/misfolded, potentially harmful proteins in yeast and mammalian cells.
Moreover, we demonstrated that several, proteins including yeast Mmi1p and Erg6p, as well
as mammalian BAX, BCL-X
L
, and TCTP, can be transferred from mitochondria to LDs via a
V-shaped domain consisting of two alpha helices [
35
]. The V-domain shows a higher binding
affinity to the LD membrane than to the outer membrane of mitochondria, which explains
the directed transfer [
7
,
35
]. Among different possible contexts, this directed protein shuttling
is of special relevance to the control of PCD/apoptosis onset mediated by the pro-apoptotic
bcl-2 family members BAX and BAK. It has to be clearly stated that apoptosis and aging are
deeply interconnected in yeast as well as in mammalian cells [
56
–
58
], and LDs seem to be
involved in both processes. In most cells, apoptosis is increased with the dysregulation of
the apoptotic program, enhancing the risk of cancer and cellular senescence [
58
]. Induced
by a plethora of potential intrinsic cell death stimuli, BAK and BAX translocate to the
mitochondrial outer membrane where they form the mitochondrial-apoptosis-induced
channel (MAC), resulting in MOMP. As a consequence, the release of cytochrome C from
the mitochondrial intermembrane space to the cytosol promotes apoptosome formation,
caspase 9 activation, and the terminal progression of intrinsic apoptotic signaling [
59
,
60
].
Particularly under cellular stress conditions, the anti-apoptotic mammalian bcl-2 family
member BCL-X
L
, as well as TCTP, also translocate to mitochondria but suppress MOMP by
antagonizing BAX/BAK oligomerization [
61
]. In a similar way, Mmi1p, the yeast homolog
of TCTP, also participates in the apoptotic machinery, with the deletion of Mmi1p leading to
an extended replicative lifespan [
62
,
63
]. From this, it can be speculated that under a given
stress condition both pro-and anti-apoptotic proteins locate to the outer mitochondrial
membrane, continuously challenging MOMP onset. Such potentially harmful mitochondria
may be specifically removed by mitophagy, a selective mode of macroautophagy [64].
Emphasizing its specificity for mitochondria, mitophagy in yeast depends on the
activity of Uth1p which localizes to the outer mitochondrial membrane and is required
for mitophagy, but not for starvation-induced bulk macroautophagy [
65
]. As previously
stated, mitophagy is crucial to cellular maintenance under stress conditions by elimi-
nating dysfunctional mitochondria, which is complicated by the fact that stress-induced

Biomolecules 2023,13, 912 7 of 56
macroautophagy/mitophagy may confer cell protection in one stress context, but con-
versely can contribute to cell death (i.e., autophagic cell death) under different stress
conditions [
66
,
67
]. Besides BAX/BAK-mediated MAC, excessive ROS generation can lead
to the formation of another mitochondrial permeability transition pore (mPT). The mPT
pore complex is composed of VDAC (voltage-dependent anion channel) in the outer mem-
brane, cyclophilin D in the matrix, and ANT (adenine-nucleotide translocator) in the inner
membrane, and opening of the mPT, leading to mitochondrial swelling in many cases
followed by necrotic cell death [
59
]. However, mPT opening may also initiate BAX/BAK-
mediated MAC/MOMP; to a large degree, the outcome of this depends on cellular ATP
availability comprising cell death by either necrosis or apoptosis, which also may involve
enhanced autophagy/mitophagy [
68
]. Reminiscent of this, for yeast mutants lacking
Mdm38p, a K
+
/H
+
exchange-regulator residing in the inner mitochondrial membrane has
been reported, which develops a drop of the mitochondrial membrane potential that is
accompanied by mitochondrial swelling, deterioration in mitochondrial morphology, and
vacuolar changes indicative of mitophagy [69].
LDs and mitochondrial homeostasis. It has to be emphasized that mitophagy does
not necessarily need to be associated with conditions of enhanced stress, but represents
an important physiological regulator of mitochondrial homeostasis. In postmitotic mam-
malian cells, mitophagy is crucial to the control of mitochondria numbers under normal
physiologic conditions, as well as the removal of dysfunctional mitochondria in starving
cells [
70
]. In this context, the age-dependent decline in autophagic activity seen in mam-
malian cells [
71
] deserves particular attention since it may weaken the cellular clearance
from dysfunctional mitochondria. Hence, it is conceivable that additional mechanisms may
support cellular maintenance in aged cells by protecting them from the onset of prema-
ture cell death via apoptosis caused by “stressed” mitochondria. The above-mentioned
V-domain-based shuttling of Mmi1p, BAX, and other MOMP agonists to LDs could fulfill
this task considering that LDs closely locating to mitochondria are capable of sequestering
pro-apoptotic proteins, and as a result antagonize the onset of MOMP-dependent apopto-
sis [
35
]. Terminally, such potentially harmful BAX-enriched LDs will be degraded in the
yeast vacuole. Indeed, in yeast cells, we demonstrated the V-domain/LD based protection
from apoptosis, but, conversely, human HepG2 hepatoma cells treated with the apoptosis
inducer staurosporine revealed a substantially elevated susceptibility for apoptosis upon
the V-domain-mediated translocation of BAX and Bcl-X
L
from mitochondria to LDs [
35
].
Explaining this, we observed the translocation of pro-apoptotic Bcl-X
S
to the mitochondria
in staurosporine-treated HepG2 cells. Opposing anti-apoptotic Bcl-X
L
(i.e., the long iso-
form), Bcl-X
S
(the short isoform) is a pro-apoptotic splice variant of Bcl-X, the Bcl-X
L
/Bcl-X
S
ratio being defined by the cell type and cell differentiation, which are dependent (e.g., non-
transformed versus tumor cells) by numerous determinants including transcription factors
and cytokine signaling [
72
]. Importantly, we found Bcl-X
S
to be devoid of a V-domain [
23
],
which may explain the enhanced onset of apoptosis in staurosporine-treated HepG2 cells.
Taken together, this emphasizes the dependence of V-domain/LD-based MOMP inhibi-
tion on additional regulatory elements, in particular in mammalian cells, rendering the
mechanisms cell-type-specific. Ongoing research demonstrates that the V-domain-based
mitochondria to LD shuttling is not restricted to the MOMP/apoptotic settings as presented
above, but seem to play a more general role in cellular stress responses, as indicated by the
marked protein accumulation by LDs seen during replicative aging and in the initiation of
proteotoxic stress [
7
]. In good correspondence with this, Garcia et al. reported a substantial
remodeling of the LD proteome in the presence of ER stress [10].
LDs and DNA repair. Moreover, certain yeast haploid rad
∆
(radiation damage) dele-
tion strains also show altered lipid storage patterns and a reduced lifespan [
73
]. RAD genes
are involved in DNA repair (e.g., nucleotide/base excision repair) which is evolutionary
highly conserved. In yeast, repair of double-strand breaks via homologous recombina-
tion is accomplished by the MRX complex composed of the RAD gene products Mre11p,
Rad50p, and Xrs2p [
73
]. Deletion of one of these three genes leads to higher levels of TAGs

Biomolecules 2023,13, 912 8 of 56
and steryl esters, as well as characteristic changes in lipid-metabolism-associated gene
expression. The down-regulated expression of lipolysis-associated genes (e.g., TGL3) at
an augmented expression of genes involved in lipid synthesis (LPP1,SLC1), together with
high TAG levels, may readily explain the observed increase in LD numbers in rad
∆
mutants.
This is accompanied by chronological lifespan shortening and pronounced mitochondrial
fragmentation indicative of premature aging. However, as normally aged cells also dis-
played higher LD numbers, it is not clear whether the increased LD abundance simply
reflects the premature aging process of rad
∆
mutants or, vice versa, LD accumulation is
causal to chronological lifespan shortening [
73
]. Concerning the considerations made
above regarding a cytoprotective role of LD accumulation in stress adaptation, it would
be interesting to study the extent to which the severity of the phenotype is altered in rad∆
mutants devoid of LDs.
These findings account for a functional triad between LD abundance, mitochondrial
integrity, and lifespan in yeast, which is addressed by stress conditions as well as the
general aging process. Following the common view of mitochondrial dysfunction as a
hallmark of aging [
74
], the causal relationship between mitochondrial fragmentation and
chronological lifespan shortening, as seen in yeast exposed to high glucose levels [
75
],
represents a reliable means of monitoring the aging process already at early stages [
29
].
Extending this, and in line with the functional triad envisaged above, LD accumulation
in the same way may be considered a complementary biomarker for both premature and
normal aging, as suggested by Kanagavijayan et al. [
73
]. With respect to this, determinants
of LD synthesis such as the cellular levels of TAG and sterols are of prevalent meaning
to the whole context. In yeast, two enzymes are regarded as the main actors in TAG
production, Lro1p (lecithin cholesterol acyl transferase related open reading frame) and
Dga1p (diacylglycerol acyltransferase 1) [
76
]. For sterols, the acyl-CoA:sterol acyltransferase
Are1p and its paralog Are2p are the main sterol esterification tools in yeast [
77
]. Together
these enzymes regulate the TAG:sterol balance to a ratio of 1:1 in yeast LDs [
78
]. We
showed that the simultaneous overexpression of all Lro1p and Dga1p enzymes, as well as
Are1p and Are2p (single overexpression of each enzyme), yields an extension of both the
chronological and replicative lifespan of S. cerevisiae [19].
This stimulation of LD synthesis resulted in less mitochondrial fragmentation and
reduced production of ROS, which normally increase during aging. Contrarily, a mutant
strain devoid of LDs (lro1
∆
,dga1
∆
,are1
∆
,are2
∆
) suffers from a significantly shortened
chronological lifespan and experiences a burst of ROS production already within one day
of cultivation, suggesting severe mitochondrial defects [
19
]. According to the assumptions
made above, mitochondrial functionality is an essential target for age-related cellular
decline, and it seems plausible that “fitter” mitochondria with maintained integrity will be
beneficial to a prolonged lifespan.
Furthermore, mitochondria have been identified recently to assist the cytosolic pro-
teasome in protein degradation, especially during stress conditions. Underlying this is a
process termed MAGIC (
m
itochondria
a
s
g
uardian
i
n
c
ytosol), which mediates the import
of misfolded proteins into mitochondria where protein degradation is performed by the
matrix-resident protease Pim1p [
79
]. Yeast Pim1p is an ATP-dependent mitochondrial
Lon protease required for the degradation of misfolded mitochondrial proteins, which is
essential to mitochondrial function and maintenance [
80
]. With aging, the activity of Pim1p
ceases, and pim1
∆
yeast mutants lacking Pim1p are marked by a shortened replicative
lifespan and show reduced proteasomal activity connected with an increased accumulation
of oxidized and aggregated proteins in the cytosol [
80
]. In line with this, we also observed a
significant shortening of both the replicative and chronological lifespan in pim1
∆
cells [
19
].
In addition, the mitochondria of pim1
∆
cells showed an abnormal morphology accom-
panied by enhanced ROS production, enlarged LDs, and a delay in the cell cycle. This
premature aging phenotype of pim1
∆
cells could be reversed partially by overexpressing
Lro1p. [
19
] This suggests an important role of LDs in the detoxification/sequestration of

Biomolecules 2023,13, 912 9 of 56
the non-degraded, oxidized protein, which underlines the beneficial role of LDs in cell
integrity by assisting cellular clearance from protein aggregates.
It is noteworthy that the advantageous effects of LDs cannot be seen solely as a function
of LD abundance, but also as a function of LD size and morphology. This is indicated
by the observation that pim1
∆
cells treated with oleate and olive oil showed a reduced
lifespan, revealing a drop in the LD number, with the LDs themselves becoming massively
enlarged. In contrast, overexpression of Lro1p/Dga1p on the pim1
∆
restored the strains’
normal replicative lifespan but led to numerous but smaller LDs [
19
]. Furthermore, cells of
the mutant strain sei1
∆
(SEI, yeast seipin controls LD size, number, and morphology) show
a reduced replicative lifespan but no significant differences to wild-type cells in overall
neutral lipid levels. Different from the wild type, the LDs of sei1
∆
cells are smaller and
show LD clustering. Hence, LD size and distribution also obviously play an important
role in the effect of LDs on lifespan in yeast [
19
]. In this context, it is worth mentioning
that, in yeast cells, life-prolonging interventions such as caloric restriction [
81
], rapamycin
treatment (blockage of the TOR kinase; for details see the following chapters) [
82
], and
sirtuin inhibition [
83
] induce the formation of LDs [
84
–
86
]. In fact, in our own experiments
we observed a modest 1.15–1.20-fold increase in the LD content upon treatment of BY4741
cells with 10 µM resveratrol (unpublished data).
Similar research was performed in the filamentous ascomycete Podospora anserina [
33
].
Here, deletion of the gene PaATG24, encoding a sorting nexin, resulted in impaired au-
tophagy, a reduced vacuolar size, lowered growth rate, and lifespan shortening. Addition
of oleic acid stimulates LD production and gives rise to an extended lifespan in wild-type
as well as PaATG24
∆
cells, revealing a restored autophagic flux and normal vacuolar phe-
notype. Interestingly, oleic acid treatment also diminishes ROS production in Podospora as
result of a bypass of complex I and II of the mitochondrial electron transport chain [87].
Taken together, the research on LDs in yeast provides substantial evidence that LDs,
apart from their well-defined role in lipid metabolism, can also serve as hitherto underrated
“detoxification organelles”, which in orchestration with other processes involved in cellular
maintenance, in particular the autophagic flux, serve as lifespan determinants. Such
protective roles (both for proteotoxic and lipotoxic intervention) were clearly demonstrated
for the model organisms discussed below.
3. Lipid Droplets in Caenorhabditis elegans
Caenorhabditis elegans has proven to be one of the most important model organisms
in aging research. Several milestones in this specific scientific field were achieved in this
nematode. It was shown for the very first time in this worm that a mutation in a single
gene (age-1) can extend the lifespan of a whole organism [
88
]. Further aging pathways
that were unraveled in C. elegans or were studied in great detail include the insulin/IGF-1
signaling (IIS) pathway [
89
], TOR signaling pathway [
90
,
91
], caloric-restriction-induced
signaling [
92
], TGF-
β
-signaling [
93
], AMPK signaling [
91
,
94
] and the HIF-1-dependent
hypoxic response [95].
3.1. The C. elegans “Dauer-Larva”
To gain a better understanding of the role of LDs in the aging process of C. elegans,
it is appropriate to provide a short overview of its lifecycle, in particular with respect to
the diapause stage of the “dauer larva” resembling a suitable aging model. In C. elegans,
two sexes can be distinguished, self-fertilizing hermaphrodites and males, each composed
of an exactly defined number of somatic cells. Upon fertilization, eggs are laid. After
embryonic development and hatching from these eggs, the nematodes have to pass four
larval stages, each of which ends with a molt, before adulthood is reached [
96
]. Spectacular
in this life cycle and important for aging research is the formation of a so-called dauer
larva. Environmental cues including starvation-, heat-, or population-density-dependent
pheromone secretion at the L2 molt phase are potential inducers of the dauer larva. In
this specific phase, the worm stops eating and ceases muscular activity in the pharynx

Biomolecules 2023,13, 912 10 of 56
but retains full mobility. The dauer larva has a reduced intestinal lumen and specialized
cuticle. As soon as the harmful environmental influences end, the larva exits the dauer
stage and, after the third and fourth molts, forms an adult worm. Strikingly, this dauer
stage can extend the lifespan up to 70 days, which is close to four-fold the average lifespan
of an adult nematode (about 18–20 days at 20
◦
C) [
97
,
98
]. In the development of many
longevity concepts, the C. elegans dauer larva played a significant role since all of the
above-mentioned pathways (e.g., TOR, IIS, TGF-
β
signaling) that affect worm longevity also
modulate entry of the worm into the dauer larva stage. Critically, many of the mechanisms
contributing to lifespan extension have to be considered dauer-related, but also dauer-
independent [
99
]. As we discuss in the following section, this may also apply to LDs; the
dauer stage phenotype of C. elegans shows a close linkage to the detoxifying effects of LDs,
a LD function that may also play a role in cellular maintenance in the adult worm as well
as higher organisms.
3.2. Detoxifying Role of Lipid Droplets
The primary sites for fat storage in C. elegans are cells of the intestine and the hypo-
dermis. In these cells, three fat deposits were identified, namely LDs, lysosome-related
organelles (LROs), and undefined vesicles [
100
]. These sites for fat storage differ in their
abundance and lipid composition. Inspection of intestinal cells by Raman scattering mi-
croscopy revealed that 18% of the cellular area is covered by LROs and 4% by LDs [
101
].
In contrast to yeast cells in which TAG and sterol esters are stored in LDs, there is a clear
separation between LROs and LDs in C. elegans fat storage. In the worm, LDs are enriched
in TAG, whereas cholesterol is mainly deposited in LROs [
101
]. Connecting LDs with LROs,
it is speculated that LROs mediate the flux of fatty acids from LDs to either mitochondria
or peroxisomes [
102
]. Until recently, the discrimination between LROs and LDs in the lipid
management in C. elegans was widely neglected, and some phenotypes attributed to LDs
more likely may be associated with LROs. Today, however, several specific approaches
and staining protocols are available, which allow a clear distinction between these two
fat-storing cell organelles. Among these, three methods should be mentioned here briefly:
(1) In transmission electron microscopy, LDs appear to be electron lucent, whereas the
more dense LROs appear to be electron-dense and opaque [
103
]. (2) Both Nile Red and
BODIPY are established as vital dyes for monitoring LDs in a broad variety of organisms. In
C. elegans, both dyes show a high affinity for LROs, whereas Nile Red fails to stain LDs in
living nematodes [
100
]. (3) Some bona fide LD-resident proteins have been identified in
C. elegans. One of these proteins is the triacylglycerol lipase ATGL1 which, upon fusion
with GFP, specifically stains LDs but not LROs [103].
3.3. Lipid Droplets, Insulin Signaling, and Autophagy
As already addressed in the preceding chapter, caloric restriction represents one of
the best-known and most reproducible interventions to prolong eukaryotic lifespan. This
phenomenon was first observed in rodents [
104
], and among others was confirmed in yeast
cells [
81
], C. elegans [
92
], Drosophila melanogaster [
105
], and primates [
106
]. In C. elegans,
the intestine as a central organ of the worm is tightly linked to the aging process [
107
].
Therefore, nutrition is of eminent importance to C. elegans and all life-prolonging processes
relate directly or indirectly to caloric restriction. In C. elegans, nutritional supply is covered
by the ingestion of bacteria, and reducing the number of bacteria that are experimentally
fed allows extension of the lifespan of up to 70%. This effect was observed in all phases of
the worm’s life cycle (either growth, reproduction, or post-reproduction phase) [
108
,
109
].
Caloric restriction leads to obvious changes in the C. elegans phenotype, foremost the
reduction in body size, and leads to characteristic changes in lipid metabolism as reflected
by an increased TAG: protein ratio observed in L4 larvae as well as in the adult worm.
As consequence, this increase in TAG levels also manifests in LD size and abundance.
For the wild type, depending on body region and developmental stage, an up to 15-fold
increase in the number of enlarged LDs upon caloric restriction has been reported. The

Biomolecules 2023,13, 912 11 of 56
very same enlargement in LDs was seen in eat-2 mutant worms (suffering from a feeding
defect) that serve as a genetic model for caloric restriction [
108
]. Moreover, in L2 larva,
starvation can induce development to the dauer larva state. This transition is marked
by fat accumulation serving as an internal energy reserve, which occurs in conjunction
with a substantial increase in LD number and density in the dauer larvae [
110
]. The exact
mechanics underlying the outcome of caloric restriction are not completely clear, but have
to be considered multifactorial. Involved processes may, inter alia, comprise (i) the down-
regulation of insulin/insulin-like growth factor 1 (IGF-1) signaling (IIS), (ii) a decline in
TOR signaling yielding elevated autophagy, (iii) increased activation of sirtuins resulting in
gene silencing, and (iv) a more complex regulated decline of the metabolic rate [111–113].
IIS-pathway/FOXO. The evolutionary highly conserved IIS pathway plays an impor-
tant role in nutrient sensing and maintenance of glucose homeostasis. Central to this is
the IIS-regulated expression of a set of genes involved in stress response, energy genera-
tion, drug metabolism, and chaperone activity [
114
]. Concerning the heavily discussed
life-prolonging effect of caloric restriction, certain arguments account for a contribution
of IIS, while others are conflicting, such as the additive effect of caloric restriction and IIS
repression on life extension [
111
,
113
]. In fact, entry of C. elegans L2 larva into the dauer
larva stage is blocked by activated IIS. Screens searching for mutations that promote the
L2/dauer larva transition led to the identification of several IIS-pathway elements. In
toto, the associated genes were given the name daf, as an abbreviation for “dauer for-
mation” variant [
99
]. The first, upstream component of the IIS pathway is a receptor
tyrosine kinase (DAF-2). Upon binding of insulin-like molecules, DAF-2 activates the PI3P
pathway that comprises sequential signaling via phosphoinositide-3-kinase (AGE-1), the
3-phosphoinositide-dependent kinase 1 (PDK-1), and the serine-threonine kinases AKT-
1/2. The final targets of this kinase cascade are the transcription factors DAF-16 (a FOXO
transcription factor; FOXO, forkhead box O) and SKN-1 (a Nrf1,2,3 transcription factor)
which, upon phosphorylation, are blocked from entering the nucleus [
99
,
115
]. Hence, re-
duced IIS upon caloric restriction will allow DAF16 (FOXO) shuttling into the nucleus and
promotion of its activity as a transcriptional regulator. With DAF-16, a central pleiotropic
mediator of cellular stress responses was identified in C. elegans that increases resistance
against stressors such as heat or pro-oxidant regimens, but also promotes fat storage [
116
].
Strikingly, most of the mutations that affect genetic control of IIS in a way that terminally
promotes shuttling of non-phosphorylated DAF-16 into the nucleus prolong the lifespan
of C. elegans in a drastic way; daf-2 [117], age-1 [88], and pdk-1 [118] are such examples. On
the contrary, mutations in daf-16 itself suppress the increased longevity [
119
]. Furthermore,
either mutation or knockdown of some of these IIS-associated genes resulted in a clear
increase in the cellular LD content [
20
,
120
]. This suggests an association of IIS-controlled
LD biogenesis with longevity in C. elegans. Supportive of this, Suriyalaksh et al. showed
that long-lived worms reveal a strong tendency for an increased LD content [
20
]. However,
it is also reported that an extreme excess of LDs upon passing a certain cut-off is negatively
associated with lifespan [
20
]. These findings perfectly match with our observations in
yeast. These demonstrate that a moderate increase in LDs (achieved by overexpression of
Dga1p and Lro1p) results in the prolongation of both replicative and chronological lifespan,
while overloading yeast cells with oleate (i.e. monounsaturated fatty acids) resulted in
super-sized LDs and a clear trend to lifespan shortening (unpublished data and [19]).
TOR pathway and nutrient sensing. Another important rheostat of caloric restriction
responses is the nutrient-sensing TOR (target of rapamycin) complex that exists in all eu-
karyotes, from yeast to humans. As the name indicates, the central component of the TOR
pathway is the serine/threonine protein kinase TOR, and it is best described in mammals
(termed mTORC, mammalian TOR complex). This kinase is either associated with the bind-
ing protein raptor (Regulatory Associated protein of mTOR), forming the TOR Complex 1
(TORC1) or rictor (Rapamycin-Insensitive Companion of mTOR) forming the TOR Complex
2 (TORC2) [
121
]. The regulation of TOR activity is highly complex, combining several input
signals, such as availability of nutrients (e.g., glucose), growth factors, amino acids, and

Biomolecules 2023,13, 912 12 of 56
oxygen. Of importance, active TOR/mTORC inhibits autophagy, the depletion of nutrients
such as those seen under starvation conditions, but also stress-derived signals, as well as
the pharmacological inducer rapamycin leading to TORC decomposition, in which the
deactivation of TORC results in the de-repression of autophagy [
122
]. Relevant to the role
of IIS in caloric restriction, mTOR signaling shares a certain cross-talk with the IIS-pathway.
Underlying this is the IIS-related activation of AKT-1/2, which leads to phosphorylation
and inactivation of the tuberous sclerosis complex (TSC) consisting of TSC1 and TSC2. As
part of an active TSC, TSC2 serves as a GTPase-activating protein (GAP) for Rheb, a small
GTPase acting as a positive regulator of mTORC1. Upon IIS/AKT-dependent phosphoryla-
tion, TSC2 becomes destabilized, rendering the TSC inactive, which results in mTORC1
activation [
123
]. In the same way, TORC1 activity is also regulated by TORC2, which also
leads to AKT-1/2 phosphorylation [
124
]. In addition, TORC1 is associated with further
GTPases such as the Rag GTPases RAGA and RAGC, which are controlled by glucose-
as well as amino-acid-pool-dependent signaling [
125
]. Among the manifold downstream
targets of TOR, 4E-BP (eIF4E-binding protein) and S6K1 (S6 kinase 1) are the best known.
4E-BP is an inhibitor of the eIF4E translation initiation factor 4E and, by forming a complex
with eIF4E, blocks translation. Upon TOR-mediated phosphorylation, 4E-BP is released
from the eIF4E/4E-BP complex and translation is initiated [
126
]. S6K1 is another target
of TOR belonging to the AGC family of protein kinases. TOR together with PDK1 phos-
phorylates S6K1 and, in a progression of this kinase cascade, leads to phosphorylation
of the ribosomal protein S6, resulting in the translation of specific mRNAs [
127
]. Most
(but not all) components of these highly conserved pathways are existent in C. elegans and
higher eukaryotes (the C. elegans homologues are given in brackets): TOR (LET-363); Raptor
(DAF-15); Rictor (RICT-1); RHEB (RHEB-1); RAGA (RAGA-1); RAGC (RAGC-1); 4E-BP
(IFET-1); S6K1 (RSKS-1) [
121
]. In C. elegans, it was shown that deletion of the TOR homolog
LET-363, and as a result deletion, of the central element of the TOR pathway, leads to an
arrest of the larva in the L3 stage [
128
]. On the other hand, the RNAi-mediated knockdown
of let-363 resulted in a dramatically increased mean lifespan and elevated lipid accumu-
lation that is seen most obviously in intestinal cells [
128
]. Moreover, as revealed either
by gene mutation or knockdown (RNAi) experiments, the reduced expression of TOR-
pathway-associated compounds RICT-1 [
129
], DAF-15 [
90
], RHEB-1 [
130
], RSKS-1 [
131
],
RAGA-1 [
129
], and RAGC-1 [
129
] extends the lifespan of C. elegans, thus mimicking the
effect of nutrient depletion/caloric restriction (for a detailed review see [
121
]). In addition,
mutations in rct-1 [
132
,
133
] and rsks-1 [
134
], and a RNAi-mediated knockdown of DAF-
15 [
90
], resulted in the formation of numerous enlarged LDs, especially in intestinal cells,
with the phenotype seen in daf-15 RNAi experiments marked by the increased presence of
autofluorescent granules, most probably representing LROs [
90
]. In this context, it appears
noteworthy that the aging-dependent sequestration of protein aggregates by LDs has been
reported to occur in mouse intestinal tissue, where this process may serve the removal of
protein aggregates for subsequent autophagic digest via lipophagy [
135
]. Hence, it cannot
be excluded that the appearance of enlarged LDs/LROs in intestinal cells of C. elegans
is also connected with a similar LD/autophagy-associated process of protein aggregate
clearance. The intrinsic linkage between TOR inhibition and LD synthesis seen in C. elegans
represents an evolutionary recurring motif, as discussed below for Drosophila melanogaster
and Homo sapiens, and also holds true for unicellular eukaryotes (Figure 2and Section 2).
In S. cerevisiae, treatment with substances such as rapamycin or methionine sulfoximine
inhibits TOR signaling and promotes chronological lifespan of the yeast cells [
82
]. Another
consequence of rapamycin exposure is an increased TAG synthesis that is accompanied
by increased LD numbers [
84
]. However, both investigations did not show whether the
observed life-prolonging effect of rapamycin in yeast cells is due to an interplay between
TOR inhibition and LD biogenesis, or whether these outcomes represent independent
effects of rapamycin.

Biomolecules 2023,13, 912 13 of 56
3.4. Lipid Droplets and TGF-βSignaling
As a further pathway contributing to lifespan expansion upon caloric restriction, we
discuss the influence of TGF-βsignaling on longevity in C. elegans [136]. Five members of
the TGF-
β
superfamily have been identified in C. elegans, and with respect to its implication
in aging, dauer larva formation, and fat storage, we focus on the TGF-
β
homolog DAF-
7 [
137
]. Produced under favorable conditions by sensory, amphid ASI neurons, DAF-7/TGF-
β
stimulates TGF-
β
receptor (TGF-
β
R)/Smad-based signaling. The final downstream target
affected upon DAF-7 ligation in the TGF-
β
/Smad pathway is the nuclear factor co-SMAD
DAF-3, which binds to the Sno/Ski transcriptional co-factor DAF-5 and promotes by this
expression the genes responsible for dauer larva formation. Homodimeric DAF-7 binds to
a heterotetramer consisting of two molecules, DAF-1 and DAF-4, resembling the TGF-
β
R
homologous receptor localizing to the plasma membrane in C. elegans. In a canonical mode,
DAF-7 ligation leads to activation of DAF-4, a Type II TGF-
β
R which phosphorylates and
activates the type I TGF-
β
R DAF-1 that itself is a serine/threonine kinase. Downstream
to this, activated DAF-1 phosphorylates the R-Smad homologs DAF-8 (Smad2) and DAF-
14 (Smad8) which, upon heterodimerization, translocate to the nucleus. In the nucleus,
heterodimeric DAF-8/DAF-14 inhibits the Co-factor/Co-Smad DAF-3 (Smad4) and the
Sno/Ski homologous transcription factor DAF-5, and, due to this, blocks the transcription
of dauer-specific genes [
138
–
140
]. Each intervention that blocks TGF-
β
/Smad signaling
(i.e., mutations in daf-7, daf-4, daf-8, daf-1, and daf-14) prolongs C. elegans lifespan, whereas
each opposite intervention boosting TGF-
β
signaling (mutations in daf-3 and daf-5) short-
ens the lifespan [
141
]. In animals with mutated daf-7, daf-1, and daf-4, the improved
lifespan was paralleled by a 2.5-fold increased fat accumulation, most probably in LDs com-
pared to wild-type worms. It is noteworthy that this increased fat storage was independent
of a reduced food intake, but was a specific outcome of defective (inhibited) TGF-
β
/Smad
signaling [142].
3.5. Significance of Lipid Droplet Accumulation to C. elegans Lifespan
These findings together demonstrate the strong interference of aging-associated path-
ways with nutrient sensing (IIS, TOR signaling) and developmental growth regulation
(TGF-
β
signaling) in controlling/blocking dauer larva transition [
143
]. Accordingly, in-
activation of each of these pathways will support longevity by promoting the exit from
normal development to the dauer larva state. Of high relevance to this, all of the “anti-aging
interventions” in C. elegans addressed in this review were accompanied by a strong accu-
mulation of LDs. Since the formation of dauer larvae is also inextricably linked to a shifted
LD abundance, it may be questioned whether this reflects an evolutionary developed mech-
anism of intrinsic energy supply under poor environmental conditions and/or the degree
to which LD accumulation is an active, driving force in the aging process. The following
section aims at addressing this question by discussing two findings, which suggest an
active role of LDs in coping with cellular stress conditions.
The Lapierre group showed that overexpression of the autophagy receptor sequesto-
some (SQST-1) resulted in a decreased lifespan of C. elegans at 25
◦
C [
144
]. Upon applying
a genome-wide RNAi screen, they identified candidates that were able to alter the protein
content of a SQST-1-GFP fusion protein, and the candidate list revealed a huge overlap with
proteins that were found to be part of the LD proteome of nematodes. In order to boost
the cellular LD content, the atgl-1 lipase was silenced, which shifted LDs in numbers and
size, and, as a result, replicated observations frequently made in long-lived worms. This
increase in lipid storage was accompanied by a strong lifespan extension and accumulation
of SQST-1 at LDs. Interestingly, the SQST-1 relocalization to LDs was not restricted to
this autophagic receptor, but was also observed for misfolded as well as ubiquitinylated
proteins, suggesting a general role of LDs in protein homeostasis [
144
]. This matches
perfectly with our observations made in yeast cells, showing that stimulating LDs can
prolong both the replicative and chronological lifespan, most likely by detoxifying harmful
proteins [19,35].

Biomolecules 2023,13, 912 14 of 56
Independent of caloric restriction, but the same as in S. cerevisiae, there appears to be
an interplay between lipid droplets and the ER-associated degradation machinery (ERAD).
It was shown that an oleate-rich diet stimulates LD levels, ERAD activity, and longevity in
C. elegans. The life-prolonging effect of oleate was also strongly dependent on LD-associated
proteins such as Plin-1 and Fitm-2 (fat-storage-inducing transmembrane 2) [
145
]. Plin-1 is
the only known perilipin in C. elegans [
146
,
147
], whereas Fitm-2 is essential for the budding
of LDs from the ER [
148
]. Another publication also confirmed the life-prolonging effect of
monounsaturated fatty acids such as oleate [149].
As seen in most eukaryotes, a fraction of LDs can reside in the nucleoplasm; in
C. elegans, such nuclear LDs are found in the nuclei of intestinal cells, especially under
stress conditions [
89
]. Some of these LDs were shown to be covered by heterochromatin,
which is translocated apart from the nuclear lamina. Mosquera et al. speculate that this
inward movement of heterochromatin could result in relieved gene silencing and, as a
result, promote the aging process. Furthermore, the authors also reported that giant nuclear
LDs may come in close contact with the nuclear lamina, especially in areas devoid of
lamina. According to the authors’ assumptions, these giant LD/nuclear lamina-contact
zones could be responsible for ruptures of the nuclear lamina, which are seen frequently in
C. elegans intestinal cells [
150
]. Although this may hold true, it is tempting to speculate in a
different direction by considering the lamina/nuclear LD/chromatin association areas as
sites specialized for chromatin/DNA repair.
In the model organism Drosophila melanogaster, with the development of specialized
cells and tissues, the evaluation of the role of LDs in the aging process is much more
difficult, but here, too, a clear interconnection between lifespan extension, LDs, mTOR, and
IIS is beginning to emerge, as the following section shows.
4. Lipid Droplets in Drosophila melanogaster
The fruit fly Drosophila melanogaster is a well-described animal model organism in
genetics, developmental biology, and cell and molecular biological research on the mechan-
ics of senescence and aging [
151
,
152
]. Throughout evolution, the “hallmarks” defined for
mammalian aging [
74
] are highly conserved and can also be investigated in Drosophila.
Consequently, studies in flies identified evolutionary conserved gene mutations, endocrine
and cellular signaling mechanisms, and tissue- and environment-specific factors including
their interactions with the genetic background, that affect lifespan [
153
]. Focusing on the
metabolic aspect, several lipid-metabolism-associated contexts of substantial physiological
and pathophysiological relevance have been addressed in Drosophila. These comprise
research on TAG storage and mobilization from LDs that have been addressed in Drosophila,
which also reflect lipid metabolism in humans, including age- and lipid-associated diseases.
As will be outlined in this section, evidence is increasing that LDs are deeply involved in
the interconnection of nutritional, metabolic, and stress-associated signaling, emphasizing
their role as cell organelles with multifaceted implications in lifespan control.
4.1. Lipid Droplets and Drosophila Development
Development of the fruit fly proceeds in an indirect mode, with each developmental
stage (egg/embryo–larva–pupa/metamorphosis–imago) differing under nutritional as-
pects. Lipid homeostasis is regulated in a food-dependent mode during the larval-hood and
in the adult fly (imago). In contrast, “nutritional supply” for embryogenesis depends on the
maternal deposition of LDs during oocyte maturation, and the energy needed for metamor-
phosis (pupa, imaginal disc development) is supplied by the LD-rich fat body established
during the larval stage, which shares functional equivalence with the mammalian liver and
adipose tissue. Reflecting the adverse effects of diet-associated obesity in higher organisms
predisposing for a number of pathological conditions, excessive consumption of a high-fat
diet decreases lifespan in Drosophila [
154
]. In line with this, Drosophila mutants devoid of
adipokinetic hormone Akh (a functional analog of glucagon), serving as a genetic model
of obesity, suffer from lifespan shortening accompanied by characteristic, age-dependent

Biomolecules 2023,13, 912 15 of 56
changes in the lipid profile that especially affects the TAG signature [
155
]. This accounts
for a selective TAG degradation from LDs in moribund flies yielding a senescence-specific
lipid signature.
As stated above, LDs play crucial roles during all stages of Drosophila ontogenesis. For
instance, oocytes are loaded with TAG-rich LDs to comply with the metabolic demands
of embryogenesis [
156
–
159
]. This maternally driven process is regulated by the Drosophila
perilipin 2 (PLIN2) homologue LSD-2 (lipid storage droplet 2) [
160
–
162
]. Beyond this, LSD-2
and LSD-1, the homologue of human perilipin 1(PLIN1) [
163
], act as central regulators of LD
growth and fat storage over the whole lifespan of Drosophila [
164
]. During the larval stage,
LDs are indispensable to fat body growth, which is controlled via metabolic signaling involv-
ing the IIS/FOXO (dFOXO in Drosophila) pathway as well as endocrine signaling [
165
–
167
].
Moreover, LDs connect fat body growth with molting, since synthesis of ecdysone (a pre-
cursor of the molting regulatory steroid hormone 20-hydroxyecdysoen) requires cholesterol
trafficking from LDs to autophagosomes, cholesterol-rich LDs accumulating in the larva if
autophagy is inhibited by the accumulation of fat [
168
]. Larval development is influenced
in a nutrition-dependent mode by the LD-associated protein CG9186/Sturkopf, which
regulates larval growth by connecting LD biogenesis to nutritional supply via interaction
with the IIS/dFOXO pathway and hormone (juvenile hormone) signaling [
169
]. In the
absence of LDs, the CG9186/Sturkopf protein localizes to the ER, but translocates to LDs
upon induction of lipid storage [
170
]. Interestingly, in Drosophila CG9186/Sturkopf null
mutants, TAG storage is neither affected in embryos (laid down by mutant mothers) nor in
mutant larva containing normal LDs, an effect that is not completely understood at present,
but could be based on distinct dFOXO targets such as the Drosophila lipase brummer, an an-
tagonist of LD-regulatory LSD-2 (see the next section) [
169
]. In contrast, CG9186/Sturkopf
null mutations become manifest only in adult flies, which show a markedly reduced TAG
storage. In addition, adult CG9186/Sturkopf mutants also reveal a markedly increased
protection from desiccation stress (supposedly due to a changed hydrocarbon composition
of the cuticula) and reduced locomotor activity [
169
]. Moreover, as we discuss below,
the reduced LD content seen in adult CG9186/Sturkopf mutants negatively affects stress
resistance as well as lifespan of the adult flies.
4.2. Control of the Lipid Droplet Pool in Drosophila Adipocytes
Interestingly, Drosophila adipocytes bear spatially and functionally distinct LD pools
that access distinct lipid pools for their individual maintenance [
171
]. Larger LDs, residing
in the central cell body, require supply by fatty acid synthetase FASN-1 de novo lipogenesis,
whereas smaller LDs locating to the cell periphery require gut-derived lipophorin shuttle
(Lpp)-dependent lipid supply. The population of small peripheral LDs stays in direct
contact with the plasma membrane and its organization changes during fasting periods.
This starvation-associated effect is regulated by the protein Snazarus, which binds LDs at
the ER-plasma membrane contact sites via a C-Nexin domain. TAG storage is enhanced
upon Snazarus overexpression, which confers resistance to starvation conditions and yields
lifespan prolongation [
171
]. Conversely, starvation also leads to the up-regulation of the
Drosophila lipase brummer (bmm), an orthologue of mammalian adipose triglyceride lipase,
which binds to LDs via a so-called brummer domain and elevates TAG mobilization from
LDs [
157
]. Under normal feeding conditions, loss of bmm activity causes a moderate
lifespan reduction, with the mutant flies developing an obese phenotype with adipocytes
containing markedly enlarged LDs. In contrast, starving bmm mutants show a marked
lifespan extension [
157
], which is considered to be due to the decelerated TAG mobilization
from LDs [
172
], albeit the involvement of other lipases such as doppelgänger von brummer
(dob) or other putative starvation-induced lipases may also play a role [157].
As mentioned before, the LD pool of Drosophila adipocytes is controlled by LSD-1
(PLIN-1) and LSD-2 (PLIN-2), which affect bmm lipase activity in opposite directions. While
LSD-1 supports lipolysis by recruiting bmm to LDs [
173
], LSD-2 antagonizes bmm access
and protects LDs from TGA mobilization [
173
]. Conversely, binding of bmm to LDs under

Biomolecules 2023,13, 912 16 of 56
starvation conditions antagonizes the anti-lipolytic activity of LSD-2 [
157
]. Concerning
the LD phenotype, LSD-2 mutants show no peculiar alterations [
164
]. In contrast, LSD-1
deficiency promotes LD accumulation [
168
], with the adipocytes developing a reduced
number of markedly enlarged, “giant” LDs during the larval hood and in the adult fly [
163
].
Moreover, it is clearly shown that the lifespan of LSD-1 mutants is reduced under starvation
conditions [
173
]. Bi and co-workers proposed that LSD-1 and LSD-2 regulate lipolysis in
an opponent, LD-size-dependent mode, with LSD-1 promoting TAG mobilization from
large LDs, but LSD-2 protecting small LDs from bmm-induced lipolysis [
173
]. The same
study also demonstrated that LSD-1 is able to adopt the anti-lipolytic property of LSD-2
under certain conditions, which points to a functional redundancy between these perilipin
homologs in Drosophila.
LSD-2 affects the Drosophila LD pool already at the earliest developmental stages, with
the oocytes of LSD-2 mutant females showing a substantially reduced TAG content, causing
impaired embryogenesis [
162
]. Similarly, the fat body of LSD-2
−/−
homozygous larva
contains less than 75% TAG as seen in normal flies. Pointing to additional functions not
directly related to lipid storage, LSD-2 is also required for the endoreplication of cells in the
larval salivary gland, with the loss of LSD-2 activity resulting in enhanced ROS generation
and stimulation of JNK (c-jun amino-terminal kinase)-dependent apoptotic cell death [
174
].
Similarly, LSD-2 provides lipid storage in imaginal discs during metamorphosis, but
also plays a distinct role in imaginal disc cell growth and differentiation. For instance,
LSD-2 expression is stimulated in the wing imaginal disc upon overexpression of the gene
vestigial (vg), which encodes a transcription factor controlling wing cell proliferation and
differentiation [
175
]. Correspondingly, knockdown of LSD-2 promotes cell death in the
wing imaginal disc, which involves the dFOXO-dependent up-regulation of pro-apoptotic
reaper [
176
]. Notably, reaper is an orthologue of human autophagy/mTOR-regulatory
TSC1 (see Section 3.3) and stimulates apoptosis in Drosophila by suppressing anti-apoptotic
Diap1 [
177
]. Diap1, in turn, represents the orthologue of mammalian caspase-activation-
inhibiting XIAP, which connects developmental (TGF-
β
1/BMP) and (oxidative) stress-
associated (NF-
κ
B and Nrf2) signaling with apoptosis in mammalian cells [
178
–
180
]. These
findings point to a distinct role of LDs and LD-associated perilipins (LSD-1, LSD-2) in
cellular growth control, interconnecting metabolic regulation with cell cycle/cell death
checkpoints, which also affects Drosophila lifespan. As already stated, this was shown for
LSD-1 under starvation conditions [
173
] and the involvement of LSD-2 in lifespan control
under a high-fat diet has also been discussed [181].
4.3. Lipid Droplets and Lifespan Extension in Drosophila
Complementary to studies on Drosophila lipid/LD biology, further investigations
addressed the effects of caloric restriction, pharmacological intervention, and the combina-
tion of both on the genotype–environment interaction and the underlying mechanisms in
Drosophila [
182
–
184
]. In agreement with the above-mentioned lifespan-shortening effect
seen for the Akh-deficiency-based genetic model of obesity [
155
], these studies revealed
a positive effect of dietary restriction and nutritional balance on longevity in Drosophila,
for which nutrient sensing and the associated downstream signaling is of pivotal rele-
vance [
184
,
185
]. Similar to the role of IIS/FOXO in C. elegans dauer larva transition and,
as stated, for the involvement of CG9186/Sturkopf in Drosophila development, the IIS-
dependent regulation of dFOXO is also involved in Drosophila lifespan control. This is
indicated by the finding that reduced IIS (as seen upon caloric/dietary restriction) results
in enhanced dFOXO activity and lifespan extension in the fly [
186
,
187
]. The IIS-dependent
effects are largely mediated by the JNK/dFOXO stress response pathway [
187
–
190
] and
other transcription factors acting downstream of dFOXO such as AOP (Anterior open,
an E-twenty-six (ETS)-family transcriptional repressor), which, in a coordinated manner,
mediate the lifespan extension in Drosophila [
191
]. It is noteworthy that dietary inputs
address organ-specific interactions. For instance, overexpression of dFOXO in the fat
body of the head leads to an extended lifespan only under high protein conditions [
192
].

Biomolecules 2023,13, 912 17 of 56
Interestingly, flies bearing lifespan-extending deteriorations in IIS, such as those caused
by the deletion of insulin-like peptide 2, show an increased body fat storage and are more
resistant to lipophilic toxins and oxidative stress [
193
]. In sum, the regulatory network
mediated by IIS that controls LD size and number is highly complex. In Drosophila nurse
cells (which dump LDs to the oocyte), strong IIS stimulation, e.g., due to the loss of the
IIS antagonist PTEN (phosphatase and tensin homologue), leads to the accumulation of
abnormally enlarged LDs [
194
,
195
]. This is counterbalanced by dFOXO, which induces
lipases such as brummer [
196
], which is antagonized by LSD-2, protecting LDs from bmm
lipase access as explained above. In this context, it is worthwhile to mention again that
LSD-2 can only be found on small LDs, whereas LSD-1 can anchor to LDs of different
sizes [
173
]. Hence, IIS/dFOXO-dependent responses are specifically controlled at the LD
level by LSD-1 and LSD-2.
Moreover, excessive intestinal stem cell (ISC) proliferation causing intestinal dysplasia
in aged flies is suppressed by reduced IIS/JNK signaling, which restores proliferative
homeostasis and extends Drosophila lifespan [
197
]. Connected with this, ER stress-associated
UPR (UPR
ER
, see Section 2) has been shown to also play an important regulatory role in ISC
proliferation, with the chronic UPR
ER
-dependent hyper-stimulation of ISC proliferation
being causal to age-related intestinal dysplasia [
198
]. Central to this UPR
ER
response in
ISC proliferation in Drosophila is an orchestrated interplay involving (i) the activity of PKR-
like ER kinase (PERK) which is controlled by the JAK/Stat pathway, (ii) transcriptional
control via Ire1 (endoribonuclease 1)-mediated splicing of the transcription factor Xbp1
(X-Box binding protein 1), and (iii) the activation of another transcriptional regulator
ATF6 [
198
,
199
]. With respect to lipid metabolism, UPR
ER
-dependent Ire1/Xbp1 signaling
is of special relevance since it connects ER stress to triacylglycerol synthesis and lifespan
extension in Drosophila [
200
]. Upon caloric restriction, Ire1/Xbp1 signaling promotes
lipogenesis and TAG accumulation in intestinal enterocytes and prolongs lifespan of the
fly; this effect also involves activity of the transcription factor sugarbabe (a Gli-like zinc
finger transcription factor involved in the carbohydrate metabolism). With respect to
this, the concept that an Ire1/DGAT2-based shift of LD biogenesis, such as that seen
under conditions of ER stress in the mouse liver, could improve LD-mediated protection
from phospholipid oxidation [
47
] (see Section 5), is of particular interest. Indeed, LDs
have been shown to confer antioxidant properties in Drosophila by incorporating TAG
redistributed from PUFAs and, as a result, protect PUFAs from lipid peroxidation in neural
glial cells [
201
]. Hence, it would be interesting to investigate whether aging- and/or
starvation-induced ER stress leading to the stimulation of Ire-1 and Xbp1-dependent LD
biogenesis also confers lifespan extension in Drosophila by improving the anti-oxidative
capacity. If so, however, this protection will demand tight control of LD abundance since
the ROS-stimulated accumulation of LDs is at risk of promoting neurodegeneration in
Drosophila [
46
]. According to this study, ROS generation emerging from mitochondrial
dysfunction is causal to glial LD accumulation, a finding that is considered to indicate
an evolutionary conserved process of pathogenic relevance to neurodegeneration. It is
tempting to speculate whether harmful mitochondrial proteins can also be shuttled to LDs
in Drosophila, as shown in yeast [
7
]. Pointing in this direction, analysis of the LD proteome in
embryonic [
202
] and adult [
203
] tissue in Drosophila revealed the presence of mitochondrial
proteins in LDs. In addition, similar to the binding of protein-aggregate-enriched IBs by
LDs in yeast [
8
], protein aggregates formed upon ER stress/oxidative stress could also be
sequestered by specific LD binding and, as a result, confer protection in the neuroglia. In
fact, LD binding of protein aggregates has been demonstrated recently for mouse intestinal
tissue which may be followed by lipophagic digest [
135
]. Thus, a similar process could
contribute to the degradation of protein aggregates via lipophagy in neuroglia, but also
other tissues, such as the intestine in Drosophila.
Autophagy–TOR pathway. Similar to C. elegans, the autophagy–regulatory TOR
pathway is also intimately involved in metabolic homeostasis and lifespan control in
Drosophila [
204
]. In general, the inhibition of mTOR signaling results in lowered trans-

Biomolecules 2023,13, 912 18 of 56
lational activity and elevated autophagy which improves proteostasis [
205
], a critical
hallmark of aging [
74
]. It should already be mentioned at this point that, in Drosophila,
both nutrient deficiency and TOR inhibition also lead to an increase in LD size [
206
].
The necessity of enhanced, functional autophagy for lifespan extension in Drosophila was
demonstrated by feeding experiments in adult flies using the TOR inhibitor rapamycin, and
it was shown that the stimulation of autophagy resulted in prolonged survival of starving
wild-type animals but also enhanced the lifespan of Drosophila IIS mutants [
207
]. Moreover,
this investigation also demonstrated that rapamycin-induced autophagy also improves
the resistance towards paraquat (1,1
0
-Dimethyl-4,4
0
-bipyridin)-induced oxidative stress.
Paraquat serves as an insecticide via the cytochrome P450/Fenton-reaction-dependent
formation of free hydroxyl radicals [
208
], which impair intestinal regeneration and cause
substantial cell damage in the aging fly [
170
]. Revealing the involvement of LDs, flies
with a reduced LD content such as that found in adult CG9186/Sturkopf mutants show
reduced survival and a decreased lifespan upon paraquat treatment [
169
]. Pointing further
to the role of autophagy, Drosophila mutants lacking a proper autophagic flux due to mu-
tation of the autophagy–regulatory gene Atg8a (autophagy-related 8a) show a shortened
lifespan accompanied by enhanced protein oxidation and ubiquitination, effects which
are aggravated by pro-oxidant conditions [
209
]. Contrarily, the age-dependent decline
in autophagy seen in Drosophila, especially in the nervous tissue as a result of reduced
Atg-expression, is counteracted by Atg8 overexpression, which improves oxidative stress
tolerance and longevity in aged flies. As stated above, the transfer of cholesterol from LDs
to Atg8-rich vesicles (autophagosomes) is essential to larval hormone synthesis, with the
up-regulation of TOR limiting Atg8 expression and autophagosome formation, which leads
to LD accumulation [
168
]. With respect to this, it is tempting to hypothesize about a similar
Atg8-related mechanism shifting LD numbers in the adult fly in support of cell surveillance
if autophagy declines. In addition to Atg8, a connection with increased longevity and
enhanced stress responses was reported also for other compounds, enhancing autophagy
especially in the nervous system of Drosophila. These include AUTEN-67 and 99 (autophagy
enhancer-67 and -99), acting downstream of TOR at the level of autophagosome membrane
formation [
210
,
211
] and spermidine, which elevates autophagy upon interference with
epigenetic control and yields lifespan expansion in yeast, C. elegans,Drosophila, and human
cell lines [210–212].
Furthermore, autophagy is also stimulated by nutritional factors such as flavonoids,
a class of plant polyphenolic compound with well-known antioxidant properties [
213
].
Among these, isoquercetin and xanthohumol have been proven to boost LD formation in
Drosophila, especially in cells of the nervous system [
214
]. In addition, xanthohumol was
shown to increase the resistance of adult flies to several stressors (e.g., hydrogen peroxide,
paraquat, starvation, and heat) and prolong lifespan in Drosophila [
215
]. As discussed below,
flavonoids also exert similar protective, antioxidant effects in the human system [216]; for
instance, quercetin counteracts liver steatosis by inhibiting lipid peroxidation [217].
As mentioned above, TOR (mTOR) signaling directly affects the translational activa-
tor S6 kinase [
204
,
207
] and the translational repressor 4EBP [
207
,
218
,
219
]. In Drosophila,
additional nutritional sensors have been described that modulate lifespan, including the
transcription factor ATF4 [
220
], amino acid deprivation–activated kinase GCN2 acting
upstream of ATF4 [
221
], GCN2 deficiency leading to a massive loss of TAGs and thus
LDs [
222
], and AMP-activated protein kinase (AMPK). From the mechanistic point of
view, AMPK acts as a central, positive regulator of autophagy by inhibiting mTORC1 (via
phosphorylation of TSC-associated TSC2). AMPK extends the lifespan of adult flies in an
autophagy-dependent mode, specifically affecting the central nervous system and intesti-
nal tissue [
223
,
224
]. In contrast, reduced AMPK activity is associated with an enhanced
susceptibility to starvation-induced lethality (especially in the larva; AMPK null mutants
are larval lethal) and abnormal lipid accumulation marked by the accumulation of enlarged
LDs in larval tissue [
225
]. Of note, this LD-related phenotype is seen in normally fed,
mutant larva lacking AMPK activity and resembles the phenotype observed in oenocytes

Biomolecules 2023,13, 912 19 of 56
of starving wild-type larva [
226
]. Oenocytes are the primary site of LD accumulation seen
in starving Drosophila larva and, analogous to the role of human hepatocytes in systemic
lipid homeostasis, are sites of lipid release from the larval fat body during starvation.
Strengthening the crucial role of autophagy on lifespan control, rapamycin partially im-
proved the survival of adult flies with reduced AMPK activity under starvation conditions.
Interestingly, the type-2 diabetes therapeutic compound metformin, which antagonizes IIS
but stimulates AMPK signaling, and, as a result, also shifts autophagy (recently reviewed
in [
227
]), extended lifespan in C. elegans and healthy mice [
228
,
229
]. However, Slack et al.
reported that metformin failed to prolong lifespan in adult Drosophila; at the same time,
the metformin-mediated activation of AMPK resulted in a drop in TAG levels [
230
]. With
respect to this, it is noteworthy that metformin inhibits TOR in Drosophila independently
of AMPK stimulation [
231
] and Slack et al. suspected a potential negative interference
of metformin effects, not related to AMPK, with positive effects of AMPK activation on
Drosophila lifespan [
230
]. Support to this is provided by the finding that overexpression of
the serine/threonine kinase LKB1, another positive regulator of AMPK, indeed extends
lifespan in Drosophila [
232
]. LKB1-null mutant flies with reduced AMPK activation show
decreased TAG levels, a phenotype that can be compensated by transgenic expression of
wild-type AMPK [
233
]. Accordingly, this suggests that AMPK activation at reduced LD
numbers does not affect lifespan, whereas AMPK activation at an increased LD abundance
extends the lifespan in Drosophila.
4.4. Lipid Droplets, Transsulfuration, and Cellular Antioxidant Defenses
Together, these findings emphasize the pivotal role of metabolic regulation and LD
homeostasis in lifespan determination. At its most basic, this applies to the maintenance
of metabolite pools, especially under dietary restriction [
134
], and in particular to the
amino acid balance, which is controlled on the anabolic (translation) and catabolic (protein
degradation, autophagy) levels as a fundament of cellular proteostasis, with its dysregu-
lation representing another hallmark of aging [
74
]. This was demonstrated in a study by
Grandsion et al., which shows that dietary-restriction-based lifespan extension in Drosophila
is abolished by feeding the flies a mix containing all essential amino acids (EAA feed),
while using an EEA-feed omitting methionine does not affect starvation-induced lifespan
prolongation [
234
]. Further research revealed that trans-sulfuration (i.e., the production
of cysteine from methionine-derived homocysteine or cystathionine) plays an important
role in extending survival under starvation conditions in Drosophila [
235
]. Upon dietary
restriction, enhanced trans-sulfuration preserves lipid storage in LDs (to levels seen in
fully fed flies) and, due to the excessive consumption of methionine, lowers overall protein
synthesis. With respect to this, the positive effect of limited methionine availability on
lifespan seems paradoxical. However, the beneficial effect of methionine restriction on
longevity is conserved from yeast to mammals as demonstrated by the improved longevity
seen in yeast, Drosophila, and rodents upon sulfur-amino acid starvation (SAAR, also refer-
ring to cysteine) [
236
–
239
]. Of particular relevance to this, in yeast, methionine starvation
contributes to longevity by stimulating autophagy and an improved vacuolar acidifica-
tion [
239
]. This puts emphasis on the specific role of sulfur-containing methionine and
cysteine pools in cellular responses to nutritional stress and aging, primarily acting on
the autophagic flux. Similar to starving flies in which inhibition of trans-sulfuration (re-
sembling SAAR) lowers TAG levels [
235
], dietary SAAR reduces fat deposition and the
TAG content in the rodent liver [
240
,
241
]. Stressing the aspect of autophagy, SAAR acts
on the same metabolic regulators in the mammalian system—GCN2, ATF4, and AMPK
(for a review see [
236
])—that link nutritional sensing to the onset of autophagy, and act as
lifespan modulators in Drosophila as outlined above.
Pointing to a further, indirectly nutrition-associated aspect of profound relevance to
lifespan control, the extended survival seen in rodents upon SAAR is accompanied by
increased systemic levels of the cysteine-containing antioxidant glutathione (GSH) [
237
].
This agrees well with the inverse correlation existing between aging and systemic GSH

Biomolecules 2023,13, 912 20 of 56
levels [
242
]. In addition, GSH levels are also elevated in long-lived flies upon dietary
restriction [
235
], an effect that is attributable to the trans-sulfuration-based refueling of the
cellular cysteine pool. Explaining this, dietary restriction up-regulates the expression of
cystathionine β-synthase (CBS), a key enzyme catalyzing the conversion of homocysteine
to the cysteine precursor cystathionine, and elevation of CBS synthesis is essential to
lifespan extension in the starving fly [
235
]. In good accordance with this, blocking the final
step of trans-sulfuration-based cysteine synthesis, which is catalyzed by cystathionine-
γ
-
lyase, leads to a marked drop in GSH levels and abolishes the starvation-based lifespan–
extension in Drosophila [
235
]. Connecting the trans-sulfuration pathway to LD biogenesis,
inhibition of cystathionine-
γ
-lyase also lowers the overall LD content in Drosophila [
235
].
Of note, this effect of trans-sulfuration inhibition on LD abundance seems to represent an
evolutionarily conserved motive, since in human ovarian cancer cells the knockdown of
CBS also results in reduced LD numbers [
243
]. Taking into consideration the crucial role
of the GSH/GSSG (the oxidized GS=SG disulfide) redox balance in cellular antioxidant
capacity, the elevation of trans-sulfuration has to be considered pivotal to cellular stress
adaption by connecting metabolic competence to pro-/antioxidant balance. Accordingly, it
makes sense that an increase in GSH is associated with an increase in LDs under oxidative
stress. Considering the pivotal role of GSH in cellular detoxification of peroxides (H
2
O
2
,
but also lipoperoxides) and lipid-peroxidation-derived metabolites such as 4-hydroxy-2-
nonenal (HNE), limitations of GSH availability, ensuing from enhanced GSH consumption
and/or inadequate GSH synthesis, are at considerable risk of promoting oxidative damage
to lipids, DNA, and proteins [
244
,
245
]. Hence, LDs could support cell survival under
conditions of GHS depletion by eliminating oxidized lipids and misfolded proteins as
discussed above. Serving a similar, cytoprotective task, LDs have been shown to adopt the
role of a cellular antioxidant in Drosophila larva by sequestering polyunsaturated fatty acids
from the cell membranes, which yields protection of these lipids from peroxidation [201].
Moreover, as a sulfhydryl group donor, methionine also undertakes the synthesis of iron–
sulfur clusters (Fe-S) via the mitochondrial ISCU (iron–sulfur cluster forming unit) [
246
]. Fe-S
clusters are indispensable for electron transfer in the respiratory chain and energy charge in all
aerobic organisms. In addition, Fe-S clusters are essential to the citrate dehydrogenase activity
of aconitase and are co-factors of DNA repair enzymes [
247
,
248
]. Interestingly, ablation of
ISCU-mediated Fe-S biogenesis leads to increased citrate concentrations, generated from
glucose-derived acetyl CoA, an elevated fatty acid synthesis, and the accumulation of
LDs in human HEK293 embryonic kidney cells [
249
]. Furthermore, in a mouse model
of Friedreich’s Ataxia (FRDA), an autosomal recessive disease marked by substantially
reduced levels of the mitochondrial ISCU regulatory protein frataxin (Ftx) [
250
,
251
], the
absence of Ftx function resulted in Fe-S protein deficiency, mitochondrial iron accumulation,
and increased LD abundance in cardiac muscle cells [
252
]. Similar effects were seen in a
Drosophila model for FRDA, where the Ftx deficiency stimulated both fatty acid synthesis
and lipid peroxidation, as well as LD accumulation in glial cells [
253
]. Both findings suggest
that hampered Fe-S cluster synthesis caused by Ftx deficiency leads to a disturbance of lipid
homeostasis. However, it should not be overlooked that the FRDA/Ftx deficiency also shifts
the iron content of mitochondria as demonstrated by the FRDA mouse model. It is well
known that labile iron (i.e., free Fe
2+
) acts as a central cellular source for the Fenton reaction-
based generation of hydroxyl radicals. In turn, these hydroxyl radicals readily react with
polyunsaturated fatty acids (PUFAs) and, as a result, initiate the lipid peroxidation chain
reaction (LPO), causing potentially lethal cellular damage, which is aggravated by the geno-
and cytotoxic properties of LPO metabolites such as malondialdehyde and HNE [
254
–
256
].
Therefore, the LD accumulation seen in FRDA/Ftx-deficiency could indicate the specific
up-regulation of LD biogenesis as cytoprotective response to iron-mediated oxidative stress.
Support to this come from the recent finding that LDs participate in cellular responses to
the pro-oxidant effects of paraquat (also a source for Fenton chemistry-based hydroxyl
radical formation) in Drosophila [
257
]. It was shown that the RNA binding protein Spen
(Split ends; the Drosophila orthologue of SPEN/SHARP, a regulator of NOTCH signaling)

Biomolecules 2023,13, 912 21 of 56
modulates the LD content in adult glial cells and provides protection from paraquat
cytotoxicity. Conversely, LD biogenesis can also be stimulated by iron deficiency, which
was demonstrated in human ARPE19 retinal pigment epithelium cells treated with the
iron chelator deferiprone (DFP) [
258
]. DFP induces marked changes in lipid metabolism,
including enhanced TAG synthesis, and leads to the accumulation of LDs in proximity
to mitochondria followed by mitophagy. Diacylglycerol O-acyltransferase 1 has been
identified as a stimulus of LD biogenesis under conditions of iron depletion, enabling the
re-esterification of fatty acids regenerated upon macroautophagy [258].
In summary, these findings link metabolic homeostasis to stress tolerance and shed
light on a particular role of LDs in cellular antioxidant defense and cytoprotection also
affecting longevity. Evidence exists that the LD-mediated clearance of oxidized com-
pounds is essential to this task, as suggested by the sequestration of LPO-products by LDs
in Drosophila protecting larval tissue, and especially neuroblasts in imaginal discs, from
hypoxia-induced oxidative stress [
201
]. Therefore, LDs could participate in cellular mainte-
nance under pro-oxidant conditions by acting as a sink for lipid peroxidation products and
other oxidized cellular compounds. However, considering the transient, dynamic nature of
LDs, such “sinks” do not necessarily need to resemble long-term deposits for the potentially
harmful “oxidized waste”. Findings in glial cells of the Drosophila eye point in this direction.
In these cells, loss of the metalloproteinase ADAM17, a trigger of tumor necrosis factor
(TNF)-based signaling, as well as lack of TNF and the Drosophila TNF receptor homologue
Grindelwald [
259
], causes an age-related degeneration of neuronal and glial cells [
260
]. In
this case, accumulation of LDs in glial cells prior to the degradative process confers an
initial protection from glia- and neuron-derived ROS, while the subsequent metabolic
decomposition of LDs leads to the release of toxic lipid peroxides causing cell damage
and neurodegeneration. Hence, LDs may exert opposing effects in oxidative stress/LPO-
dependent contexts, with the outcome being strongly dependent on additional factors such
as nutritional sensing and lipid turnover, as well as different stress qualities (also in terms
of stress duration: short, intermittent, or chronic), linking the protective capacity of LDs to
the aging process.
4.5. Lipid Droplets and (Epi)Genetic Control
Addressing a further aspect of LD biology that may also be connected to cytoprotec-
tion, LDs may locate to the cytosol as well as to the cell nucleus. As already stated for
C. elegans, nuclear LDs (nLDs) exist in many organisms, including Drosophila. Analysis of
the nLD-proteome isolated from rat liver identified a number of proteins, including histones,
cytoskeletal elements (e.g., cytokeratins), proteins involved in transcriptional and transla-
tional control, protein folding, and post-translational modification, and lipid-metabolism-
associated carboxylesterase 1d (Ces1d; cholesteryl-ester hydrolase) [
261
]. Among different
functions in lipid metabolism, mammalian Ces1d contributes to cellular detoxification by
hydrolyzing lipid esters, either derived from xenobiotic or endogenous sources, especially
in the liver and intestine [
262
]. With respect to LD biogenesis, Ces1d deficiency has no effect
on cytosolic (ER-based) LD formation itself, but yields increased numbers of small-sized
cytosolic LDs, an effect that is attributable to a lower lipid transfer to LDs [
263
,
264
]. Refer-
ring to the presence of Ces1d in nLDs in Drosophila, carboxylesterases may also participate
in nLD biogenesis. In mammalian cells, nLDs are formed either de novo at the inner
nuclear membrane or upon translocation of cytosolic LDs, originating from ER resident
lipoprotein precursors, to the nucleoplasmic reticulum terminally moving to the nucle-
oplasm (recently reviewed by Fujimoto [
265
]). In hepatocytes, ER stress promotes LD
shuttling to the nucleus and requires the activity of promyelocytic leukemia protein (PML)
locating to the inner nuclear membrane. PML, which is critical to nuclear signaling, can
be retained in nLDs (then termed lipid-associated PML structures; LAPS) and due to the
PML binding properties, nLDs/LAPS may regulate the PML-mediated control of the gene-
expression, for instance, as part of lipid stress responses [
265
,
266
]. The above-mentioned
association between stress responses and nLDs in C. elegans (see Section 3) may point in a

Biomolecules 2023,13, 912 22 of 56
similar direction and it cannot be excluded that nLDs assist cytosolic LDs in directing the
expression of lipid-metabolism-linked genes. Moreover, the association between nLDs and
heterochromatin seen in C. elegans under stress conditions could also indicate a functional
role of nLDs in aging-related gene silencing [
118
]. Indeed, aging in Drosophila is associated
with changes in heterochromatin structure yielding a repression of gene silencing, which
also activates the expression of transposable elements (TEs) residing in heterochromatic
areas of adipocyte nuclei in the fat body (as mentioned, the Drosophila equivalent to the
human liver) and brain tissue [
267
,
268
]. Antagonistic to this, dietary restriction counteracts
TE activation and extends lifespan. Similarly, mutation of the Drosophila gene Argonaute
2(Ago2), a regulator of TE silencing [
269
], results in enhanced TE expression, impaired
neuronal function, and reduced longevity [
270
]. Of note, the enhanced TE expression seen
in the aged fat body of old flies is accompanied by increased DNA damage and declined
levels of the nuclear-lamina protein lamin-B, with the depletion of lamin-B yielding a
similar phenotype (enhanced TE expression and DNA damage) in the fat tissue of larva
and young adults [
267
]. These findings shed light on the critical role of the nuclear lamina
in gene silencing, including TF expression and genome/DNA integrity surveillance in
lifespan control.
Interestingly, in early Drosophila development, binding of extranuclear histones to
cytosolic LDs enables the storage of histones required for chromatin organization, a task that
is conferred by the protein Jabba [
18
,
271
]. Extranuclear histone stores are specific for very
early, syncytial blastoderm stages of Drosophila embryogenesis that are marked by a rapid
series of consecutive nuclear divisions (without accompanying cell divisions, thus forming
a syncytium), with the Jabba-based recruitment of histones to LDs providing an adequate
extranuclear histone supply. Although this mode of Jabba-aided histone-to-LD binding
may be specific for early Drosophila embryogenesis, it is tempting to speculate that similar
histone–LD interactions play a role in aging-associated changes in chromatin organization
in adult flies as well as higher organisms. These may comprise chromatin remodeling
and histone methylation, which are both considered further hallmarks of aging [
74
]. In
Drosophila, repressive histone methylations contribute to heterochromatin stability and their
disruption results in lifespan shortening [
272
,
273
]. Histones found on LDs in Drosophila
include histones H2A and H2B, which both show an age-specific ubiquitination [
18
].
Reduction of the ubiquitinated form of H2A prolongs the flies’ lifespan [
274
]. Notably,
binding of H2A by LDs would naturally serve the same purpose. In H2B, the ubiquitination
is a prerequisite for the trimethylation of the histone H3K4 [
18
], which upon trimethylation
(H3K4me3) promotes the aging process in Drosophila [
275
]. Accordingly, the possible
binding of H2B by LDs would lead to a reduced amount of H3K4me3 and thus prolong the
lifespan of the flies.
Moreover, the aging-associated loss of histones (originally identified as a lifespan-
restricting determinant in yeast by Feser et al. [
276
]) and changes in the eu-/heterochromatin
ratio are linked to changed histone methylation patterns that affect gene expression and may
promote aging-associated DNA damage [
277
,
278
]. This fits well with the afore-mentioned
Argonaute 2 mutant phenotype in Drosophila marked by lamin depletion, enhanced TE
expression, and DNA damage. Similarly, down-regulation of lamin-B1 accelerates the
senescence of proliferating human fibroblast cells and has a profound impact on chromatin
structure and gene expression [
279
]. Finally, it has been demonstrated that recruitment of
the chromatin remodeler SNF2h (enhancing DNA accessibility in DNA repair) by sirtuin
(SIRT6), which deacetylates histones, protects human and mouse cells from genotoxic
damage [
280
]. On the other hand, SIRT6 also serves as a negative regulator of lipid
metabolism [
281
], with the EGF-dependent down-regulation of SIRT6 (FOXO3/SIRT6)
resulting in enhanced LD biogenesis in human colon cancer cells [
282
]. Taking into ac-
count the inverse nature of SIRT6 regulation on LD formation and the protective role of
SIRT6 on DNA/chromatin integrity, sirtuins (histone deacetylases) could play a critical
role in balancing LD abundance. However, this could also limit the cellular “lipid mass”,
including LDs, serving as substrate for lipid-peroxidation-derived genotoxic effects [
283
]

Biomolecules 2023,13, 912 23 of 56
under pro-oxidant conditions. Summarizing, these findings account for a distinct role of
cytosolic and nuclear LDs in chromatin and genome surveillance with a particular impact
on the aging process, for which the underlying mechanisms remain to be elucidated by
further investigation.
4.6. Intracellular Lipid Droplet Trafficking
Finally, additional findings point to intracellular LD trafficking as an important further
aspect in LD biology. In Drosophila, LD transport is mediated by the interaction of the motor
proteins kinesin and dynein with microtubules [284–287]. Involved in the coordination of
LD movement conferred by cytoskeletal interaction are proteins such as Bicaudal D (inter-
acting with dynein) [
288
] and perilipin-homolog LSD-2 [
289
], which physically interacts
with the gene product klarsicht (Klar is identical to the Drosophila gene marbles) [
286
,
290
].
Klarsicht mutants develop more or less normally, but due to disturbed LD transport show a
markedly reduced lipid/LD deposition in the blastoderm, yielding enhanced transparency
of the embryo (hence the name of the mutant). In addition, absence of Klar also leads
to mispositioning of photoreceptor nuclei in the developing eye and affects trafficking
of secretory vesicles in the salivary gland [
291
]. Three Klar isoforms (Klar
α
,
β
,
γ
) have
been described [
292
]. While no function is known for Klar
γ
, the isoform Klar
α
is in-
volved in linking cytosolic and nucleoplasmic proteins and, as a result, affects positioning
of photoreceptor nuclei. Although not shown, it cannot be excluded that Klar
α
and
LD-associated Klar
β
together with LD-resident Jabba participate in the recruitment of
extranuclear histones to LDs during early Drosophila embryogenesis, as discussed above.
Klar
β
, in a Klar
α
-analogous fashion, targets LDs for cytoskeletal interaction, which is
mediated by a distinct, C-terminal LD domain [
292
]. It is noteworthy that Klar via its LD
domain serves intracellular, microtubule-based transport of LDs, not only in embryonic but
also in adult Drosophila tissue, as well as in cultured insect S2 cells [
293
]. Summarizing, this
indicates that the association between LDs and the cytoskeletal/microtubule network may
enable distinct intracellular LD positioning mechanics in developmental, physiological,
and pathological contexts.
Taken together, Drosophila is a model organism with outstanding findings for LD
biology that foreshadow the integrative functionality of LDs in higher organisms con-
necting environmental, dietary, and hormonal inputs, and assisting in their translation to
metabolism and signaling cascades. The interaction with other cellular organelles, primar-
ily mitochondria and the ER, make LDs a useful and highly dynamic organelle apart from
lipid storage. However, it has to be admitted that the complexity of the LD interaction
network increases with the number of different cell types and tissues, although common
central regulatory pathways are conserved from yeast to the mammalian system (Figure 2).
In the following chapter dedicated to mammals, we show that LDs, as a kind of “Janus-
faced organelle”, essentially fulfill a cytoprotective function, but contribute to age-related
diseases if LD accumulation becomes inadequately excessive.
5. LDs in Human Disease
5.1. Caloric Restriction, Lifespan Control, and Age-Related Disease
It is a common thread in evolution that caloric restriction promotes health and prolongs
lifespan, which has been documented for a variety of organisms such as S. cerevisiae, proto-
zoans, rotifers, crustaceans, C. elegans and nematodes in general, Drosophila melanogaster,
and fish (Lebistes reticulates and Danio rerio) [
32
,
105
,
108
,
294
–
299
]. The underlying molecular
mechanisms seem to be similar or identical in all the organisms studied and converge
to common pathways—TGF-
β
signaling, IIS/IGF-1 signaling, mTOR, and stimulation of
autophagy—as illustrated in Figure 2. As outlined in this figure, these processes also share
the common motive that they stimulate or are otherwise associated with LD biosynthesis.
Aging research has identified several compounds that preserve health and extend lifespan
in different model organisms: resveratrol [
83
], rapamycin [
82
], spermidine [
212
], 2-deoxy-D-
glucose [
300
], curcumin [
301
], quercetin [
83
], metformin [
302
], and NAD
+
precursors [
303
].

Biomolecules 2023,13, 912 24 of 56
Many if not all of these substances have been shown to be caloric-restriction mimetics [
304
]
and, as discussed in the previous sections, many of these substances also stimulate LD
biogenesis. This provokes the central question of whether the life-prolonging effects of
caloric-restriction mimetics observed in non-mammalian animal model systems also apply
to mammals and humans, and whether these processes are also LD-driven.
In rodents, the life-prolonging effects of caloric restriction have been known for a long
time [
104
,
305
,
306
]. Over the last few years, data on the life-prolonging effect of caloric
restriction also became available for non-human primates, which now allows conclusions
to be drawn for humans. Since the 1980s, the effects of caloric restriction have been investi-
gated in the rhesus monkey Macaca mulatta by three different organizations (University of
Maryland, University of Wisconsin Madison, and the National Institute on Aging). After
settling some controversy over the study design, the participating organizations agreed in
concluding that caloric restriction has a positive effect on survival and aging-associated
diseases [
106
]. Since these studies did not investigate the involvement of LDs in the life-
prolonging effect seen in the rhesus monkeys, a distinct role of LDs in lifespan control can
only be conjectured from observations made in other model organisms. Indeed, various
findings in mammals, including humans, suggest a positive effect of caloric restriction on
LD biology. In humans, a distinction can be made between white, brown, and beige adipose
tissue. For a long time it was considered that brown adipose tissue is present only in the
newborn, contributing to the regulation of body temperature, and that it is rapidly lost after
birth. More recently, however, brown adipose tissue was also found in adults [
307
], being
involved in lipid and glucose oxidation as well as insulin-independent glucose uptake [
308
].
Based on this, it could be shown that activity of brown adipose tissue increases during
adolescence and rapidly ceases at higher age [
309
]. The implication of brown adipose tissue
in the aging process is also supported by the finding that the activity of brown adipose
tissue is significantly higher in long-lived than in short-lived animals [
308
]. Importantly,
the influence of caloric restriction leads to the “browning” of white adipose tissue, meaning
that adipocytes, instead of forming one large lipid droplet (as in white adipose tissue),
constitute many small LDs in brown adipose tissue [
310
]. This is accompanied by distinct
changes in the LD proteome [
311
]. Due to the long life of primates (in rhesus monkeys,
between 30–40 years), the study on caloric restriction referenced above is the only one of
its kind hitherto that proves the influence of an anti-aging strategy on primates/humans.
Since no data are available at present referring to LDs at the organismic level in aged indi-
viduals, we set the focus of this chapter to diseases whose prevalence and severity generally
increases with age, and are hence widely accepted as being age-associated [
312
], and for
which, in many cases, a contribution of LDs to the disease pattern have been reported.
In contrast to the previously discussed models such as C. elegans and D. melanogaster,
the role of LDs in human and mammalian aging is poorly defined. Representing a general
conceptual flaw for most pathologies that show elevated cellular LD levels, it is not clear
at present whether this LD accumulation is causal to the diseased state or rather is a
consequence of disease-related changes in lipid metabolism. In particular, evidence for
a regulatory, cytoprotective effect of increased LD levels, such as that indicated by the
lifespan-extending effects seen in other, less-complex model systems, is widely missing
in the mammalian/human system. Nevertheless, the research listed in Table 1provides
clear evidence that LD levels increase in several, if not all, age-related diseases (ARDs).
In ARDs, the irreversible cessation of cell proliferation that demarcates the progression
to cellular senescence is discussed as a major pathological criterion [
313
]. Essential to
this, the deregulation of cell-cycle-regulating genes is considered a hallmark of cellular
senescence which, surprisingly, also applies to lipid-related pathways [
314
]. Evidence is
accumulating that the deregulation of nutrient-sensing pathways, such as growth hormone
and IIS pathway [
315
–
317
], autophagy–regulatory mTOR and AMPK signaling, and the
histone deacetylase activity of sirtuins, play key roles in ARD development [
318
]. As
discussed above, exactly the same pathways have been shown to be important stimuli of
LD biogenesis in the other animal models of aging. Insulin signaling is also intrinsically

Biomolecules 2023,13, 912 25 of 56
tied to trafficking and storage of lipids in lipid droplets. With respect to the wide range of
ADR-associated processes, this review focuses on aging-associated aspects of LD biology
in several selected tissues.
Table 1. Age-related diseases associated with an increase in cellular LD numbers.
Disease Main Affected Cell Type/Tissue References
Alzheimer’s disease neurons, glia, myeloid cells, ependymal
cells, astrocytes [319–323]
Parkinson’s disease neurons, microglia [319,324,325]
Age-related macular
degeneration retinal pigment epithelium [326]
Stroke microglia [327]
Atherosclerosis Foam cells [328,329]
Cardiovascular disease myocardium [330,331]
Sarcopenia muscle cells [332,333]
Rheumatoid arthritis T-cells [334]
Chronic obstructive pulmonary disease (COPD) macrophages [327]
Periodontitis monocytes, macrophages [335,336]
Osteopenia osteoblasts, osteocytes [337,338]
Osteoarthritis chondrocytes, cartilage [339–341]
Diabetes β-cells [342,343]
Liver disease (NAFLD) 1parenchymal hepatocytes [344]2
Cancer several [343,345,346]
Senescence several [347–349]
1
Non-alcoholic fatty liver disease per se is characterized by progressive steatosis involving LD accumulation.
2Review of the aging aspect of NAFLD to HCC progression.
5.2. Bone Marrow Aging–Epigenetic Mechanisms
In both children and adults, bone mineral density is inversely correlated with bone
marrow fat abundance [
350
,
351
] and the aging process in bone marrow is characterized
by an expansion in marrow adipose tissue (MAT), which impairs bone stability, thereby
yielding an enhanced bone fracture risk [
352
]. Essential to this, non-differentiated bone
marrow stromal cells (BMSCs), amongst others, serve as progenitors for osteoblast and
adipocyte differentiation. BMSCs are marked by a low LD content but LD abundance
increases upon osteogenesis due to enhanced energy demands, and the blockade of LD
formation by Triacsin C (an inhibitor of fatty acyl CoA synthase) results in impaired os-
teogenic differentiation [
353
]. In line with this, LD-associated PLIN2 is generally expressed
in bone tissue with the highest levels seen in osteo-progenitor cells [
353
]. Increased serum
levels of fatty acids stimulate the adipocytic differentiation of bone marrow progenitor
cells [
354
], the MAT-resident adipocytes containing large LDs that serve as fatty acid and
adipokine reservoirs [
355
]. Of special relevance, MAT expansion may promote free-fatty-
acid-based lipotoxicity and negatively affect bone marrow osteoblast proliferation [
356
],
and the inhibition of adipocyte-derived fatty acid synthesis may protect osteoblasts from
the lipotoxic effect [357].
Epigenetic mechanisms may play a crucial role in bone marrow aging. Underlying this,
histone deacetylases (HDACs) such as sirtuins remove acetyl groups from lysine residues
of histone tails, which leads to chromatin condensation and thereby alters gene expression.
For example, activation of the murine class III HDAC sirtuin 1 (SIRT1), a nutritional sensor
responding to NAD
+
/NADH changes, directs mouse mesenchymal C3H10T1/2 cell lines
and primary rat bone marrow stromal cells towards enhanced osteoblastic and reduced

Biomolecules 2023,13, 912 26 of 56
adipocytic differentiation, while SIRT1 inhibition shows exactly the opposite effect [
358
].
It is also worth mentioning that LD-derived mono-unsaturated fatty acids are strong
activators of SIRT1 [
359
]. Moreover, a direct interaction has been shown between the
peroxisome proliferator-activated receptor
γ
2 (PPAR
γ
2), a key transcription factor for the
differentiation of progenitors into adipocytes, and Sirt1 through its catalytic core domain,
forming a stable transcription-inhibiting complex. Binding of this complex to the SIRT1
promoter represses SIRT1 transcription via a self-regulatory feedback loop [
360
]. This
report shows a decline in SIRT1 mRNA and protein levels in older compared to younger
human lung, heart, and fat tissues, thus indicating that the SIRT1/PPAR
γ
interaction is
a senescence-associated (epi)genetic mechanism [
360
]. In accordance with this, elevated
PPAR
γ
2 levels were also detected in aged compared to young bone marrow stromal
cells [
361
] and it is proposed that the SIRT1/PPAR
γ
2 negative feedback loop lowers
SIRT1 expression in bone marrow, and, as a result, promotes MAT expansion and the
aging process.
In addition, studies in mice have shown that deletion of another HDAC—Hdac3—leads
to lipid accumulation in osteochondrocyte progenitor cells and promotes MAT expansion in
young mice [
337
]. Compared to the wild type, the Hdac3 knockout also leads to a substan-
tial shift in LD/lipid storage-associated Plin1 and Fsp27/Cidec (Fspe27, with fat-specific
protein 27 belonging to the family of death-inducing DFF45-like effectors (CIDE) [
362
]),
and a minor but still significant increase in lipolysis-associated lipases (Pnpla2 and Lipe).
Further transcriptome analysis in Hdac3 knockout mice revealed a highly abundant expres-
sion of 11b-hydroxysteroid dehydrogenase type 1 (Hsd11b1), a gene encoding an enzyme
involved in the activation of intracellular glucocorticoids participating in glucocorticoid
receptor-based signaling. Inhibition of Hsd11b1 by carbenoxolone resulted in reduced
expression of Plin1 and Cidec in Hdac3-deficient cells, which identifies Hdac3 as a crucial
regulator of glucocorticoid-induced LD formation in osteoprogenitor cells [337].
5.3. Lipid Droplets in Neurodegeneration
Neurodegenerative diseases such as Morbus Alzheimer and Morbus Parkinson rep-
resent another highly prevalent complex of age-related pathologies. Aging of the central
nervous system is associated with progressive myelin degeneration at a reduced myelin
renewal [
363
], overall loss of total brain volume [
364
], glia activation (microglia and astro-
cytes) and cilia loss in ependymal cells [
365
], and changes in neuron morphology leading
to neuron dysfunction [
366
]. Lipid homeostasis is of central importance to the functionality
of the nervous system since neuron function is considerably impaired by the accumulation
of fatty acids that promotes ER stress, lipotoxicity, and mitochondrial damage (recently
reviewed in [
319
]). Notably, the lipid composition of the normal human brain is about
60% by dry weight, which ranks directly after adipose tissue in terms of the tissues with
the highest fat fraction, and the brain fat content varies markedly between different brain
areas, being highest in myelin (78–81% of the dry weight) and lowest in grey matter
(36–40%) [
367
]. Triacylglycerol levels are low in neurons, probably due to the constant lipid
turnover generating the phospholipid mass required for cell membrane maintenance [
368
].
Although only sparse evidence exists for the
in vivo
LD formation in neurons, the LD
content of neurons increases under “lipid stress” arising from fatty acid treatment [
369
,
370
].
In addition, LD numbers are raised upon expression of mutant huntingtin protein [
371
]. In
the aged brain, LD accumulation has been shown in neurons, microglia, astrocytes, and
ependymal cells [
372
,
373
]. ROS-induced LPO, considered a hallmark of neurodegenera-
tion, serves as a driving force since, similar to the case in Drosophila, LPO caused by ROS
originating from mitochondrial dysfunction stimulates LD biogenesis by JNK-mediated
responses, which precedes the neurodegenerative process in the mammalian system [
46
].
Astrocytes are important regulators of oxidative stress adaptation in the brain by provid-
ing homeostatic, antioxidant support for neurons. This macroglial cell type shows an
enhanced resistance towards oxidative stress regimens [
374
], which might be attributable
to its well-equipped antioxidant defenses such as the glutathione redox system, manganese

Biomolecules 2023,13, 912 27 of 56
superoxide dismutase, and catalase [
375
]. Intriguingly, it has been shown that peroxidized
lipids (Lox) formed by ROS/LPO in neurons bind to apolipoproteins (especially ApoE)
and these ApoE/Lox complexes can be shuttled from neurons to astrocytes, where they are
decomposed by lysosomal processing and the liberated fatty acids are stored transiently in
LDs [
370
]. Terminally, the “imported”, LD-bound Lox are oxidized in mitochondria via
β
-oxidation, which stimulates ROS formation; however, due to the enhanced antioxidant
defense, astrocytes may successfully prevent substantial ROS build-up. In good accordance
with the findings of Schroeter et al. [
375
], Ioannu et al. [
370
] identified the up-regulated
expression of several genes associated with antioxidant defenses (Gpx8, glutathione peroxi-
dase 8; superoxide dismutase Sod1 and Sod3; catalase and fatty acid transporters FabP5
and FabP7) in cultured, LD-rich astrocytes. This emphasizes the integration of LDs in
enhanced astrocytic antioxidant defenses that confer particular antioxidant robustness
needed in these macroglial cells to serve as a “detoxifying recipient” for neuron-derived
LPO products. Hence, it appears likely that disturbances of this neuron-to-astrocyte lipid
transfer and LD storage detoxification mechanism will enhance the risk of neuron dam-
age and the development of Alzheimer’s disease (AD). In humans, three APO-E alleles
have been described: APOE2, APOE3, and APOE4. Compared to APOE2 homozygotes,
individuals bearing the APOE4 allele either in a heterozygous or homozygous genotype
show a 9–15-fold increased risk of acquiring AD, rendering the APOE4 allele as one of the
main risk factors for AD [
376
,
377
]. In fact, it was shown that the APOE4 allele dampens the
“neuron to glia” lipid transfer, thus promoting neurodegeneration [
378
]. With respect to
this, it is tempting to speculate that, downstream of ApoE4/Lox-shuttling, the inability of
astrocytes to store Lox/lipid peroxides in LDs also promotes the risk of developing AD.
LDs and
α
-synuclein. In Parkinson’s disease (PD), the second most common neu-
rodegenerative disease after AD, LDs also move into the focus of research. Central to PD
pathogenesis is the loss of dopaminergic neurons in the substantia nigra pars compacta
accompanied by the appearance of Lewy bodies [
379
]. Lewy bodies are insoluble aggre-
gates of misfolded proteins, of which
α
-synuclein comprises the main constituent [
379
,
380
].
Oligomers of
α
-synuclein are neurotoxic to themselves and are considered as the main
drivers of neurodegeneration [
381
], with mutations of
α
-synuclein dramatically increasing
the prevalence of PD [
382
]. It was shown that
α
-synuclein forms di- and trimers that
accumulate at the surface of LDs in human embryonic kidney cells as well as hippocam-
pal neurons treated with high concentrations of oleic acid [
325
]. Taking into account the
observations made in S. cerevisiae, C. elegans, and Drosophila, this suggests the existence
of LD-based detoxification mechanisms for
α
-synuclein aggregates in higher organisms,
which hypothetically may provide neuroprotection. Whether this holds true and the extent
to which it may play a role in PD pathogenesis remains to be addressed by further research.
LD accumulation in Huntington’s disease. Apart from AD and PD, accumulation
of LDs is seen in several other neurological and neurodegenerative disorders, including
Huntington’s disease and amyotrophic lateral sclerosis (ALS), as reviewed recently by
Islyme et al. [
383
]. However, the authors also leave open the question of whether LD
accumulation mitigates or promotes the progression of neurodegenerative disorders. This
emphasizes the ambiguity of LD biogenesis in the pathogenic context. LD accumulation
may enable dynamic lipid storage, and, as a result, act as a potential substrate for LPO,
in one pathological condition, but opposite to this, in another diseased state, resemble a
sink for the “safe” sequestration of LPO-derived compounds including peroxidized fatty
acids as well as aggregates of potentially harmful proteins such as
α
-synuclein oligomers.
It is reasonable to consider that the direction of the LD-based response is dependent on the
influence of additional factors. For instance, in Huntington’s disease, the mutation-based
excessive N-terminal poly-glutamylation of the protein huntingtin (polyG-Htt) is causative
of neuronal cell death, which to a substantial degree is due to the disturbed interaction
between poly-glutamylated huntingtin and the cytoskeleton [
384
]. Huntingtin is involved
in several cellular transport processes and, in concert with the microtubule network in
particular, participates in the axonal transport of organelles and neurotransmitters in neu-

Biomolecules 2023,13, 912 28 of 56
rons [
385
]. Interestingly, in a yeast model of poly-glutamylation, polyG-Htt aggregation
leads to the formation of inclusion bodies and cell death, the degree of which correlates with
an aberrant LD morphology that is indicative of a disturbed TAG storage function [
386
].
Taking into consideration the role of cytoskeletal alterations in neurodegenerative dis-
ease [
387
], an inadequate binding of protein aggregates to LDs could hypothetically also
result in a disturbance of cytoskeleton-based intracellular LD/lipid trafficking, for instance,
by impairing the afore-mentioned interaction of LDs with motor proteins (see Section 4),
and, as a result, aggravate disease progression. Therefore, although the clearance of poten-
tially dangerous protein aggregates by LDs may be considered beneficial to cell integrity,
evaluating the impact of this LD-based mechanism on disease outcome deserves a rather
holistic approach integrating a multiplicity of interlinked accessory factors.
5.4. Lipid Droplets in Metabolic Disease
Intimately associated with the essential, lipid-metabolism-linked function of LDs, the
metabolic process itself represents a critical element of aging. This is indicated by the
fact that the risk for metabolic diseases such as the metabolic syndrome, type II diabetes,
non-alcoholic fatty liver disease/steatohepatitis (NASH), and cardiovascular diseases and
atherosclerosis, is increasing with age [
388
]. For example, in the pathogenesis of type II
diabetes, the development of insulin resistance represents an early, critical issue leading
to the disturbance of glucose homeostasis. As a pathophysiological response, insulin
production by the
β
-cells of the pancreatic Langerhans islets becomes elevated, which
partially compensates for the incremental insulin resistance. However, with progression of
the diabetic condition, the
β
-cell mass declines and, with this, insulin production ceases.
Nutrition-derived lipid stress arising from a high-fat diet is under discussion as a main
driver of type II diabetes development, with LDs playing a central role in
β
-cell lipid
management [
342
]. In fact, it was shown that knockdown of perilipin 2 (PLIN2), an
essential LD scaffold protein, resulted in reduced insulin production by
β
-cells, whereas
PLIN2 overexpression boosted insulin secretion. Moreover,
β
-cells devoid of LDs are prone
to ER stress, which results in an impairment of
β
-cell functionality [
343
]. This accounts for
a critical role of LDs in protecting
β
-cells from the toxic effects of lipids, and, as a result,
acting as an antagonist of type II diabetes progression.
LD accumulation in NASH. Insulin resistance is closely associated with obesity marked
by the enhanced accumulation of LDs in epithelial cells and other non-adipose tissues,
which establishes a pro-inflammatory microenvironment in the affected tissues as com-
monly found in metabolic diseases. This also applies to the development of NASH, the
inflammatory type of non-alcoholic fatty liver disease (NAFLD) [
389
]. NASH, which is
closely associated with insulin resistance, represents a well-studied liver pathology emerg-
ing from chronic fat-rich alimentation. NASH is characterized by the marked accumulation
of triglycerides in liver epithelial cells (i.e., parenchymal hepatocytes) causing liver steatosis
and the accompanying, chronic inflammation that promotes fibrotic/cirrhotic remodeling
of liver tissue (recently reviewed in [
390
]). Of particular relevance, the development of
NASH is accompanied by several processes including mitochondrial dysfunction, ER stress,
and enhanced ROS formation, as well as tissue-specific changes, primarily the activation of
hepatic stellate cells (HSCs), which promotes the inflammatory process and liver fibrosis
upon trans-differentiation of activated HSCs into the extracellular matrix (ECM) producing
myofibroblast-like cells [
344
]. In the “multiple hit pathogenesis” of NASH, toxic lipids play an
essential role, affecting different liver cell populations, in particular parenchymal hepato-
cytes, HSCs/myofibroblast-like cells, and Kupffer cells (i.e., liver macrophages) in different
ways [
391
]. Contrasting with the well-defined general understanding of NASH develop-
ment and its clinical manifestation, little is known about age-associated alterations in lipid
metabolism and LD biogenesis in NAFLD/NASH progression. Nevertheless, ROS and the
ROS-mediated senescence of hepatic cells may also play a pivotal role in NAFLD/NASH
pathogenesis. For instance, it was shown that liver steatosis is promoted by a decline in

Biomolecules 2023,13, 912 29 of 56
mitochondrial fatty acid metabolism in senescent hepatocytes, an effect that was abolished
by the antioxidant, lipid peroxidation inhibitory flavonoid quercetin [217,392].
Role of ROS. Several investigations account for a distinct role of the ECM-associated
matrix protein CCN1 (central communication network factor 1; formerly termed Cyr61,
cysteine-rich protein 61 [
393
]) in NASH progression. CCN1 plays an important role in
wound-repair-associated ECM remodeling via binding to integrin
αVβ
3 and
αVβ
5 of
epithelial cells and myofibroblast integrin
α
6
β
1 [
394
]. In line with this and accounting
for a role in liver fibrosis, an increased expression of CCN1 has been shown in the liver
of NASH patients [
395
], in hepatocytes of the human cirrhotic liver, and as a reaction to
liver injury [
396
]. Furthermore, it has been shown that CCN1 stimulates hepatic steatosis in
obese mice and promotes LD accumulation in hepatocytes treated with free fatty acids [
395
].
Of considerable relevance, CCN1 expression is enhanced by ROS. This was demonstrated in
skin fibroblasts exposed to hydrogen peroxide, which resulted in the c-jun/AP1-dependent
up-regulation of CCN1 expression yielding a repression of collagen synthesis and fibroblast
senescence [
397
,
398
]. Similarly, the overexpression of CCN1 also lowers the production of
collagen type1
α
1 (col1
α
1) in HSCs [
399
]. Moreover, CCN1, by acting via integrin
α
6
β
1,
shifts ROS/RAC1-dependent NOX1 (NADPH oxidase 1) activity and promotes senescence
of HSCs as well as myofibroblasts, as a result conferring an anti-fibrotic response, and
in addition stimulates liver-regeneration-associated signaling via IL-6 (interleukin-6) and
CXCR2 (chemokine receptor 2) ligands [396,400].
These observations suggest a seemingly ambiguous involvement of CCN1 in NASH:
promoting steatosis by LD accumulation in parenchymal hepatocytes, but counteracting
NASH-associated fibrosis by mediating senescence, in non-parenchymal HSCs and my-
ofibroblasts. It is noteworthy that in HSCs, the overexpression of CCN1 is also able to
trigger ER stress and UPR due to the high abundance of CCN1 protein, which renders
these cells susceptible to apoptotic cell death [
399
]. Hence, elevated CCN1 levels may exert
a particular challenge for cell integrity in HSCs. Importantly, parenchymal hepatocytes,
and not HSCs or Kupffer cells, represent the major hepatic source of CCN1. This was
shown by employing CCl
4
, a hepatotoxic compound that generates free radicals upon
cytochrome P450-based metabolization and shifts CCN1 expression only in parenchymal
hepatocytes [
396
]. It has to be mentioned that CCN1 null mutant mice reveal a normal
hepatic function in the absence of CCl
4
, which suggests a specific association of hepatocytic
CCN1 expression with stress/ROS-mediated conditions, such as existing in liver injury and
inflammation. This notion is supported by the further observation that CCN1 expression
in parenchymal hepatocytes is also up-regulated by the pro-inflammatory cytokine TNF
α
in a ROS-dependent mode [
401
]. Conversely, the before-mentioned stimulation of LD bio-
genesis upon CCN1 overexpression in primary mouse hepatocytes is accompanied by an
enhanced expression of TNF
α
[
42
]. TNF
α
, acting via the TNF
α
receptor (TNFR), stimulates
mitochondrial ROS generation via JNK signaling and downstream ER stress, which leads
to the activation of ATF6 (activating transcription factor 6) and elF2
α
(eukaryotic initiation
factor 2
α
). In turn, ATF6 and elF2
α
transduce the signal to nuclear transcriptional control
via C/EBP (CCAAT/enhancer-binding protein
α
) homologous protein (CHOP), which may
stimulate ER-stress-dependent apoptosis in hepatocytes [402].
Taken together, these findings account for the existence of an autocrine amplification
loop in hepatocytes, established by CCN1 and TNF
α
/TNFR under pro-oxidant conditions
that enables the secretion of CCN1 to the extracellular space and drives activated HSCs
and myofibroblasts towards senescence, eventually counteracting the HSC/myofibroblast-
driven fibrotic process. Indeed, an inflammation-associated feed-forward loop of cytokine
secretion including TNF
α
has been discussed recently [
402
]. In extension to this, CCN1-
stimulated LD biogenesis could play a central, albeit ambiguous, role in this regulatory
network. With respect to the causative role of increased CCN1 and TNF
α
levels in ER
stress, UPR, and apoptosis in HSCs, as well as hepatocytes, it is appropriate to consider
that hepatocytes, as the main hepatic source for CCN1 under stress conditions, need to
be protected from the cytotoxic potential established by the self-amplifying CCN1–TNF
α

Biomolecules 2023,13, 912 30 of 56
circuit. With respect to this, the finding of Ju et al. revealed that overexpression of CCN1
in hepatocytes leads to both (i) the up-regulation of lipid-metabolism-associated genes (ii)
and the up-regulation of sirtuins (Sirt 1, 2 and 3), Nrf1, BMP2 (a member of the TGF-
β
superfamily), and AMP kinases [
395
]. This connects the CCN1–TNF
α
circuit to hepatic LD
biogenesis, and by this steatosis to most of the LD-associated signaling pathways discussed
above, which are capable of exerting a cytoprotective function in cells under oxidative
stress. Hence, it is tempting to speculate that enhanced LD biogenesis allows hepatocytes
to synthesize CCN1 in potentially cytotoxic amounts, which are needed for counteracting
inflammation-driven fibrosis by the paracrine induction of HCS/myofibroblast senescence.
Obviously, if this holds true, such a mechanism would assign a yet ambiguous context to LD
biogenesis in NAFLD/NASH (as well as alcoholic AFLD): slowing disease progression in a
paracrine mode via the CCN1-mediated deceleration of inflammation-associated fibrosis at
the cost of promoting steatosis progression via an autocrine amplification loop.
The stimulation of NOX1-based ROS (superoxide) production following CCN1 sig-
naling via integrin
α
6
β
1 in HSCs represents a further critical element of such auto- and
paracrine regulatory networks. Concerning this, Kim et al. proposed that HSC senescence
is caused by CCN1/NOX1-mediated ROS leading to genotoxic damage, and p53 and
p16/pRb–dependent, senescence-related responses [
396
]. Moreover, similar to LD accumu-
lation mediated by CCN1 in hepatocytes [
395
], Long et al. showed that the up-regulation
of NOX1 also stimulates lipid-metabolism-related gene expression and LD accumula-
tion in mouse hepatocytes. and also leads to the up-regulation of ER-stress-associated
genes ATF6 and eIF2, effects which were antagonized by the antioxidant N-acetylcysteine
(NAC) [
403
]. This strengthens the assumptions made above regarding a protective role of
LD biogenesis counteracting the pro-oxidant effects on the CCN1–TNF
α
circuit, as well as
altered NOX1 activity in hepatocytes under conditions of inflammation. It is noteworthy to
emphasize that in the experiments conducted by Long et al., NOX1 overexpression was
accomplished in hepatocytes via knockdown of the transcription factor hepatocyte nuclear
factor 1
β
(HNF1
β
), which identifies HNF1
β
as a negative regulator of NOX1, and, as a
result, a suppressor of both NOX1-mediated superoxide formation and LD biogenesis in
hepatocytes [
403
]. Interestingly, steatotic livers of obese mice show a reduced expression
of HNF1
β
, and treatment of mouse hepatocytes with palmitic acid also lowers HNF1
β
expression, an effect that is suppressed by NAC [
403
]. In addition, the HNF1
β
knockdown
also stimulated insulin resistance in hepatocytes, which was also ameliorated by NAC.
These observations account for a further feedback mechanism in hepatocytes, leading to
an elevated LD accumulation that is driven by the lipid/ROS-based down-regulation of
HNF1
β
, and resulting in NOX1-mediated LD biogenesis and causing increased insulin
resistance. With respect to this, HNF1
β
connects NAFLD with diabetes, which is under-
lined by the finding that mutated HNF1
β
alleles are associated with diabetes type MODY5
(maturity-onset diabetes of the young) [
404
] as well as type-II diabetes [
405
], suggesting
that HNF1βactivity mitigates insulin resistance, at least in these pathologies.
Lipid droplets, lipophagy, and hepatic lipid homeostasis. The hepatic lipid flux is
marked by diurnal oscillations of fed and fasted states reflected by LD catabolism (fed
state) and LD storage (fasted state), with the hepatocyte LD lifecycle playing a pivotal role
in systemic lipid homeostasis. Regarding this, the blood insulin concentration is critical by
linking systemic fed/fasted states to oscillating high (fed) and low (fasted) insulin signaling.
Among several physiologic contexts, this nutrition-dependent systemic insulin dynamics
also affects cellular LD dynamics, in particular intracellular LD trafficking. Similar to the
case of Drosophila (see Section 4), LDs may also become connected to the cytoskeleton
in primary hepatocytes where LD binding of the motor protein kinesin-1 mediates their
transport along microtubules, a process that is regulated by insulin signaling [
406
] and
serves the delivery of LDs to the smooth ER (sER) for VLDL (very-low-density lipoprotein)
production [
407
]. Essential to this, insulin signaling enhances the GDP-dependent binding
of GTPase ADP-ribosylation factor 1 (ARF1) to LDs, rendering them “reactive” [
408
], the
bound ARF1 in turn recruiting phospholipase-D1 (PLD1), which generates phosphatic acid

Biomolecules 2023,13, 912 31 of 56
(PA), and, as a result, shifts the PA content of reactive LDs [
409
]. As a consequence, via
binding to PA, these LDs recruit kinesin-1, which terminally mediates LD shuttling to the
sER. Therefore, in the fed state, high systemic insulin levels will promote LD–sER shuttling
and fuel VLDL production as well as secretion by hepatocytes, while the postprandial, low-
blood insulin levels characteristic of the fasted state will antagonize the shuttling process
and thus limit VLDL synthesis and secretion. Notably, starvation conditions markedly
enhance the hepatic clearance of adipose-tissue-derived lipids from the circulation, which
causes a substantial shift in the hepatocyte LD content. Hence, the down-regulation of LD
trafficking entailed by low insulin signaling serves as a “bottleneck” for VLDL production
in LD-rich hepatocytes, limiting VLDL under fasted conditions. This puts emphasis on
the role of LDs participating in the physiological regulation of systemic lipid homeostasis
serving as a hormonally controlled, dynamic lipid buffer in the liver.
At the cellular level, LDs can be selectively degraded by autophagy, a process termed
(macro)lipophagy, which sequesters cytosolic LDs for autophagolysosomal digest and
is pivotal to lipid metabolism [
410
,
411
]. In addition to its role in mobilizing fatty acids
from cellular LD-based lipid stores, lipophagy represents a pivotal “guardian” of cellular
LD abundance. This holds particularly true when LD biogenesis is substantially stimu-
lated in hepatocytes in response to an enhanced clearance of lipids from the blood stream
when facing the risk of systemic lipotoxicity arising from an excess of circulating free
fatty acids. Emphasis on this is provided by experiments showing that lipid treatment
of cultured hepatocytes stimulates lipophagy while lipophagy inhibition by the macroau-
tophagy inhibitor 3-methyladenine shifts the number of LDs in hepatocytes even under
normal culture conditions [
410
]. Moreover, lipophagy is also enhanced in the mouse liver
under starvation conditions that stimulate LD biogenesis. Importantly, these experiments
revealed further that enhanced exogenous lipid supply such as that provoked by a fat-rich
diet negatively affects LD breakdown by lipophagy. In good agreement with this, it was
reported recently for a mouse model of obesity that lipophagy declines upon feeding a
high-fat diet, resulting in liver steatosis [
412
]. Of special relevance to lipotoxic side effects,
the observed drop in autophagic/lipophagic efficiency was accompanied by the enhanced
accumulation of HNE-modified proteins. This connects lipophagy with reparative au-
tophagy (i.e., detoxification of the aggregated modified proteins), and, as a result, LD
abundance, with the critical interference existing between LPO and proteostasis. Hence,
it is not surprising that a reduced lipophagic flux plays an important pathogenic role
in NAFLD and other lipid-metabolism-associated diseases such as atherosclerosis [
413
],
and may affect the aging-associated transition from NAFLD into primary hepatocellular
carcinoma (HCC) [344].
LD accumulation, oxidative stress, and cell death. Finally, the excessive cytosolic accu-
mulation of fatty acids seen under “hyperlipidemic” states such as obesity, NAFLD/NASH,
and diabetes may lead to an elevated susceptibility to lipotoxicity-induced cell death via
apoptosis. The specific term lipoapoptosis was coined for apoptotic cell death stimulated
by fatty acid derivatives such as ceramide [
414
], and as a terminal issue in lipotoxic settings,
lipoapoptosis is of particular relevance to several lipid-associated pathologies including
NAFLD [
415
], as well as others such as vascular and cardio-metabolic diseases [
416
], which
are discussed below. In addition, ROS-stimulated LPO yields further metabolites such as
HNE, which interferes with anti-apoptotic and cell-proliferation-associated intracellular
signaling [
417
], but is also capable of inducing apoptosis per se [
418
]. Moreover, the concept
of ferroptosis, which has gained substantial interest over the last decade, also represents, in
principle, an LPO-dependent mode of cell death. In ferroptosis, LPO is initiated by iron-
derived
•
OH radicals at inadequate antioxidant defenses (i.e., weakening of the GSH/GSSG
redox system due to GSH-peroxidase 4 deficiency) leading to a non-apoptotic, necrotic
mode of cell death [
419
]. Taking into consideration that hepatocytes represent an iron-rich
cell type serving systemic iron buffering, ferroptosis represents a considerable issue in
NAFLD/NASH that also occurs with high levels of PUFAs [
420
]. In addition, it should
not be overlooked that ferroptosis may not only lead to hepatocyte loss, but as a necrotic

Biomolecules 2023,13, 912 32 of 56
mode of cell death will also aggravate the pro-inflammatory condition. Therefore, lipid
stress arising from an inappropriate lipid accumulation will affect many cellular targets and
pathways, among which LDs are of central relevance, either fueling lipid (per)oxidation or
attenuating lipotoxicity by aiding cellular lipid detoxification and interfering with cytotoxic
responses such as lipoapoptosis and possibly also other lethal outcomes such as ferroptosis.
5.5. Lipid Droplets in Vascular Disease
In hepatic Kupffer cells residing in the liver sinusoids, the augmented uptake of
cholesterol and free fatty acids enhances the development of a lipid-rich macrophage phe-
notype [
421
], characterized as foam cells, and the aggregation of such lipid-rich Kupffer
cells contributes to NAFLD/NASH-associated lipogranulomas that are built from inflam-
matory cells, ECM (collagen), and LDs [
422
,
423
]. Of particular pathological relevance,
the conversion of macrophages into foam cells represents a major issue in atherosclerosis
development (atherogenesis), since the “foamy” macrophages build up atherosclerotic
plaques in arterial walls, leading to inflammation and progressive damage to the ves-
sel wall (reviewed in [
328
]). Underlying this is the oxidation of low-density lipoprotein
(LDL) bound to proteoglycans of the extracellular matrix and the vessel endothelium,
which yields oxidized LDL (oxLDL), the oxLDL in turn triggering the release of monocyte
chemoattractant protein (MCP-1) by vascular endothelial cells and vascular smooth muscle
cells (VSMCs). The attracted monocytes migrate to the arterial wall and differentiate into
macrophages that endocytose the oxLDL particles via scavenger receptor A (SR-A) and
CD36 [
424
], although alternative uptake mechanisms may exist [
425
]. Upon lysosomal
processing of the oxLDL particles, the free fatty acids and cholesterol molecules are re-
leased to the cytoplasm, where they are either stored in LDs [
426
,
427
] or are released via
high-density lipoprotein (HDL) [
428
]. The enhanced uptake of oxLDL by macrophages
will nourish the accumulation of cholesterol-rich LDs, and, as a result, stimulate foam cell
and plaque formation [
328
]. It should not be overlooked that LDs are also essential to
reverse cholesterol transport (RCT), a process by which cholesterol sequestered from the
circulation is stored transiently in LDs, from which it can be liberated via lipophagy and be
re-released from the cell, for instance, to the bile for fecal excretion. Although RCT can be
accomplished by several cell types, macrophage-associated RCT is considered causal to the
atherosclerotic process [429].
LDs and cholesterol homeostasis. From this, it becomes clear that the ability of LDs
to store cholesterol is essential to cellular cholesterol homeostasis and the protection from
cholesterol lipotoxicity. Essential to this is the esterification of free, unesterified cholesterol
by Acetyl-coenzyme A cholesterol O-acyltransferase-1 (ACAT1) as a prerequisite for the
incorporation of the cholesteryl-esters into LDs [
430
]. In line with the protective role of LDs,
stimulation of ACAT1 is essential for proper cellular cholesterol management and, by facili-
tating LD-based cholesterol clearance, counteracts cholesterol toxicity [
431
]. In addition,
oxysterol-binding protein-related proteins ORP2, ORP5, and ORP8 can stimulate LD biogen-
esis and can bind oxysterols such as 25-hydroxycholesterol and 7-ketocholesterol, as well
as cholesterol itself, to LDs [
432
,
433
]. Considering the toxic effects of oxysterols, oxysterol
binding to LDs clearly represents a cytoprotective function of LDs. This holds particularly
true for 7-ketocholesterol, which accumulates in foam cells (reviewed in [
434
]) and is known
as a stimulator of oxiapoptophagy, a distinct mode of oxysterol/oxidative-stress-associated
cell death involving apoptosis and autophagy with particular pathogenic relevance to
age-related diseases including atherosclerosis [
435
–
439
]. The cholesteryl ester-driven bio-
genesis of LDs in vascular macrophages is considered causal to foam cell development [
429
],
which likewise also holds true for oxysterol binding to LDs, and both processes represent
driving forces of atherogenesis. As stated by Lee-Rueckert et al. [
440
], the development
of foam cells may be accompanied by the reduced expression of pro-inflammatory genes
(characteristic of the activated M1 macrophage phenotype) converting the phenotype into
an anti-inflammatory one (i.e., activated lipid-rich M2 macrophages [
441
], leading to the
concept that foam cell development represents an anti-atherogenic effect [
440
]. Hence,

Biomolecules 2023,13, 912 33 of 56
LD accumulation in atherosclerosis serves as a further example of the ambiguous role
of LDs in pathophysiological settings, as mentioned above for NAFLD: protection from
acute lipotoxicity, thus aiding cell survival at the cost of promoting a chronic process
such as atherosclerosis and liver fibrosis/cirrhosis. Interestingly, Lathe et al. followed a
similar concept in discussing the effects of the pathogen (virus)-induced stimulation of
25-hydroxycholesterol, which via ACAT1 esterification can also bind to LDs, and con-
tributes to both atherosclerosis as well as Alzheimer’s disease, postulating that
25-hydroxycholesterol protects from “infectious agents at the expense of longer-term pathol-
ogy” [
442
]. Emphasizing the ambivalent role of LD formation in foam cell development and
the atherogenic context, plaque formation is accompanied by a decline in the lipophagic
flux, which limits LD breakdown, and, as a result, excessive liberation of cholesterol [
443
],
but aggravates atherogenesis due to the continuous stimulation of foam cell formation
driven by LD accumulation. Finally, it appears noteworthy that VSMCs may translocate to
the arterial intima in the course of atherosclerosis progression and transdifferentiate into a
macrophage-like foam cell phenotype, revealing an enhanced oxLDL content, although
containing fewer LDs and showing a reduced lipophagic flux compared to macrophage-
derived foam cells [
444
,
445
]. Nevertheless, VSMC-derived foam cells can comprise about
50% of the foam cell content seen in human atherosclerotic plaques [
444
], and thus represent
an atherogenesis-associated cell population of considerable interest.
Finally, LDs may assist macrophage integrity not only by the control of lipid balance
and oxLDL/oxysterol sequestration, but also by aiding the “clearance” of other aging
and stress-associated compounds, especially protein aggregates. This is indicated by a
recent investigation that demonstrates the aging-dependent binding of protein aggregates
to LDs in mouse intestinal tissue, supposedly followed by the terminal degradation of the
critical matter via lipophagy [
135
]. It is conceivable that a similar, LD-based cytoprotective
mechanism is involved in aging and lifespan control of other organisms such as C. elegans
and Drosophila, and also likely in yeast, considering the binding of IBs to LDs as discussed
in this review.
5.6. LD Accumulation in Cardiomyocytes: Role of PPARs
As in other tissues, LDs also play a dual role in the cardiac system. To overcome
the enhanced energy demand of cardiomyocytes, long-chain fatty acids (LCFAs) such as
palmitate and oleate (due to their higher energy yield per carbon molecule as compared to
glucose) are the primary fuel needed for ATP synthesis [
330
]. Subsequent to esterification by
acyl-coenzyme A synthetase (CoA), these CoA-fatty acyls are further esterified to a glycerol
backbone and stored as TAGs in LDs. Upon LD lipolysis and lipase-mediated TAG hydrolysis,
the liberated fatty acids will fuel mitochondrial
β
-oxidation or serve as ligands for the nuclear
peroxisome proliferator-activated receptor
α
(PPAR
α
), a transcription factor that is central to
the control of intracellular TAG turnover and fatty acid metabolism [
446
,
447
]. It is noteworthy
that another PPAR species, PPAR
γ
, stimulates lipid uptake and LD biogenesis in cardiac
tissue and confers protection of cardiomyocytes from ROS-mediated damage via regulating
the expression of the Sod2 gene, encoding manganese superoxide dismutase [
448
]. Hence,
PPARs represent important determinants of cardiac LD turnover, which is central to cardiac
lipid management and protects the heart from organ dysfunction caused by lipotoxic-
ity [
330
]. Interestingly, it has been shown that PPAR
α
together with mTOR also regulate
a reciprocal mode of LD biogenesis and mTORC1-containing stress granule formation
in lipid-stressed HEK239T and SH-SY5Y cells [
449
]. It cannot be excluded that this also
applies to heart tissue and this interconnects cardiac LD turnover with autophagy via
mTOR/PPAR signaling.
Role of PLIN5 in cardiac disease. Of particular pathologic relevance, heart failure
in obesity and diabetes mellitus is associated with hyperlipidemia resulting in lipid ac-
cumulation and an expansion of the myocardial LD content [
450
]. In diabetic heart dis-
ease, particular attention has been paid to the role of the LD-associated protein perilipin
5 (PLIN5) (recently reviewed in [
451
]. Reflecting the context-dependent role of LDs in lipid

Biomolecules 2023,13, 912 34 of 56
homeostasis, PLIN5, under normal conditions, inhibits lipolysis via binding comparative
gene identification-58 (CGI-58), which otherwise binds triglyceride lipase, but under stress
conditions (e.g. fasting, exercise) stimulates lipolysis as a result of phosphorylation by PKA
(protein kinase A), leading to the release of CGI-58 from PLIN5, which in turn activates
triglyceride lipase [
452
]. Connected with this, cells with a high oxidative capacity show an
enhanced expression of PLIN5, as holds especially true for cardiomyocytes, where PLIN5
is responsible for the tethering of LDs to mitochondria [
453
]. Compared to healthy control
donors, expression of PLIN5 is reduced in samples drawn from patients with heart failure,
showing a decline in direct LD-mitochondria contacts and reduced fatty acid usage for
energy supply [
454
]. On the other hand, by inhibiting triglyceride lipase, PLIN5 supports
the sequestration of TAG by LDs, which will limit fatty acid availability, and, as a result,
protect the heart from lipotoxicity; however, dysregulation of this process upon PLIN5
overexpression will promote cardiac steatosis and hypertrophy [
455
,
456
]. Conversely,
LDs are absent from myocardial tissue in PLIN5 knockout mice and myocytes isolated
from these PLIN5
−/−
mice show an increased fatty acid oxidation
in vitro
compared to
the wild type. Moreover, ROS production is enhanced in the heart tissue of PLIN5
−/−
mice, which aggravates the age-related cardiomyopathy, but can be antagonized by the
glutathione-precursor N-acetylcysteine [
457
]. In addition, PLIN5 may also exert protection
from lipotoxicity by antagonizing ER stress, which has been shown for pancreatic
β
-cells
upon chronic exposure to free fatty acids [
458
]. Finally, a recent report demonstrated a
regulatory role for cardiac PLIN5 in cardiac Ca
2+
signaling and muscle contractility that is
based on the interaction between PLIN5 and sarcoplasmic/endoplasmic reticulum Ca
2+
ATPAase2 [
459
]. Summarizing, these findings put emphasis on the proper LD balance and
expression of LD-associated PLIN5 on heart integrity maintenance.
5.7. Lipid Droplets and Cancer—A General Outline
Growing evidence suggests a manifold involvement in LDs in cancer (reviewed
in [
460
]), however, it still is not clear whether LD accumulation plays a causative role in
carcinogenesis (as, for instance, is discussed above for the transition from NAFLD to HCC),
or is a consequence of increased lipid demands of tumor cells; or—most likely—both may
even apply. In many aspects, aging and tumorigenesis show opposing phenotypes, which
led to the proposal that anti-aging strategies can be developed based on tumor cells [
461
].
Attributable to the altered energy demands of tumor cell proliferation, elevated LD accu-
mulation is observable in different kinds of tumors, such as colorectal cancer, hepatocellular
and pancreatic carcinoma, renal cell carcinoma, prostate and breast cancer, lung cancer, and
glioblastoma [
462
–
464
]. For several cancers, a direct correlation between tumor cell sur-
vival, tumor aggressiveness, and LD numbers has been documented and tendencies exist
to consider cancer as a “LD-driven metabolic disease” [
465
]. A series of excellent reviews
addresses the question of how LDs can promote tumorigenesis [
460
,
462
,
465
], which are
recapitulated here briefly. As stated above, the most obvious role of LDs in cancer growth is
energy supply, with LD-derived FAs serving as fuel for
β
-oxidation and mitochondrial ATP
production [
466
]. LD-resident PLIN5 is essential to the FA flux between LDs and mitochon-
dria [
453
], which is essential to cellular lipid supply coping with increased energy demands
such as seen in tumor cells, but also in normal cells under stress conditions as discussed
above for cardiovascular disease. Of special pathophysiological relevance, the FAs released
from LDs are not only used for energy production, but also act as signaling molecules
(e.g., lysophosphatidic acid) regulating tumor progression and metastasis [
460
,
467
]. Fur-
thermore, LDs are able to modulate cell cycle checkpoints and gene expression in tumor
cells (e.g., G
0
/G
1
bypass and regulation of FOXO3A activity) [
460
,
468
,
469
]. In addition,
LDs also enable the intracellular trafficking of growth-signaling proteins such as PI3K,
ERK1, ERK2, p38, and PKC, as well as endo-/transcytosis-regulating caveolin, which are
also involved in tumorigenesis [470,471].
Moreover, LDs seem to be especially important for tumor initiation during early
carcinogenesis. In the so-called elimination phase, tumor defense by the both the innate

Biomolecules 2023,13, 912 35 of 56
and adaptive immune systems is based on the detection of potentially malignant cells
and their targeted elimination via apoptosis [
472
,
473
]. Besides acting inside the tumor
cells, LDs also interfere with the tumor microenvironment [
460
]. As an illustrative ex-
ample of the highly complex interactions between LDs, tumor cells, and the tumor cell
microenvironment, the role of LDs in cellular eicosanoid production [474] should be men-
tioned here. Eicosanoids (e.g., prostaglandins, leukotrienes, and lipoxins) are important
PUFA-derived (e.g., arachidonic acid) signaling molecules, which are secreted from tumor
cells into their microenvironment where they exert autocrine and paracrine activities. For
instance, prostaglandin E2 (PGE
2
) is mainly synthesized from LDs in cancer cells [
475
] and
immune suppression conferred by tumor-derived PGE
2
is deeply involved in the tumor
escape from immune surveillance [
476
]. In a complementary mode, PGE
2
is also involved
in tumor cell proliferation, angiogenesis, and metastasis [
477
]. Conversely, dendritic cells
enriched in LDs containing oxidized TAG show a dysfunctional antigen presentation [
478
],
which will also impair the host tumor defense.
Acting on the central balance of homeostatic growth control, LD accumulation may also
affect the onset of apoptosis [
479
,
480
], probably by delaying the accumulation of toxic fatty
acids inside the affected tissue [
481
,
482
]. Several findings account for the direct involvement
of LDs in cancer cell apoptosis. Notably, in both tumorous and non-tumorous cell lines,
stimulation of apoptosis occurs in conjunction with enhanced LD biogenesis [
35
,
483
], and
evidence exists that an increased LD content improves the tumor cells’ resistance to pro-
apoptotic stimuli. This may be due to the enhanced sequestration of a pro-apoptotic
stimulus by LDs, as was demonstrated for curcumin. This plant polyphenol stimulates
apoptosis via intrinsic, mitochondria-dependent signaling in several cell lines [
484
–
487
], but
fails to do so in glioblastoma cells [
488
]. In these cells, curcumin is efficiently sequestered
by the high LD content. Lowering LD numbers via inhibition of cytosolic phospholipase
A2 restores the sensitivity to curcumin-mediated apoptosis [
488
]. In a similar way, the
enhanced sequestration of chemotherapeutic drugs by LDs may render cancer treatment
inefficient and finally promote drug resistance [460,489].
However, further approaches exist to explain the anti-apoptotic role of LDs, addressing
intrinsic, mitochondria-dependent (MOMP/apoptosome), and extrinsic, death-receptor-
dependent (TNF
α
receptor superfamily /DISC) apoptotic signaling [
490
]. For instance,
it was shown that alterations of the cholesterol content of lipid rafts blocks the onset of
apoptosis induced upon TRAIL (tumor necrosis factor-related apoptosis-inducing ligand)
ligation to death receptors DR4 and DR5 in non-small cell lung carcinoma cells [
491
]. Con-
sidering that LDs serve as a reservoir for cholesterol, LDs could hypothetically contribute
to the suppression of extrinsic apoptosis. In addition, our own findings suggest a direct
involvement of LDs in intrinsic apoptotic signaling. We showed (for details see Section 2.3)
that mitochondria-localized apoptotic proteins (pro- as well as anti-apoptotic) contain a
V-domain that enables shuttling of these proteins from mitochondria to LDs. The affinity
of this V-domain is higher for LDs than for mitochondria and, upon an increase in the
cytosolic LD content (as seen in tumor cells), these apoptotic proteins are cleared from the
mitochondria, with the relocalization to LDs interrupting the apoptotic program [
35
]. In
fact, both pro- and anti-apoptotic proteins such as BAX [
12
,
35
,
492
], BCL-X
L
[
35
], Bcl-w [
12
],
AIFM1, AIFM2 [12], CCAR2 [12], API5 [492], and TPT1 [35,492] were shown to localize to
LDs in tumor cells.
Taken together, it is likely that LDs play a hitherto underestimated role in cancer biol-
ogy, addressing tumorigenesis at several critical instances. As discussed in this review, LDs
may confer protection by exerting antioxidant properties including lipid stress (LPO) under
healthy conditions. On the contrary, LDs may promote carcinogenesis in diseased contexts,
especially in chronic, inflammation-associated settings such as the “malignant” transition
from NASH to HCC, and may terminally also contribute to tumor progression and metas-
tasis by interfering with vascularization and proliferation–regulatory cell signaling in the
tumor environment.

Biomolecules 2023,13, 912 36 of 56
6. Concluding Remarks
In most model organisms, a clear picture seems to emerge that LDs, despite their
rather negative appraisal as a mere “fat-particle”, fulfill a cytoprotective role. This is due
to the “buffering” function of LDs, which enables them to take up lipid peroxides and
other oxidized lipid derivatives (e.g., oxLDL), as well as to detoxify misfolded proteins
and protein aggregates in and on various cell organelles. Accordingly, it is not surprising
that lifespan extension is positively correlated with LD abundance (at least to some extent).
Similar experimental evidence can be found throughout a diversity of biological model
systems, suggesting LDs inherit highly conserved functions. This picture is clearest in
simple organisms such as S. cerevisiae or C. elegans, but is also presented by the more
complex organism D. melanogaster. Central to this LD–aging connection seems to be
metabolic pathways such as TOR signaling or IIS (see Figure 2), which upon inhibition
lead to both prolonged lifespan and elevated LD synthesis. The situation in mammals and
humans is more difficult to interpret due to markedly larger cell numbers, the enhanced
diversity of differentiated cell types, and the complex interaction among diverse tissues. It
is striking that LDs are concomitant to age-related disease. Concerning this, however, we
want to question the still prevailing concept that LDs, by oversimplification understood as
“monofunctional” fat-accumulating vesicles, are causative of the pathogenesis of age-related
diseases. Taken together, the existing literature advocates a different view, suggesting that
LDs represent multifunctional organelles of particular physiological relevance that play a
subtle, Janus-faced role in disease: LDs essentially fulfilling a protective, retarding function
during early pathogenetic stages, but converting to the opposite function in the course
of disease progression when an excessive accumulation of LDs amplifies a phenotype
characteristic of advanced disease states.
Moreover, disregarding the pathogenic aspect, there is evidence for a physiological
role of LDs as important “players” in healthy aging in humans. It is well accepted that
the Mediterranean diet has numerous beneficial effects on human health. Many studies
have shown that this diet reduces mortality and lowers the risk of developing cancer,
neurodegenerative diseases, and cardiovascular diseases [
493
]. Some of the effects of
the Mediterranean diet can be attributed to sirtuins [
494
], which have been addressed
as regulators of LD biogenesis at several instances in this review. This puts emphasis
on beneficial nutritional aspects, in particular focusing on two essential pillars of the
Mediterranean diet: red wine and olive oil. In fact, it was shown that resveratrol, a
polyphenol enriched in red wine, is an activator of sirtuin Sir2p (the yeast homologue of
Sirt1) that has the capability to extend the lifespan in a broad variety of organisms [
83
]. It
has to be noted critically that the activation of yeast Sirt1 by resveratrol occurs in an indirect
mode via the cAMP-Epac1-AMPK-Sirt1 pathway, with Sirt1 being likely to be activated by
increased cellular amounts of NAD
+
[
495
]. Recently it was shown that monounsaturated
fatty acids such as oleic acid, the main component of olive oil, allosterically activate Sirt1 at
a magnitude many times higher than that of resveratrol [
359
]. Considering the connection
between Sirt signaling and LD biogenesis, it would be thrilling to see, in the future, if some
of the positive effects of the Mediterranean diet can be attributed to the stimulation of
LD biogenesis.
In synopsis, it is obvious that lipid metabolism is closely linked to aging and cellular
stress responses via highly complex interactions that are not yet fully understood. Accord-
ing to our recent knowledge, it can be concluded that LDs participate in these complex
metabolic, aging-associated networks by playing a “Janus-faced” role, as illustrated in
Figure 3, and it will be the subject of future investigation to elucidate the exact, underlying
contexts in detail.

Biomolecules 2023,13, 912 37 of 56
Biomolecules 2023, 13, 912 37 of 58
to the opposite function in the course of disease progression when an excessive
accumulation of LDs amplifies a phenotype characteristic of advanced disease states.
Moreover, disregarding the pathogenic aspect, there is evidence for a physiological
role of LDs as important “players” in healthy aging in humans. It is well accepted that the
Mediterranean diet has numerous beneficial effects on human health. Many studies have
shown that this diet reduces mortality and lowers the risk of developing cancer,
neurodegenerative diseases, and cardiovascular diseases [493]. Some of the effects of the
Mediterranean diet can be aributed to sirtuins [494], which have been addressed as
regulators of LD biogenesis at several instances in this review. This puts emphasis on
beneficial nutritional aspects, in particular focusing on two essential pillars of the
Mediterranean diet: red wine and olive oil. In fact, it was shown that resveratrol, a
polyphenol enriched in red wine, is an activator of sirtuin Sir2p (the yeast homologue of
Sirt1) that has the capability to extend the lifespan in a broad variety of organisms [83]. It
has to be noted critically that the activation of yeast Sirt1 by resveratrol occurs in an
indirect mode via the cAMP-Epac1-AMPK-Sirt1 pathway, with Sirt1 being likely to be
activated by increased cellular amounts of NAD+ [495]. Recently it was shown that
monounsaturated fay acids such as oleic acid, the main component of olive oil,
allosterically activate Sirt1 at a magnitude many times higher than that of resveratrol [359].
Considering the connection between Sirt signaling and LD biogenesis, it would be
thrilling to see, in the future, if some of the positive effects of the Mediterranean diet can
be aributed to the stimulation of LD biogenesis.
In synopsis, it is obvious that lipid metabolism is closely linked to aging and cellular
stress responses via highly complex interactions that are not yet fully understood.
According to our recent knowledge, it can be concluded that LDs participate in these
complex metabolic, aging-associated networks by playing a “Janus-faced” role, as
illustrated in Figure 3, and it will be the subject of future investigation to elucidate the
exact, underlying contexts in detail.
Figure 3. Model for the Janus-faced role of LDs in the aging process. Central to this explanatory
approach is the bifunctional involvement of LDs in cellular maintenance, with LDs serving as both
Figure 3.
Model for the Janus-faced role of LDs in the aging process. Central to this explanatory
approach is the bifunctional involvement of LDs in cellular maintenance, with LDs serving as both
(i) a dynamic lipid /fat buffer, and (ii) a “sink” for toxic compounds upon LD clearance (proteins,
lipids, and toxic compounds). In addition, LDs may also provide lipid protection by preserving
PUFAs from excessive LPO and can quench stress derived from extrinsic factors. Under stress
conditions, such as those created by ROS, the ER, or mitochondria, stimulation of LD biogenesis
becomes the most important factor, and excessive LD formation depends on a variety of intrinsic
factors (TOR, IIS, and TGF-
β
). The increase in the LD pool establishes a delicate balance between
LD accumulation and LD-based detoxification. This balance determines the outcome of the stress
response, which protects against cell death but may result in a chronic process (disease/inflammation)
based on it. In contrast with the initial beneficial effects, the accumulation of LD accelerates the
progression of chronic disease and thus the “aging” process. Hence, in stressed cells, LD biogenesis
and LD functionality, both indirectly and directly, intervenes with multifaceted cellular life–death
decisions (shaded blue area).
Author Contributions:
Conceptualization, M.R.; writing—original draft preparation, N.B., M.K.,
A.L., T.K.F. and M.R.; writing—review and editing, N.B., M.K., A.L., T.K.F. and M.R.; visualization,
N.B. and M.R.; funding acquisition, M.R. All authors have read and agreed to the published version
of the manuscript.
Funding:
This research was funded by the Austrian Science Fund (FWF) with the grant P33511
to M.R.
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement: Not applicable.
Acknowledgments:
This scientific work is dedicated to the great scientist Breitenbach, with whom it
was always a pleasure to cooperate and discuss science.

Biomolecules 2023,13, 912 38 of 56
Conflicts of Interest: The authors declare no conflict of interest.
Abbreviations
AD, Alzheimer’s disease; AMPK, AMP-activated protein kinase; ANT, adenine-
nucleotide translocator; Apo and APO, apolipoproteins; ARDs, age-related diseases; ATF6,
activating transcription factor 6; ATG, autophagy related genes; ALS, amyotrophic lateral
sclerosis; CBS, cystathionine
β
-synthase; Ces1d, cholesteryl-ester hydrolase; CCN1, cen-
tral communication network factor 1; DFP, deferiprone; ECM, extracellular matrix; ER,
endoplasmic reticulum; ESCRT, endosomal sorting complexes required for transport; Fe-S,
iron-sulfur clusters; FOXO, forkhead box O; FRDA, Friedreich’s Ataxia; Ftx, frataxin; GAP,
GTPase activating protein; GSH, glutathione (reduced from); GSSG, the oxidized (disul-
fide) form of GSH; HCC, hepatocellular carcinoma; HDACs, histone deacetylases; HNE,
4-hydroxy-2-nonenal; HNF1
β
, hepatocyte nuclear factor 1
β
; HSCs, hepatic stellate cells;
IBs, inclusion bodies; IIS, Insulin/Insulin growth factor -1 signaling; ISC, intestinal stem
cell; ISCU, iron–sulfur cluster forming unit; JNK, c-jun amino-terminal kinase; LDs, Lipid
droplets; LDL, low-density lipoprotein; oxLDL, oxidized LDL; Lox, peroxidized lipids; LPO,
lipid peroxidation; LROs, lysosome-related organelles; MAC, mitochondrial-apoptosis-
induced channel; MAGIC, mitochondria as guardian in cytosol; MAT, marrow adipose
tissue; MOMP, mitochondrial outer membrane permeabilization; MSCs, bone marrow
stromal cells; mPT, mitochondrial permeability transition pore; NAFLD, non-alcoholic fatty
liver disease; NASH, non-alcoholic steatohepatitis; nLDs, nuclear LDs; NOX1, NADPH
oxidase 1; PA, phosphatic acid; PCD, programmed cell death; PD, Parkinson’s disease;
PERK, PKR-like ER kinase; PLIN, perilipin; PML, promyelocytic leukemia protein; PPAR,
peroxisome proliferator-activated receptor; PUFAs, polyunsaturated fatty acids; rad, ra-
diation damage; raptor, regulatory associated protein of mTOR; RCT, reverse cholesterol
transport; rictor, rapamycin-insensitive companion of mTOR; ROS, reactive oxygen species;
SIRT, sirtuin; TAG, triacylglycerol(s); TE, transposable element; TNF
α
, tumor necrosis
factor
α
; TNFR, TNF
α
-receptor; TOR, target of rapamycin; mTORC, mammalian TOR
complex; TSC, trans-sulfuration pathway; UPR, unfolded protein response; UPR
ER
, ER
stress-associated UPR; TSC, tuberous sclerosis complex; VDAC, voltage-dependent anion
channel; VLDL, very-low-density lipoprotein; VSMCs, vascular smooth muscle cells; Xbp1,
X-Box binding protein 1.
References
1. Murphy, D.J.; Vance, J. Mechanisms of lipid-body formation. Trends Biochem. Sci. 1999,24, 109–115. [CrossRef]
2.
Martin, S.; Parton, R.G. Lipid droplets: A unified view of a dynamic organelle. Nat. Rev. Mol. Cell Biol.
2006
,7, 373–378. [CrossRef]
[PubMed]
3. Wilfling, F.; Haas, J.T.; Walther, T.C.; Farese, R.V., Jr. Lipid droplet biogenesis. Curr. Opin. Cell Biol. 2014,29, 39–45. [CrossRef]
4.
Geltinger, F.; Schartel, L.; Wiederstein, M.; Tevini, J.; Aigner, E.; Felder, T.K.; Rinnerthaler, M. Friend or foe: Lipid droplets
as organelles for protein and lipid storage in cellular stress response, aging and disease. Molecules
2020
,25, 5053. [CrossRef]
[PubMed]
5.
Plotz, T.; Hartmann, M.; Lenzen, S.; Elsner, M. The role of lipid droplet formation in the protection of unsaturated fatty acids
against palmitic acid induced lipotoxicity to rat insulin-producing cells. Nutr. Metab. 2016,13, 16. [CrossRef]
6.
Listenberger, L.L.; Han, X.; Lewis, S.E.; Cases, S.; Farese, R.V., Jr.; Ory, D.S.; Schaffer, J.E. Triglyceride accumulation protects
against fatty acid-induced lipotoxicity. Proc. Natl. Acad. Sci. USA 2003,100, 3077–3082. [CrossRef] [PubMed]
7.
Geltinger, F.; Tevini, J.; Briza, P.; Geiser, A.; Bischof, J.; Richter, K.; Felder, T.; Rinnerthaler, M. The transfer of specific mitochondrial
lipids and proteins to lipid droplets contributes to proteostasis upon stress and aging in the eukaryotic model system Saccharomyces
cerevisiae.Geroscience 2020,42, 19–38. [CrossRef]
8.
Moldavski, O.; Amen, T.; Levin-Zaidman, S.; Eisenstein, M.; Rogachev, I.; Brandis, A.; Kaganovich, D.; Schuldiner, M. Lipid
Droplets Are Essential for Efficient Clearance of Cytosolic Inclusion Bodies. Dev. Cell 2015,33, 603–610. [CrossRef]
9.
Vevea, J.D.; Garcia, E.J.; Chan, R.B.; Zhou, B.; Schultz, M.; Di Paolo, G.; McCaffery, J.M.; Pon, L.A. Role for Lipid Droplet
Biogenesis and Microlipophagy in Adaptation to Lipid Imbalance in Yeast. Dev. Cell 2015,35, 584–599. [CrossRef]

Biomolecules 2023,13, 912 39 of 56
10.
Garcia, E.J.; Liao, P.C.; Tan, G.; Vevea, J.D.; Sing, C.N.; Tsang, C.A.; McCaffery, J.M.; Boldogh, I.R.; Pon, L.A. Membrane dynamics
and protein targets of lipid droplet microautophagy during ER stress-induced proteostasis in the budding yeast, Saccharomyces
cerevisiae.Autophagy 2021,17, 2363–2383. [CrossRef]
11.
Kumar, R.; Nawroth, P.P.; Tyedmers, J. Prion Aggregates Are Recruited to the Insoluble Protein Deposit (IPOD) via Myosin
2-Based Vesicular Transport. PLoS Genet. 2016,12, e1006324. [CrossRef]
12.
Bersuker, K.; Peterson, C.W.H.; To, M.; Sahl, S.J.; Savikhin, V.; Grossman, E.A.; Nomura, D.K.; Olzmann, J.A. A Proximity Labeling
Strategy Provides Insights into the Composition and Dynamics of Lipid Droplet Proteomes. Dev. Cell
2018
,44, 97–112.e117.
[CrossRef]
13.
Greenberg, A.S.; Egan, J.J.; Wek, S.A.; Garty, N.B.; Blanchette-Mackie, E.J.; Londos, C. Perilipin, a major hormonally regulated
adipocyte-specific phosphoprotein associated with the periphery of lipid storage droplets. J. Biol. Chem.
1991
,266, 11341–11346.
[CrossRef] [PubMed]
14.
Cho, S.Y.; Shin, E.S.; Park, P.J.; Shin, D.W.; Chang, H.K.; Kim, D.; Lee, H.H.; Lee, J.H.; Kim, S.H.; Song, M.J.; et al. Identification of
mouse Prp19p as a lipid droplet-associated protein and its possible involvement in the biogenesis of lipid droplets. J. Biol. Chem.
2007,282, 2456–2465. [CrossRef] [PubMed]
15.
Turro, S.; Ingelmo-Torres, M.; Estanyol, J.M.; Tebar, F.; Fernandez, M.A.; Albor, C.V.; Gaus, K.; Grewal, T.; Enrich, C.; Pol, A.
Identification and characterization of associated with lipid droplet protein 1: A novel membrane-associated protein that resides
on hepatic lipid droplets. Traffic 2006,7, 1254–1269. [CrossRef]
16.
Onal, G.; Kutlu, O.; Gozuacik, D.; Dokmeci Emre, S. Lipid Droplets in Health and Disease. Lipids Health Dis.
2017
,16, 128.
[CrossRef]
17.
Renne, M.F.; Hariri, H. Lipid Droplet-Organelle Contact Sites as Hubs for Fatty Acid Metabolism, Trafficking, and Metabolic
Channeling. Front. Cell Dev. Biol. 2021,9, 726261. [CrossRef] [PubMed]
18.
Li, Z.; Thiel, K.; Thul, P.J.; Beller, M.; Kühnlein, R.P.; Welte, M.A. Lipid droplets control the maternal histone supply of Drosophila
embryos. Curr. Biol. 2012,22, 2104–2113. [CrossRef]
19.
Kovacs, M.; Geltinger, F.; Verwanger, T.; Weiss, R.; Richter, K.; Rinnerthaler, M. Lipid Droplets Protect Aging Mitochondria and
Thus Promote Lifespan in Yeast Cells. Front. Cell Dev. Biol. 2021,9, 774985. [CrossRef]
20.
Suriyalaksh, M.; Raimondi, C.; Mains, A.; Segonds-Pichon, A.; Mukhtar, S.; Murdoch, S.; Aldunate, R.; Krueger, F.; Guimera, R.;
Andrews, S.; et al. Gene regulatory network inference in long-lived C. elegans reveals modular properties that are predictive of
novel aging genes. Iscience 2022,25, 103663. [CrossRef]
21.
Zhao, X.; Li, X.; Shi, X.; Karpac, J. Diet-MEF2 interactions shape lipid droplet diversification in muscle to influence Drosophila
lifespan. Aging Cell 2020,19, e13172. [CrossRef]
22.
Zimmermann, A.; Hofer, S.; Pendl, T.; Kainz, K.; Madeo, F.; Carmona-Gutierrez, D. Yeast as a tool to identify anti-aging
compounds. Fems Yeast Res. 2018,18, foy020. [CrossRef]
23.
Stefanini, I.; De Filippo, C.; Cavalieri, D. Yeast as a Model in High-Throughput Screening of Small-Molecule Libraries. In
Diversity-Oriented Synthesis; Wiley Online Library: Hoboken, NJ, USA, 2013; pp. 455–482.
24.
Steinkraus, K.A.; Kaeberlein, M.; Kennedy, B.K. Replicative aging in yeast: The means to the end. Annu. Rev. Cell Dev. Biol.
2008
,
24, 29–54. [CrossRef] [PubMed]
25. Rockenfeller, P.; Madeo, F. Apoptotic death of ageing yeast. Exp. Gerontol. 2008,43, 876–881. [CrossRef] [PubMed]
26.
Fabrizio, P.; Longo, V.D. The chronological life span of Saccharomyces cerevisiae.Methods Mol. Biol.
2007
,371, 89–95. [CrossRef]
[PubMed]
27.
Bitterman, K.J.; Medvedik, O.; Sinclair, D.A. Longevity regulation in Saccharomyces cerevisiae: Linking metabolism, genome
stability, and heterochromatin. Microbiol. Mol. Biol. Rev. 2003,67, 376–399. [CrossRef]
28.
Pringle, J.R. Staining of bud scars and other cell wall chitin with calcofluor. Methods Enzymol.
1991
,194, 732–735. [CrossRef]
[PubMed]
29.
Klinger, H.; Rinnerthaler, M.; Lam, Y.T.; Laun, P.; Heeren, G.; Klocker, A.; Simon-Nobbe, B.; Dickinson, J.R.; Dawes, I.W.;
Breitenbach, M. Quantitation of (a)symmetric inheritance of functional and of oxidatively damaged mitochondrial aconitase in
the cell division of old yeast mother cells. Exp. Gerontol. 2010,45, 533–542. [CrossRef]
30. Hayflick, L.; Moorhead, P.S. The serial cultivation of human diploid cell strains. Exp. Cell Res. 1961,25, 585–621. [CrossRef]
31.
Sinclair, D.A. Studying the Replicative Life Span of Yeast Cells. In Biological Aging: Methods and Protocols, 2nd ed.; Springer:
Berlin/Heidelberg, Germany, 2013; Volume 1048, pp. 49–63. [CrossRef]
32.
Lin, S.J.; Defossez, P.A.; Guarente, L. Requirement of NAD and SIR2 for life-span extension by calorie restriction in Saccharomyces
cerevisiae.Science 2000,289, 2126–2128. [CrossRef]
33.
Goldberg, A.A.; Bourque, S.D.; Kyryakov, P.; Boukh-Viner, T.; Gregg, C.; Beach, A.; Burstein, M.T.; Machkalyan, G.; Richard, V.;
Rampersad, S.; et al. A novel function of lipid droplets in regulating longevity. Biochem. Soc. Trans.
2009
,37, 1050–1055. [CrossRef]
34.
Hiltunen, J.K.; Mursula, A.M.; Rottensteiner, H.; Wierenga, R.K.; Kastaniotis, A.J.; Gurvitz, A. The biochemistry of peroxisomal
beta-oxidation in the yeast Saccharomyces cerevisiae.FEMS Microbiol. Rev. 2003,27, 35–64. [CrossRef] [PubMed]
35.
Bischof, J.; Salzmann, M.; Streubel, M.K.; Hasek, J.; Geltinger, F.; Duschl, J.; Bresgen, N.; Briza, P.; Haskova, D.; Lejskova, R.; et al.
Clearing the outer mitochondrial membrane from harmful proteins via lipid droplets. Cell Death Discov.
2017
,3, 17016. [CrossRef]
[PubMed]

Biomolecules 2023,13, 912 40 of 56
36.
Beas, A.O.; Gordon, P.B.; Prentiss, C.L.; Olsen, C.P.; Kukurugya, M.A.; Bennett, B.D.; Parkhurst, S.M.; Gottschling, D.E.
Independent regulation of age associated fat accumulation and longevity. Nat. Commun. 2020,11, 2790. [CrossRef] [PubMed]
37.
Martínez, G.; Duran-Aniotz, C.; Cabral-Miranda, F.; Vivar, J.P.; Hetz, C. Endoplasmic reticulum proteostasis impairment in aging.
Aging Cell 2017,16, 615–623. [CrossRef]
38.
Cui, H.-J.; Liu, X.-G.; McCormick, M.; Wasko, B.M.; Zhao, W.; He, X.; Yuan, Y.; Fang, B.-X.; Sun, X.-R.; Kennedy, B.K.; et al. PMT1
deficiency enhances basal UPR activity and extends replicative lifespan of Saccharomyces cerevisiae.Age 2015,37, 46. [CrossRef]
39.
Walter, P.; Ron, D. The unfolded protein response: From stress pathway to homeostatic regulation. Science
2011
,334, 1081–1086.
[CrossRef]
40.
Kopito, R.R.; Sitia, R. Aggresomes and Russell bodies. Symptoms of cellular indigestion? EMBO Rep.
2000
,1, 225–231. [CrossRef]
41.
Celik, C.; Lee, S.Y.T.; Yap, W.S.; Thibault, G. Endoplasmic reticulum stress and lipids in health and diseases. Prog. Lipid Res.
2022
,
89, 101198. [CrossRef]
42.
Halbleib, K.; Pesek, K.; Covino, R.; Hofbauer, H.F.; Wunnicke, D.; Hanelt, I.; Hummer, G.; Ernst, R. Activation of the Unfolded
Protein Response by Lipid Bilayer Stress. Mol. Cell 2017,67, 673–684.e678. [CrossRef]
43.
Rubio, C.; Pincus, D.; Korennykh, A.; Schuck, S.; El-Samad, H.; Walter, P. Homeostatic adaptation to endoplasmic reticulum stress
depends on Ire1 kinase activity. J. Cell Biol. 2011,193, 171–184. [CrossRef] [PubMed]
44.
Lee, A.H.; Scapa, E.F.; Cohen, D.E.; Glimcher, L.H. Regulation of hepatic lipogenesis by the transcription factor XBP1. Science
2008,320, 1492–1496. [CrossRef] [PubMed]
45.
Luo, W.; Wang, H.; Ren, L.; Lu, Z.; Zheng, Q.; Ding, L.; Xie, H.; Wang, R.; Yu, C.; Lin, Y.; et al. Adding fuel to the fire: The lipid
droplet and its associated proteins in cancer progression. Int. J. Biol. Sci. 2022,18, 6020–6034. [CrossRef]
46.
Liu, L.; Zhang, K.; Sandoval, H.; Yamamoto, S.; Jaiswal, M.; Sanz, E.; Li, Z.; Hui, J.; Graham, B.H.; Quintana, A.; et al. Glial lipid
droplets and ROS induced by mitochondrial defects promote neurodegeneration. Cell 2015,160, 177–190. [CrossRef] [PubMed]
47. Walther, T.C.; Chung, J.; Farese, R.V., Jr. Lipid Droplet Biogenesis. Annu. Rev. Cell Dev. Biol. 2017,33, 491–510. [CrossRef]
48.
Harman, D. Aging: A Theory Based on Free Radical and Radiation Chemistry. J. Gerontol.
1956
,11, 298–300. [CrossRef] [PubMed]
49.
Cohen, A.; Weindling, E.; Rabinovich, E.; Nachman, I.; Fuchs, S.; Chuartzman, S.; Gal, L.; Schuldiner, M.; Bar-Nun, S. Water-
Transfer Slows Aging in Saccharomyces cerevisiae.PLoS ONE 2016,11, e0148650. [CrossRef] [PubMed]
50.
Gao, Q.; Binns, D.D.; Kinch, L.N.; Grishin, N.V.; Ortiz, N.; Chen, X.; Goodman, J.M. Pet10p is a yeast perilipin that stabilizes lipid
droplets and promotes their assembly. J. Cell Biol. 2017,216, 3199–3217. [CrossRef]
51.
Di Gregorio, S.E.; Duennwald, M.L. Yeast as a model to study protein misfolding in aged cells. FEMS Yeast Res.
2018
,18, foy054.
[CrossRef]
52.
Currie, E.; Guo, X.; Christiano, R.; Chitraju, C.; Kory, N.; Harrison, K.; Haas, J.; Walther, T.C.; Farese, R.V. High confidence
proteomic analysis of yeast LDs identifies additional droplet proteins and reveals connections to dolichol synthesis and sterol
acetylation. J. Lipid Res. 2014,55, 1465–1477. [CrossRef]
53.
Grillitsch, K.; Connerth, M.; Köfeler, H.; Arrey, T.N.; Rietschel, B.; Wagner, B.; Karas, M.; Daum, G. Lipid particles/droplets of
the yeast Saccharomyces cerevisiae revisited: Lipidome meets proteome. Biochim. Biophys. Acta
2011
,1811, 1165–1176. [CrossRef]
[PubMed]
54.
Wang, C.-W.; Lee, S.-C. The ubiquitin-like (UBX)-domain-containing protein Ubx2/Ubxd8 regulates lipid droplet homeostasis.
J. Cell Sci. 2012,125, 2930–2939. [CrossRef] [PubMed]
55.
Neuber, O.; Jarosch, E.; Volkwein, C.; Walter, J.; Sommer, T. Ubx2 links the Cdc48 complex to ER-associated protein degradation.
Nat. Cell Biol. 2005,7, 993–998. [CrossRef] [PubMed]
56.
Laun, P.; Büttner, S.; Rinnerthaler, M.; Burhans, W.C.; Breitenbach, M. Yeast Aging and Apoptosis. In Aging Research in Yeast;
Breitenbach, M., Jazwinski, S.M., Laun, P., Eds.; Springer: Dordrecht, The Netherlands, 2012; pp. 207–232.
57. Côrte-Real, M.; Madeo, F. Yeast Programed Cell Death and Aging. Front. Oncol. 2013,3, 283. [CrossRef] [PubMed]
58. Tower, J. Programmed cell death in aging. Ageing Res. Rev. 2015,23, 90–100. [CrossRef]
59.
Chipuk, J.E.; Bouchier-Hayes, L.; Green, D.R. Mitochondrial outer membrane permeabilization during apoptosis: The innocent
bystander scenario. Cell Death Differ. 2006,13, 1396–1402. [CrossRef]
60.
Dadsena, S.; King, L.E.; García-Sáez, A.J. Apoptosis regulation at the mitochondria membrane level. Biochim. Biophys. Acta
Biomembr. 2021,1863, 183716. [CrossRef]
61.
Subburaj, Y.; Cosentino, K.; Axmann, M.; Pedrueza-Villalmanzo, E.; Hermann, E.; Bleicken, S.; Spatz, J.; García-Sáez, A.J. Bax
monomers form dimer units in the membrane that further self-assemble into multiple oligomeric species. Nat. Commun.
2015
,
6, 8042. [CrossRef]
62.
Rinnerthaler, M.; Lejskova, R.; Grousl, T.; Stradalova, V.; Heeren, G.; Richter, K.; Breitenbach-Koller, L.; Malinsky, J.; Hasek, J.;
Breitenbach, M. Mmi1, the yeast homologue of mammalian TCTP, associates with stress granules in heat-shocked cells and
modulates proteasome activity. PLoS ONE 2013,8, e77791. [CrossRef]
63.
Rinnerthaler, M.; Jarolim, S.; Heeren, G.; Palle, E.; Perju, S.; Klinger, H.; Bogengruber, E.; Madeo, F.; Braun, R.J.;
Breitenbach-Koller, L.; et al. MMI1 (YKL056c, TMA19), the yeast orthologue of the translationally controlled tumor pro-
tein (TCTP) has apoptotic functions and interacts with both microtubules and mitochondria. Biochim. Biophys. Acta
2006
,1757,
631–638. [CrossRef]
64.
Ding, W.X.; Yin, X.M. Mitophagy: Mechanisms, pathophysiological roles, and analysis. Biol. Chem.
2012
,393, 547–564. [CrossRef]

Biomolecules 2023,13, 912 41 of 56
65.
Kissová, I.; Deffieu, M.; Manon, S.; Camougrand, N. Uth1p is involved in the autophagic degradation of mitochondria. J. Biol.
Chem. 2004,279, 39068–39074. [CrossRef]
66.
Kim, E.H.; Choi, K.S. A critical role of superoxide anion in selenite-induced mitophagic cell death. Autophagy
2008
,4, 76–78.
[CrossRef] [PubMed]
67.
Chu, C.T.; Zhu, J.; Dagda, R. Beclin 1-independent pathway of damage-induced mitophagy and autophagic stress: Implications
for neurodegeneration and cell death. Autophagy 2007,3, 663–666. [CrossRef] [PubMed]
68.
Kim, I.; Rodriguez-Enriquez, S.; Lemasters, J.J. Selective degradation of mitochondria by mitophagy. Arch. Biochem. Biophys.
2007
,
462, 245–253. [CrossRef] [PubMed]
69.
Nowikovsky, K.; Reipert, S.; Devenish, R.J.; Schweyen, R.J. Mdm38 protein depletion causes loss of mitochondrial K
+
/H+
exchange activity, osmotic swelling and mitophagy. Cell Death Differ. 2007,14, 1647–1656. [CrossRef]
70.
Terman, A.; Kurz, T.; Navratil, M.; Arriaga, E.A.; Brunk, U.T. Mitochondrial turnover and aging of long-lived postmitotic cells:
The mitochondrial-lysosomal axis theory of aging. Antioxid. Redox Signal. 2010,12, 503–535. [CrossRef]
71.
Bergamini, E. Autophagy: A cell repair mechanism that retards ageing and age-associated diseases and can be intensified
pharmacologically. Mol. Asp. Med. 2006,27, 403–410. [CrossRef]
72.
Stevens, M.; Oltean, S. Modulation of the Apoptosis Gene Bcl-x Function Through Alternative Splicing. Front. Genet.
2019
,10, 804.
[CrossRef]
73.
Kanagavijayan, D.; Rajasekharan, R.; Srinivasan, M. Yeast MRX deletions have short chronological life span and more triacylglyc-
erols. Fems Yeast Res. 2016,16, fov109. [CrossRef]
74.
López-Otín, C.; Blasco, M.A.; Partridge, L.; Serrano, M.; Kroemer, G. The hallmarks of aging. Cell
2013
,153, 1194–1217. [CrossRef]
75.
Aung-Htut, M.T.; Lam, Y.T.; Lim, Y.L.; Rinnerthaler, M.; Gelling, C.L.; Yang, H.; Breitenbach, M.; Dawes, I.W. Maintenance of
mitochondrial morphology by autophagy and its role in high glucose effects on chronological lifespan of Saccharomyces cerevisiae.
Oxidative Med. Cell. Longev. 2013,2013, 636287. [CrossRef]
76.
Sorger, D.; Athenstaedt, K.; Hrastnik, C.; Daum, G. A yeast strain lacking lipid particles bears a defect in ergosterol formation.
J. Biol. Chem. 2004,279, 31190–31196. [CrossRef] [PubMed]
77.
Valachovic, M.; Hronska, L.; Hapala, I. Anaerobiosis induces complex changes in sterol esterification pattern in the yeast
Saccharomyces cerevisiae.FEMS Microbiol. Lett. 2001,197, 41–45. [CrossRef] [PubMed]
78.
Leber, R.; Zinser, E.; Zellnig, G.; Paltauf, F.; Daum, G. Characterization of lipid particles of the yeast, Saccharomyces cerevisiae.Yeast
1994,10, 1421–1428. [CrossRef]
79.
Ruan, L.H.; Zhou, C.K.; Jin, E.L.; Kucharavy, A.; Zhang, Y.; Wen, Z.H.; Florens, L.; Li, R. Cytosolic proteostasis through importing
of misfolded proteins into mitochondria. Nature 2017,543, 443–446. [CrossRef] [PubMed]
80.
Erjavec, N.; Bayot, A.; Gareil, M.; Camougrand, N.; Nystrom, T.; Friguet, B.; Bulteau, A.L. Deletion of the mitochondrial Pim1/Lon
protease in yeast results in accelerated aging and impairment of the proteasome. Free Radic. Biol. Med.
2013
,56, 9–16. [CrossRef]
[PubMed]
81.
Lin, S.J.; Kaeberlein, M.; Andalis, A.A.; Sturtz, L.A.; Defossez, P.A.; Culotta, V.C.; Fink, G.R.; Guarente, L. Calorie restriction
extends Saccharomyces cerevisiae lifespan by increasing respiration. Nature 2002,418, 344–348. [CrossRef] [PubMed]
82.
Powers, R.W., 3rd; Kaeberlein, M.; Caldwell, S.D.; Kennedy, B.K.; Fields, S. Extension of chronological life span in yeast by
decreased TOR pathway signaling. Genes Dev. 2006,20, 174–184. [CrossRef]
83.
Howitz, K.T.; Bitterman, K.J.; Cohen, H.Y.; Lamming, D.W.; Lavu, S.; Wood, J.G.; Zipkin, R.E.; Chung, P.; Kisielewski, A.;
Zhang, L.L.; et al. Small molecule activators of sirtuins extend Saccharomyces cerevisiae lifespan. Nature
2003
,425, 191–196.
[CrossRef] [PubMed]
84.
Madeira, J.B.; Masuda, C.A.; Maya-Monteiro, C.M.; Matos, G.S.; Montero-Lomeli, M.; Bozaquel-Morais, B.L. TORC1 Inhibition
Induces Lipid Droplet Replenishment in Yeast. Mol. Cell. Biol. 2015,35, 737–746. [CrossRef] [PubMed]
85.
Vall-llaura, N.; Mir, N.; Garrido, L.; Vived, C.; Cabiscol, E. Redox control of yeast Sir2 activity is involved in acetic acid resistance
and longevity. Redox. Biol. 2019,24, 101229. [CrossRef] [PubMed]
86.
Boender, L.G.; Almering, M.J.; Dijk, M.; van Maris, A.J.; de Winde, J.H.; Pronk, J.T.; Daran-Lapujade, P. Extreme calorie restriction
and energy source starvation in Saccharomyces cerevisiae represent distinct physiological states. Biochim. Biophys. Acta
2011
,
1813, 2133–2144. [CrossRef]
87.
Schurmanns, L.; Hamann, A.; Osiewacz, H.D. Lifespan Increase of Podospora anserina by Oleic Acid Is Linked to Alterations in
Energy Metabolism, Membrane Trafficking and Autophagy. Cells 2022,11, 519. [CrossRef] [PubMed]
88.
Friedman, D.B.; Johnson, T.E. A mutation in the age-1 gene in Caenorhabditis elegans lengthens life and reduces hermaphrodite
fertility. Genetics 1988,118, 75–86. [CrossRef]
89.
Iser, W.B.; Wolkow, C.A. DAF-2/insulin-like signaling in C. elegans modifies effects of dietary restriction and nutrient stress on
aging, stress and growth. PLoS ONE 2007,2, e1240. [CrossRef]
90.
Jia, K.; Chen, D.; Riddle, D.L. The TOR pathway interacts with the insulin signaling pathway to regulate C. elegans larval
development, metabolism and life span. Development 2004,131, 3897–3906. [CrossRef]
91.
Johnson, S.C.; Rabinovitch, P.S.; Kaeberlein, M. mTOR is a key modulator of ageing and age-related disease. Nature
2013
,
493, 338–345. [CrossRef]
92.
Lakowski, B.; Hekimi, S. The genetics of caloric restriction in Caenorhabditis elegans.Proc. Natl. Acad. Sci. USA
1998
,
95, 13091–13096. [CrossRef]

Biomolecules 2023,13, 912 42 of 56
93.
Luo, S.; Kleemann, G.A.; Ashraf, J.M.; Shaw, W.M.; Murphy, C.T. TGF-beta and insulin signaling regulate reproductive aging via
oocyte and germline quality maintenance. Cell 2010,143, 299–312. [CrossRef]
94.
Greer, E.L.; Dowlatshahi, D.; Banko, M.R.; Villen, J.; Hoang, K.; Blanchard, D.; Gygi, S.P.; Brunet, A. An AMPK-FOXO pathway
mediates longevity induced by a novel method of dietary restriction in C. elegans.Curr. Biol. 2007,17, 1646–1656. [CrossRef]
95.
Leiser, S.F.; Kaeberlein, M. The hypoxia-inducible factor HIF-1 functions as both a positive and negative modulator of aging. Biol.
Chem. 2010,391, 1131–1137. [CrossRef]
96.
Golden, J.W.; Riddle, D.L. The Caenorhabditis elegans dauer larva: Developmental effects of pheromone, food, and temperature.
Dev. Biol. 1984,102, 368–378. [CrossRef]
97. Klass, M.; Hirsh, D. Non-ageing developmental variant of Caenorhabditis elegans.Nature 1976,260, 523–525. [CrossRef]
98.
Zhang, S.; Li, F.; Zhou, T.; Wang, G.; Li, Z. Caenorhabditis elegans as a Useful Model for Studying Aging Mutations. Front. Endocrinol.
2020,11, 554994. [CrossRef]
99.
Ewald, C.Y.; Castillo-Quan, J.I.; Blackwell, T.K. Untangling Longevity, Dauer, and Healthspan in Caenorhabditis elegans Insulin/IGF-
1-Signalling. Gerontology 2018,64, 96–104. [CrossRef]
100.
Zhang, S.O.; Trimble, R.; Guo, F.; Mak, H.Y. Lipid droplets as ubiquitous fat storage organelles in C. elegans.BMC Cell Biol.
2010
,
11, 96. [CrossRef]
101.
Wang, P.; Liu, B.; Zhang, D.; Belew, M.Y.; Tissenbaum, H.A.; Cheng, J.X. Imaging lipid metabolism in live Caenorhabditis elegans
using fingerprint vibrations. Angew. Chem. Int. Ed. 2014,53, 11787–11792. [CrossRef] [PubMed]
102. Srinivasan, S. Regulation of body fat in Caenorhabditis elegans.Annu. Rev. Physiol. 2015,77, 161–178. [CrossRef] [PubMed]
103.
Mak, H.Y. Lipid droplets as fat storage organelles in Caenorhabditis elegans: Thematic Review Series: Lipid Droplet Synthesis and
Metabolism: From Yeast to Man. J. Lipid Res. 2012,53, 28–33. [CrossRef] [PubMed]
104.
McCay, C.M.; Crowell, M.F.; Maynard, L.A. The effect of retarded growth upon the length of life span and upon the ultimate
body size. J. Nutr. 1935,10, 63–79. [CrossRef]
105. Partridge, L.; Piper, M.D.; Mair, W. Dietary restriction in Drosophila.Mech. Ageing Dev. 2005,126, 938–950. [CrossRef] [PubMed]
106.
Mattison, J.A.; Colman, R.J.; Beasley, T.M.; Allison, D.B.; Kemnitz, J.W.; Roth, G.S.; Ingram, D.K.; Weindruch, R.; de Cabo, R.;
Anderson, R.M. Caloric restriction improves health and survival of rhesus monkeys. Nat. Commun.
2017
,8, 14063. [CrossRef]
[PubMed]
107.
McGhee, J.D. The C. elegans intestine. In WormBook: The Online Review of C. elegans Biology; WormBook: Pasadena, CA, USA, 2007.
108.
Klass, M.R. Aging in Nematode Caenorhabditis-elegans—Major Biological and Environmental-Factors Influencing Life-Span. Mech.
Ageing Dev. 1977,6, 413–429. [CrossRef]
109.
Hosono, R.; Nishimoto, S.; Kuno, S. Alterations of life span in the nematode Caenorhabditis elegans under monoxenic culture
conditions. Exp. Gerontol. 1989,24, 251–264. [CrossRef] [PubMed]
110.
Mullaney, B.C.; Ashrafi, K. C. elegans fat storage and metabolic regulation. Biochim. Biophys. Acta
2009
,1791, 474–478. [CrossRef]
111.
Wu, Z.; Isik, M.; Moroz, N.; Steinbaugh, M.J.; Zhang, P.; Blackwell, T.K. Dietary Restriction Extends Lifespan through Metabolic
Regulation of Innate Immunity. Cell Metab. 2019,29, 1192–1205.e1198. [CrossRef] [PubMed]
112.
Houthoofd, K.; Gems, D.; Johnson, T.E.; Vanfleteren, J.R. Dietary restriction in the nematode Caenorhabditis elegans.Interdiscip. Top.
Gerontol. 2007,35, 98–114. [CrossRef]
113.
Walker, G.; Houthoofd, K.; Vanfleteren, J.R.; Gems, D. Dietary restriction in C. elegans: From rate-of-living effects to nutrient
sensing pathways. Mech. Ageing Dev. 2005,126, 929–937. [CrossRef]
114.
McElwee, J.; Bubb, K.; Thomas, J.H. Transcriptional outputs of the Caenorhabditis elegans forkhead protein DAF-16. Aging Cell
2003,2, 111–121. [CrossRef]
115.
Sun, X.; Chen, W.D.; Wang, Y.D. DAF-16/FOXO Transcription Factor in Aging and Longevity. Front. Pharmacol.
2017
,8, 548.
[CrossRef] [PubMed]
116.
Baumeister, R.; Schaffitzel, E.; Hertweck, M. Endocrine signaling in Caenorhabditis elegans controls stress response and longevity.
J. Endocrinol. 2006,190, 191–202. [CrossRef] [PubMed]
117.
Kenyon, C.; Chang, J.; Gensch, E.; Rudner, A.; Tabtiang, R. A C. elegans Mutant That Lives Twice as Long as Wild-Type. Nature
1993,366, 461–464. [CrossRef]
118.
Paradis, S.; Ailion, M.; Toker, A.; Thomas, J.H.; Ruvkun, G. A PDK1 homolog is necessary and sufficient to transduce AGE-1 PI3
kinase signals that regulate diapause in Caenorhabditis elegans.Gene Dev. 1999,13, 1438–1452. [CrossRef]
119.
Ogg, S.; Paradis, S.; Gottlieb, S.; Patterson, G.I.; Lee, L.; Tissenbaum, H.A.; Ruvkun, G. The Fork head transcription factor DAF-16
transduces insulin-like metabolic and longevity signals in C. elegans.Nature 1997,389, 994–999. [CrossRef]
120.
Zecic, A.; Braeckman, B.P. DAF-16/FoxO in Caenorhabditis elegans and Its Role in Metabolic Remodeling. Cells
2020
,9, 109.
[CrossRef]
121.
Blackwell, T.K.; Sewell, A.K.; Wu, Z.; Han, M. TOR Signaling in Caenorhabditis elegans Development, Metabolism, and Aging.
Genetics 2019,213, 329–360. [CrossRef] [PubMed]
122. Jung, C.H.; Ro, S.H.; Cao, J.; Otto, N.M.; Kim, D.H. mTOR regulation of autophagy. FEBS Lett. 2010,584, 1287–1295. [CrossRef]
123.
Inoki, K.; Li, Y.; Zhu, T.Q.; Wu, J.; Guan, K.L. TSC2 is phosphorylated and inhibited by Akt and suppresses mTOR signalling. Nat.
Cell Biol. 2002,4, 648–657. [CrossRef]
124. Dibble, C.C.; Cantley, L.C. Regulation of mTORC1 by PI3K signaling. Trends Cell Biol. 2015,25, 545–555. [CrossRef]

Biomolecules 2023,13, 912 43 of 56
125.
Lapierre, L.R.; Hansen, M. Lessons from C. elegans: Signaling pathways for longevity. Trends Endocrinol. Metab.
2012
,23, 637–644.
[CrossRef] [PubMed]
126.
Gingras, A.C.; Gygi, S.P.; Raught, B.; Polakiewicz, R.D.; Abraham, R.T.; Hoekstra, M.F.; Aebersold, R.; Sonenberg, N. Regulation
of 4E-BP1 phosphorylation: A novel two-step mechanism. Genes Dev. 1999,13, 1422–1437. [CrossRef] [PubMed]
127. Powers, T. TOR signaling and S6 kinase 1: Yeast catches up. Cell Metab. 2007,6, 1–2. [CrossRef]
128.
Vellai, T.; Takacs-Vellai, K.; Zhang, Y.; Kovacs, A.L.; Orosz, L.; Muller, F. Genetics: Influence of TOR kinase on lifespan in C. elegans.
Nature 2003,426, 620. [CrossRef] [PubMed]
129.
Robida-Stubbs, S.; Glover-Cutter, K.; Lamming, D.W.; Mizunuma, M.; Narasimhan, S.D.; Neumann-Haefelin, E.; Sabatini, D.M.;
Blackwell, T.K. TOR Signaling and Rapamycin Influence Longevity by Regulating SKN-1/Nrf and DAF-16/FoxO. Cell Metab.
2012,15, 713–724. [CrossRef]
130.
Honjoh, S.; Yamamoto, T.; Uno, M.; Nishida, E. Signalling through RHEB-1 mediates intermittent fasting-induced longevity in
C. elegans.Nature 2009,457, 726–730. [CrossRef]
131.
Hansen, M.; Taubert, S.; Crawford, D.; Libina, N.; Lee, S.J.; Kenyon, C. Lifespan extension by conditions that inhibit translation in
Caenorhabditis elegans.Aging Cell 2007,6, 95–110. [CrossRef]
132.
Jones, K.T.; Greer, E.R.; Pearce, D.; Ashrafi, K. Rictor/TORC2 regulates Caenorhabditis elegans fat storage, body size, and
development through sgk-1. PLoS Biol. 2009,7, e60. [CrossRef]
133.
Soukas, A.A.; Kane, E.A.; Carr, C.E.; Melo, J.A.; Ruvkun, G. Rictor/TORC2 regulates fat metabolism, feeding, growth, and life
span in Caenorhabditis elegans.Genes Dev. 2009,23, 496–511. [CrossRef]
134.
Hara, K.; Maruki, Y.; Long, X.; Yoshino, K.; Oshiro, N.; Hidayat, S.; Tokunaga, C.; Avruch, J.; Yonezawa, K. Raptor, a binding
partner of target of rapamycin (TOR), mediates TOR action. Cell 2002,110, 177–189. [CrossRef]
135.
Long, L.; Liu, W.; Ruan, P.; Yang, X.; Chen, X.; Li, L.; Yuan, F.; He, D.; Huang, P.; Gong, A.; et al. Visualizing the Interplay of
Lipid Droplets and Protein Aggregates During Aging via a Dual-Functional Fluorescent Probe. Anal. Chem.
2022
,94, 2803–2811.
[CrossRef] [PubMed]
136.
Fletcher, M.; Kim, D.H. Age-Dependent Neuroendocrine Signaling from Sensory Neurons Modulates the Effect of Dietary
Restriction on Longevity of Caenorhabditis elegans.PLoS Genet. 2017,13, e1006544. [CrossRef] [PubMed]
137. Gumienny, T.L.; Savage-Dunn, C. TGF-Beta Signaling in C. elegans; WormBook: Pasadena, CA, USA, 2013; pp. 1–34. [CrossRef]
138.
Liu, T.; Zimmerman, K.K.; Patterson, G.I. Regulation of signaling genes by TGFbeta during entry into dauer diapause in C. elegans.
BMC Dev. Biol. 2004,4, 11. [CrossRef] [PubMed]
139.
Hu, M.H.; Crossman, D.; Prasain, J.K.; Miller, M.A.; Serra, R.A. Transcriptomic Profiling of DAF-7/TGF beta Pathway Mutants in
C. elegans.Genes 2020,11, 288. [CrossRef]
140.
Lant, B.; Storey, K.B. An Overview of Stress Response and Hypometabolic Strategies in Caenorhabditis elegans: Conserved and
Contrasting Signals with the Mammalian System. Int. J. Biol. Sci. 2010,6, 9–50. [CrossRef]
141.
Shaw, W.M.; Luo, S.; Landis, J.; Ashraf, J.; Murphy, C.T. The C. elegans TGF-beta Dauer pathway regulates longevity via insulin
signaling. Curr. Biol. 2007,17, 1635–1645. [CrossRef]
142.
Greer, E.R.; Perez, C.L.; Van Gilst, M.R.; Lee, B.H.; Ashrafi, K. Neural and molecular dissection of a C. elegans sensory circuit that
regulates fat and feeding. Cell Metab. 2008,8, 118–131. [CrossRef]
143.
Rashid, S.; Pho, K.B.; Mesbahi, H.; MacNeil, L.T. Nutrient Sensing and Response Drive Developmental Progression in Caenorhab-
ditis elegans.Bioessays 2020,42, e1900194. [CrossRef]
144.
Kumar, A.V.; Mills, J.; Parker, W.M.; Leitão, J.A.; Rodriguez, D.I.; Ng, C.; Patel, R.; Aguilera, J.L.; Johnson, J.R.; Wong, S.Q.; et al.
Lipid droplets modulate proteostasis, SQST-1/SQSTM1 dynamics, and lifespan in C. elegans.bioRxiv
2022
, 2021.04.22.440991.
[CrossRef]
145.
Castillo-Quan, J.I.; Steinbaugh, M.J.; Fernández-Cárdenas, L.P.; Pohl, N.K.; Wu, Z.; Zhu, F.; Moroz, N.; Teixeira, V.; Bland, M.S.;
Lehrbach, N.J.; et al. An anti-steatosis response regulated by oleic acid through lipid droplet-mediated ERAD enhancement.
bioRxiv 2022, 2022.06.15.496302. [CrossRef]
146.
Na, H.; Zhang, P.; Chen, Y.; Zhu, X.; Liu, Y.; Liu, Y.; Xie, K.; Xu, N.; Yang, F.; Yu, Y.; et al. Identification of lipid droplet
structure-like/resident proteins in Caenorhabditis elegans.Biochim. Biophys. Acta (BBA)—Mol. Cell Res.
2015
,1853, 2481–2491.
[CrossRef]
147.
Chughtai, A.A.; Kaššák, F.; Kostrouchová, M.; Novotný, J.P.; Krause, M.W.; Saudek, V.; Kostrouch, Z.; Kostrouchová, M.
Perilipin-related protein regulates lipid metabolism in C. elegans.PeerJ 2015,3, e1213. [CrossRef] [PubMed]
148.
Choudhary, V.; Ojha, N.; Golden, A.; Prinz, W.A. A conserved family of proteins facilitates nascent lipid droplet budding from the
ER. J. Cell Biol. 2015,211, 261–271. [CrossRef] [PubMed]
149.
Papsdorf, K.; Miklas, J.W.; Hosseini, A.; Cabruja, M.; Morrow, C.S.; Savini, M.; Yu, Y.; Silva-García, C.G.; Haseley, N.R.;
Murphy, L.M.; et al. Lipid droplets and peroxisomes are co-regulated to drive lifespan extension in response to mono-unsaturated
fatty acids. Nat. Cell Biol. 2023,25, 672–684. [CrossRef] [PubMed]
150.
Mosquera, J.V.; Bacher, M.C.; Priess, J.R. Nuclear lipid droplets and nuclear damage in Caenorhabditis elegans.PLoS Genet.
2021
,
17, e1009602. [CrossRef]
151. He, Y.; Jasper, H. Studying aging in Drosophila.Methods 2014,68, 129–133. [CrossRef]
152.
Piper, M.D.W.; Partridge, L. Drosophila as a model for ageing. Biochim. Biophys. Acta Mol. Basis Dis.
2018
,1864, 2707–2717.
[CrossRef]

Biomolecules 2023,13, 912 44 of 56
153.
Proshkina, E.N.; Shaposhnikov, M.V.; Sadritdinova, A.F.; Kudryavtseva, A.V.; Moskalev, A.A. Basic mechanisms of longevity: A
case study of Drosophila pro-longevity genes. Ageing Res. Rev. 2015,24, 218–231. [CrossRef]
154.
Liao, S.; Amcoff, M.; Nässel, D.R. Impact of high-fat diet on lifespan, metabolism, fecundity and behavioral senescence in
Drosophila.Insect Biochem. Mol. Biol. 2021,133, 103495. [CrossRef]
155.
Hofbauer, H.F.; Heier, C.; Sen Saji, A.K.; Kühnlein, R.P. Lipidome remodeling in aging normal and genetically obese Drosophila
males. Insect Biochem. Mol. Biol. 2021,133, 103498. [CrossRef]
156.
Buszczak, M.; Lu, X.; Segraves, W.A.; Chang, T.Y.; Cooley, L. Mutations in the midway gene disrupt a Drosophila acyl coenzyme A:
Diacylglycerol acyltransferase. Genetics 2002,160, 1511–1518. [CrossRef]
157.
Gronke, S.; Mildner, A.; Fellert, S.; Tennagels, N.; Petry, S.; Muller, G.; Jackle, H.; Kuhnlein, R.P. Brummer lipase is an evolutionary
conserved fat storage regulator in Drosophila.Cell Metab. 2005,1, 323–330. [CrossRef] [PubMed]
158.
Song, Y.; Park, J.O.; Tanner, L.; Nagano, Y.; Rabinowitz, J.D.; Shvartsman, S.Y. Energy budget of Drosophila embryogenesis. Curr.
Biol. 2019,29, R566–R567. [CrossRef] [PubMed]
159.
Tennessen, J.M.; Barry, W.E.; Cox, J.; Thummel, C.S. Methods for studying metabolism in Drosophila.Methods
2014
,68, 105–115.
[CrossRef] [PubMed]
160.
Bickel, P.E.; Tansey, J.T.; Welte, M.A. PAT proteins, an ancient family of lipid droplet proteins that regulate cellular lipid stores.
Biochim. Biophys. Acta 2009,1791, 419–440. [CrossRef]
161.
Miura, S.; Gan, J.W.; Brzostowski, J.; Parisi, M.J.; Schultz, C.J.; Londos, C.; Oliver, B.; Kimmel, A.R. Functional conservation
for lipid storage droplet association among Perilipin, ADRP, and TIP47 (PAT)-related proteins in mammals, Drosophila, and
Dictyostelium. J. Biol. Chem. 2002,277, 32253–32257. [CrossRef]
162.
Teixeira, L.; Rabouille, C.; Rørth, P.; Ephrussi, A.; Vanzo, N.F. Drosophila Perilipin/ADRP homologue Lsd2 regulates lipid
metabolism. Mech. Dev. 2003,120, 1071–1081. [CrossRef]
163.
Beller, M.; Bulankina, A.V.; Hsiao, H.H.; Urlaub, H.; Jackle, H.; Kuhnlein, R.P. PERILIPIN-dependent control of lipid droplet
structure and fat storage in Drosophila.Cell Metab. 2010,12, 521–532. [CrossRef]
164.
Grönke, S.; Beller, M.; Fellert, S.; Ramakrishnan, H.; Jäckle, H.; Kühnlein, R.P. Control of fat storage by a Drosophila PAT domain
protein. Curr. Biol. 2003,13, 603–606. [CrossRef]
165.
Kayukawa, T.; Jouraku, A.; Ito, Y.; Shinoda, T. Molecular mechanism underlying juvenile hormone-mediated repression of
precocious larval-adult metamorphosis. Proc. Natl. Acad. Sci. USA 2017,114, 1057–1062. [CrossRef]
166.
Mirth, C.K.; Shingleton, A.W. Integrating body and organ size in Drosophila: Recent advances and outstanding problems. Front.
Endocrinol. 2012,3, 49. [CrossRef]
167.
Mirth, C.K.; Tang, H.Y.; Makohon-Moore, S.C.; Salhadar, S.; Gokhale, R.H.; Warner, R.D.; Koyama, T.; Riddiford, L.M.; Shingleton,
A.W. Juvenile hormone regulates body size and perturbs insulin signaling in Drosophila.Proc. Natl. Acad. Sci. USA
2014
,
111, 7018–7023. [CrossRef]
168.
Texada, M.J.; Malita, A.; Christensen, C.F.; Dall, K.B.; Faergeman, N.J.; Nagy, S.; Halberg, K.A.; Rewitz, K. Autophagy-Mediated
Cholesterol Trafficking Controls Steroid Production. Dev. Cell 2019,48, 659–671.e654. [CrossRef] [PubMed]
169.
Werthebach, M.; Stewart, F.A.; Gahlen, A.; Mettler-Altmann, T.; Akhtar, I.; Maas-Enriquez, K.; Droste, A.; Eichmann, T.O.;
Poschmann, G.; Stuhler, K.; et al. Control of Drosophila Growth and Survival by the Lipid Droplet-Associated Protein
CG9186/Sturkopf. Cell Rep. 2019,26, 3726–3740.e3727. [CrossRef] [PubMed]
170.
Thiel, K.; Heier, C.; Haberl, V.; Thul, P.J.; Oberer, M.; Lass, A.; Jäckle, H.; Beller, M. The evolutionarily conserved protein CG9186
is associated with lipid droplets, required for their positioning and for fat storage. J. Cell Sci. 2013,126, 2198–2212. [CrossRef]
171.
Ugrankar, R.; Bowerman, J.; Hariri, H.; Chandra, M.; Chen, K.; Bossanyi, M.F.; Datta, S.; Rogers, S.; Eckert, K.M.; Vale, G.; et al.
Drosophila Snazarus Regulates a Lipid Droplet Population at Plasma Membrane-Droplet Contacts in Adipocytes. Dev. Cell 2019,
50, 557–572.e555. [CrossRef] [PubMed]
172.
Blumrich, A.; Vogler, G.; Dresen, S.; Diop, S.B.; Jaeger, C.; Leberer, S.; Grune, J.; Wirth, E.K.; Hoeft, B.; Renko, K.; et al. Fat-body
brummer lipase determines survival and cardiac function during starvation in Drosophila melanogaster. Iscience
2021
,24, 102288.
[CrossRef] [PubMed]
173.
Bi, J.; Xiang, Y.; Chen, H.; Liu, Z.; Grönke, S.; Kühnlein, R.P.; Huang, X. Opposite and redundant roles of the two Drosophila
perilipins in lipid mobilization. J. Cell Sci. 2012,125, 3568–3577. [CrossRef]
174.
Binh, T.D.; Nguyen, Y.D.H.; Pham, T.L.A.; Komori, K.; Nguyen, T.Q.C.; Taninaka, M.; Kamei, K. Dysfunction of lipid storage
droplet-2 suppresses endoreplication and induces JNK pathway-mediated apoptotic cell death in Drosophila salivary glands. Sci.
Rep. 2022,12, 4302. [CrossRef]
175.
Fauny, J.D.; Silber, J.; Zider, A. Drosophila Lipid Storage Droplet 2 gene (Lsd-2) is expressed and controls lipid storage in wing
imaginal discs. Dev. Dyn. 2005,232, 725–732. [CrossRef]
176.
Binh, T.D.; Pham, T.L.A.; Men, T.T.; Dang, T.T.P.; Kamei, K. LSD-2 dysfunction induces dFoxO-dependent cell death in the wing
of Drosophila melanogaster. Biochem. Biophys. Res. Commun. 2019,509, 491–497. [CrossRef]
177.
Goyal, L.; McCall, K.; Agapite, J.; Hartwieg, E.; Steller, H. Induction of apoptosis by Drosophila reaper, hid and grim through
inhibition of IAP function. EMBO J. 2000,19, 589–597. [CrossRef] [PubMed]
178.
Eckelman, B.P.; Salvesen, G.S.; Scott, F.L. Human inhibitor of apoptosis proteins: Why XIAP is the black sheep of the family.
EMBO Rep. 2006,7, 988–994. [CrossRef] [PubMed]
179. Hanifeh, M.; Ataei, F. XIAP as a multifaceted molecule in Cellular Signaling. Apoptosis 2022,27, 441–453. [CrossRef]

Biomolecules 2023,13, 912 45 of 56
180.
Lu, M.; Lin, S.C.; Huang, Y.; Kang, Y.J.; Rich, R.; Lo, Y.C.; Myszka, D.; Han, J.; Wu, H. XIAP induces NF-kappaB activation via the
BIR1/TAB1 interaction and BIR1 dimerization. Mol. Cell. 2007,26, 689–702. [CrossRef] [PubMed]
181.
Yan, Y.; Wang, H.; Hu, M.; Jiang, L.; Wang, Y.; Liu, P.; Liang, X.; Liu, J.; Li, C.; Lindström-Battle, A.; et al. HDAC6 Suppresses
Age-Dependent Ectopic Fat Accumulation by Maintaining the Proteostasis of PLIN2 in Drosophila.Dev. Cell
2017
,43, 99–111.e115.
[CrossRef] [PubMed]
182.
Katewa, S.D.; Akagi, K.; Bose, N.; Rakshit, K.; Camarella, T.; Zheng, X.; Hall, D.; Davis, S.; Nelson, C.S.; Brem, R.B.; et al.
Peripheral Circadian Clocks Mediate Dietary Restriction-Dependent Changes in Lifespan and Fat Metabolism in Drosophila.Cell
Metab. 2016,23, 143–154. [CrossRef] [PubMed]
183. Katewa, S.D.; Kapahi, P. Dietary restriction and aging, 2009. Aging Cell 2010,9, 105–112. [CrossRef]
184. Skorupa, D.A.; Dervisefendic, A.; Zwiener, J.; Pletcher, S.D. Dietary composition specifies consumption, obesity, and lifespan in
Drosophila melanogaster. Aging Cell 2008,7, 478–490. [CrossRef]
185.
Solon-Biet, S.M.; McMahon, A.C.; Ballard, J.W.; Ruohonen, K.; Wu, L.E.; Cogger, V.C.; Warren, A.; Huang, X.; Pichaud, N.;
Melvin, R.G.; et al. The ratio of macronutrients, not caloric intake, dictates cardiometabolic health, aging, and longevity in ad
libitum-fed mice. Cell Metab. 2014,19, 418–430. [CrossRef]
186.
Puig, O.; Marr, M.T.; Ruhf, M.L.; Tjian, R. Control of cell number by Drosophila FOXO: Downstream and feedback regulation of
the insulin receptor pathway. Genes Dev. 2003,17, 2006–2020. [CrossRef] [PubMed]
187.
Slack, C.; Giannakou, M.E.; Foley, A.; Goss, M.; Partridge, L. dFOXO-independent effects of reduced insulin-like signaling in
Drosophila.Aging Cell 2011,10, 735–748. [CrossRef]
188.
Martins, R.; Lithgow, G.J.; Link, W. Long live FOXO: Unraveling the role of FOXO proteins in aging and longevity. Aging Cell
2016,15, 196–207. [CrossRef] [PubMed]
189.
Wang, M.C.; Bohmann, D.; Jasper, H. JNK signaling confers tolerance to oxidative stress and extends lifespan in Drosophila.Dev.
Cell 2003,5, 811–816. [CrossRef]
190.
Yamamoto, R.; Tatar, M. Insulin receptor substrate chico acts with the transcription factor FOXO to extend Drosophila lifespan.
Aging Cell 2011,10, 729–732. [CrossRef]
191.
Alic, N.; Giannakou, M.E.; Papatheodorou, I.; Hoddinott, M.P.; Andrews, T.D.; Bolukbasi, E.; Partridge, L. Interplay of dFOXO and
two ETS-family transcription factors determines lifespan in Drosophila melanogaster. PLoS Genet.
2014
,10, e1004619. [CrossRef]
[PubMed]
192.
Min, K.J.; Yamamoto, R.; Buch, S.; Pankratz, M.; Tatar, M. Drosophila lifespan control by dietary restriction independent of
insulin-like signaling. Aging Cell 2008,7, 199–206. [CrossRef]
193.
Gronke, S.; Clarke, D.F.; Broughton, S.; Andrews, T.D.; Partridge, L. Molecular evolution and functional characterization of
Drosophila insulin-like peptides. PLoS Genet. 2010,6, e1000857. [CrossRef]
194.
Vereshchagina, N.; Wilson, C. Cytoplasmic activated protein kinase Akt regulates lipid-droplet accumulation in Drosophila nurse
cells. Development 2006,133, 4731–4735. [CrossRef]
195.
DiAngelo, J.R.; Birnbaum, M.J. Regulation of fat cell mass by insulin in Drosophila melanogaster. Mol. Cell. Biol.
2009
,29,
6341–6352. [CrossRef]
196.
Wang, B.; Moya, N.; Niessen, S.; Hoover, H.; Mihaylova, M.M.; Shaw, R.J.; Yates, J.R.; Fischer, W.H.; Thomas, J.B.; Montminy, M. A
Hormone-Dependent Module Regulating Energy Balance. Cell 2011,145, 596–606. [CrossRef]
197.
Biteau, B.; Karpac, J.; Supoyo, S.; Degennaro, M.; Lehmann, R.; Jasper, H. Lifespan extension by preserving proliferative
homeostasis in Drosophila.PLoS Genet. 2010,6, e1001159. [CrossRef]
198.
Wang, L.; Zeng, X.; Ryoo, H.D.; Jasper, H. Integration of UPRER and oxidative stress signaling in the control of intestinal stem cell
proliferation. PLoS Genet. 2014,10, e1004568. [CrossRef] [PubMed]
199.
Wang, L.; Ryoo, H.D.; Qi, Y.; Jasper, H. PERK Limits Drosophila Lifespan by Promoting Intestinal Stem Cell Proliferation in
Response to ER Stress. PLoS Genet. 2015,11, e1005220. [CrossRef] [PubMed]
200.
Luis, N.M.; Wang, L.F.; Ortega, M.; Deng, H.S.; Katewa, S.D.; Li, P.W.L.; Karpac, J.; Jasper, H.; Kapahi, P. Intestinal IRE1 Is
Required for Increased Triglyceride Metabolism and Longer Lifespan under Dietary Restriction. Cell Rep.
2016
,17, 1207–1216.
[CrossRef] [PubMed]
201.
Bailey, A.P.; Koster, G.; Guillermier, C.; Hirst, E.M.; MacRae, J.I.; Lechene, C.P.; Postle, A.D.; Gould, A.P. Antioxidant Role for
Lipid Droplets in a Stem Cell Niche of Drosophila.Cell 2015,163, 340–353. [CrossRef]
202.
Cermelli, S.; Guo, Y.; Gross, S.P.; Welte, M.A. The lipid-droplet proteome reveals that droplets are a protein-storage depot. Curr.
Biol. 2006,16, 1783–1795. [CrossRef] [PubMed]
203.
Beller, M.; Riedel, D.; Jansch, L.; Dieterich, G.; Wehland, J.; Jackle, H.; Kuhnlein, R.P. Characterization of the Drosophila lipid
droplet subproteome. Mol. Cell. Proteom. 2006,5, 1082–1094. [CrossRef]
204.
Kapahi, P.; Zid, B.M.; Harper, T.; Koslover, D.; Sapin, V.; Benzer, S. Regulation of lifespan in Drosophila by modulation of genes in
the TOR signaling pathway. Curr. Biol. 2004,14, 885–890. [CrossRef]
205. Taylor, R.C.; Dillin, A. Aging as an event of proteostasis collapse. Cold Spring Harb. Perspect. Biol. 2011,3, a004440. [CrossRef]
206.
Marshall, L.; Rideout, E.J.; Grewal, S.S. Nutrient/TOR-dependent regulation of RNA polymerase III controls tissue and organismal
growth in Drosophila.EMBO J. 2012,31, 1916–1930. [CrossRef] [PubMed]
207.
Bjedov, I.; Toivonen, J.M.; Kerr, F.; Slack, C.; Jacobson, J.; Foley, A.; Partridge, L. Mechanisms of life span extension by rapamycin
in the fruit fly Drosophila melanogaster. Cell Metab. 2010,11, 35–46. [CrossRef]

Biomolecules 2023,13, 912 46 of 56
208.
Reeg, S.; Grune, T. Protein Oxidation in Toxicology. In Studies on Experimental Toxicology and Pharmacology; Oxidative Stress in
Applied Basic Research and Clinical Practice; Humana Press: Cham, Switzerland, 2015; pp. 81–102. [CrossRef]
209.
Simonsen, A.; Cumming, R.C.; Brech, A.; Isakson, P.; Schubert, D.R.; Finley, K.D. Promoting basal levels of autophagy in the
nervous system enhances longevity and oxidant resistance in adult Drosophila.Autophagy 2008,4, 176–184. [CrossRef]
210.
Kovacs, T.; Billes, V.; Komlos, M.; Hotzi, B.; Manzeger, A.; Tarnoci, A.; Papp, D.; Szikszai, F.; Szinyakovics, J.; Racz, A.; et al. The
small molecule AUTEN-99 (autophagy enhancer-99) prevents the progression of neurodegenerative symptoms. Sci. Rep.
2017
,
7, 42014. [CrossRef] [PubMed]
211.
Papp, D.; Kovács, T.; Billes, V.; Varga, M.; Tarnóci, A.; Hackler, L., Jr.; Puskás, L.G.; Liliom, H.; Tárnok, K.; Schlett, K.; et al.
AUTEN-67, an autophagy-enhancing drug candidate with potent antiaging and neuroprotective effects. Autophagy
2016
,
12, 273–286. [CrossRef]
212.
Eisenberg, T.; Knauer, H.; Schauer, A.; Büttner, S.; Ruckenstuhl, C.; Carmona-Gutierrez, D.; Ring, J.; Schroeder, S.; Magnes, C.;
Antonacci, L.; et al. Induction of autophagy by spermidine promotes longevity. Nat. Cell Biol. 2009,11, 1305–1314. [CrossRef]
213.
Pang, X.; Zhang, X.; Jiang, Y.; Su, Q.; Li, Q.; Li, Z. Autophagy: Mechanisms and Therapeutic Potential of Flavonoids in Cancer.
Biomolecules 2021,11, 135. [CrossRef]
214.
Fantin, M.; Garelli, F.; Napoli, B.; Forgiarini, A.; Gumeni, S.; De Martin, S.; Montopoli, M.; Vantaggiato, C.; Orso, G. Flavonoids
Regulate Lipid Droplets Biogenesis in Drosophila melanogaster. Nat. Prod. Commun. 2019,14, 1934578X19852430. [CrossRef]
215.
Wongchum, N.; Dechakhamphu, A. Xanthohumol prolongs lifespan and decreases stress-induced mortality in Drosophila
melanogaster. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 2021,244, 108994. [CrossRef]
216.
Gajender; Mazumder, A.; Sharma, A.; Azad, M.A.K. A Comprehensive Review of the Pharmacological Importance of Dietary
Flavonoids as Hepatoprotective Agents. Evid.-Based Complement. Altern. Med. 2023,2023, 4139117. [CrossRef]
217.
Heijnen, C.G.; Haenen, G.R.; Oostveen, R.M.; Stalpers, E.M.; Bast, A. Protection of flavonoids against lipid peroxidation: The
structure activity relationship revisited. Free Radic. Res. 2002,36, 575–581. [CrossRef]
218.
Partridge, L.; Alic, N.; Bjedov, I.; Piper, M.D. Ageing in Drosophila: The role of the insulin/Igf and TOR signalling network. Exp.
Gerontol. 2011,46, 376–381. [CrossRef]
219.
Zid, B.M.; Rogers, A.N.; Katewa, S.D.; Vargas, M.A.; Kolipinski, M.C.; Lu, T.A.; Benzer, S.; Kapahi, P. 4E-BP extends lifespan upon
dietary restriction by enhancing mitochondrial activity in Drosophila.Cell 2009,139, 149–160. [CrossRef] [PubMed]
220.
Vattem, K.M.; Wek, R.C. Reinitiation involving upstream ORFs regulates ATF4 mRNA translation in mammalian cells. Proc. Natl.
Acad. Sci. USA 2004,101, 11269–11274. [CrossRef] [PubMed]
221.
Kang, M.J.; Vasudevan, D.; Kang, K.; Kim, K.; Park, J.E.; Zhang, N.; Zeng, X.; Neubert, T.A.; Marr, M.T., 2nd; Ryoo, H.D. 4E-BP is
a target of the GCN2-ATF4 pathway during Drosophila development and aging. J. Cell Biol. 2017,216, 115–129. [CrossRef]
222.
Srivastava, A.; Lu, J.; Gadalla, D.S.; Hendrich, O.; Grönke, S.; Partridge, L. The Role of GCN2 Kinase in Mediating the Effects of
Amino Acids on Longevity and Feeding Behaviour in Drosophila.Front. Aging 2022,3, 944466. [CrossRef] [PubMed]
223.
Mair, W.; Morantte, I.; Rodrigues, A.P.; Manning, G.; Montminy, M.; Shaw, R.J.; Dillin, A. Lifespan extension induced by AMPK
and calcineurin is mediated by CRTC-1 and CREB. Nature 2011,470, 404–408. [CrossRef]
224.
Stenesen, D.; Suh, J.M.; Seo, J.; Yu, K.; Lee, K.S.; Kim, J.S.; Min, K.J.; Graff, J.M. Adenosine nucleotide biosynthesis and AMPK
regulate adult life span and mediate the longevity benefit of caloric restriction in flies. Cell Metab.
2013
,17, 101–112. [CrossRef]
[PubMed]
225.
Johnson, E.C.; Kazgan, N.; Bretz, C.A.; Forsberg, L.J.; Hector, C.E.; Worthen, R.J.; Onyenwoke, R.; Brenman, J.E. Altered
metabolism and persistent starvation behaviors caused by reduced AMPK function in Drosophila.PLoS ONE
2010
,5, e12799.
[CrossRef]
226.
Gutierrez, E.; Wiggins, D.; Fielding, B.; Gould, A.P. Specialized hepatocyte-like cells regulate Drosophila lipid metabolism. Nature
2007,445, 275–280. [CrossRef]
227.
Triggle, C.R.; Mohammed, I.; Bshesh, K.; Marei, I.; Ye, K.; Ding, H.; MacDonald, R.; Hollenberg, M.D.; Hill, M.A. Metformin: Is it
a drug for all reasons and diseases? Metabolism 2022,133, 155223. [CrossRef] [PubMed]
228.
Anisimov, V.N.; Berstein, L.M.; Egormin, P.A.; Piskunova, T.S.; Popovich, I.G.; Zabezhinski, M.A.; Tyndyk, M.L.; Yurova, M.V.;
Kovalenko, I.G.; Poroshina, T.E.; et al. Metformin slows down aging and extends life span of female SHR mice. Cell Cycle
2008
,
7, 2769–2773. [CrossRef]
229.
Cabreiro, F.; Au, C.; Leung, K.Y.; Vergara-Irigaray, N.; Cocheme, H.M.; Noori, T.; Weinkove, D.; Schuster, E.; Greene, N.D.;
Gems, D. Metformin retards aging in C. elegans by altering microbial folate and methionine metabolism. Cell 2013,153, 228–239.
[CrossRef]
230.
Slack, C.; Foley, A.; Partridge, L. Activation of AMPK by the putative dietary restriction mimetic metformin is insufficient to
extend lifespan in Drosophila.PLoS ONE 2012,7, e47699. [CrossRef] [PubMed]
231.
Kalender, A.; Selvaraj, A.; Kim, S.Y.; Gulati, P.; Brûlé, S.; Viollet, B.; Kemp, B.E.; Bardeesy, N.; Dennis, P.; Schlager, J.J.; et al.
Metformin, independent of AMPK, inhibits mTORC1 in a rag GTPase-dependent manner. Cell Metab.
2010
,11, 390–401.
[CrossRef] [PubMed]
232.
Funakoshi, M.; Tsuda, M.; Muramatsu, K.; Hatsuda, H.; Morishita, S.; Aigaki, T. A gain-of-function screen identifies wdb and lkb1
as lifespan-extending genes in Drosophila.Biochem. Biophys. Res. Commun. 2011,405, 667–672. [CrossRef]
233.
Choi, S.; Lim, D.S.; Chung, J. Feeding and Fasting Signals Converge on the LKB1-SIK3 Pathway to Regulate Lipid Metabolism in
Drosophila.PLoS Genet. 2015,11, e1005263. [CrossRef]

Biomolecules 2023,13, 912 47 of 56
234.
Grandison, R.C.; Piper, M.D.W.; Partridge, L. Amino-acid imbalance explains extension of lifespan by dietary restriction in
Drosophila.Nature 2009,462, 1061–1064. [CrossRef]
235.
Kabil, H.; Kabil, O.; Banerjee, R.; Harshman, L.G.; Pletcher, S.D. Increased transsulfuration mediates longevity and dietary
restriction in Drosophila.Proc. Natl. Acad. Sci. USA 2011,108, 16831–16836. [CrossRef]
236.
Jonsson, W.O.; Margolies, N.S.; Anthony, T.G. Dietary Sulfur Amino Acid Restriction and the Integrated Stress Response:
Mechanistic Insights. Nutrients 2019,11, 1349. [CrossRef]
237.
Richie, J.P., Jr.; Leutzinger, Y.; Parthasarathy, S.; Malloy, V.; Orentreich, N.; Zimmerman, J.A. Methionine restriction increases
blood glutathione and longevity in F344 rats. Faseb J. 1994,8, 1302–1307. [CrossRef]
238.
Miller, R.A.; Buehner, G.; Chang, Y.; Harper, J.M.; Sigler, R.; Smith-Wheelock, M. Methionine-deficient diet extends mouse
lifespan, slows immune and lens aging, alters glucose, T4, IGF-I and insulin levels, and increases hepatocyte MIF levels and stress
resistance. Aging Cell 2005,4, 119–125. [CrossRef]
239.
Ruckenstuhl, C.; Netzberger, C.; Entfellner, I.; Carmona-Gutierrez, D.; Kickenweiz, T.; Stekovic, S.; Gleixner, C.; Schmid, C.; Klug,
L.; Sorgo, A.G.; et al. Lifespan Extension by Methionine Restriction Requires Autophagy-Dependent Vacuolar Acidification. PLoS
Genet. 2014,10, e1004347. [CrossRef]
240.
Malloy, V.L.; Perrone, C.E.; Mattocks, D.A.; Ables, G.P.; Caliendo, N.S.; Orentreich, D.S.; Orentreich, N. Methionine restriction
prevents the progression of hepatic steatosis in leptin-deficient obese mice. Metabolism 2013,62, 1651–1661. [CrossRef]
241.
Perrone, C.E.; Mattocks, D.A.; Jarvis-Morar, M.; Plummer, J.D.; Orentreich, N. Methionine restriction effects on mitochondrial
biogenesis and aerobic capacity in white adipose tissue, liver, and skeletal muscle of F344 rats. Metabolism
2010
,59, 1000–1011.
[CrossRef]
242.
Nakata, K.; Kawase, M.; Ogino, S.; Kinoshita, C.; Murata, H.; Sakaue, T.; Ogata, K.; Ohmori, S. Effects of age on levels of cysteine,
glutathione and related enzyme activities in livers of mice and rats and an attempt to replenish hepatic glutathione level of mouse
with cysteine derivatives. Mech. Ageing Dev. 1996,90, 195–207. [CrossRef] [PubMed]
243.
Chakraborty, P.K.; Xiong, X.; Mustafi, S.B.; Saha, S.; Dhanasekaran, D.; Mandal, N.A.; McMeekin, S.; Bhattacharya, R.; Mukherjee,
P. Role of cystathionine beta synthase in lipid metabolism in ovarian cancer. Oncotarget
2015
,6, 37367–37384. [CrossRef] [PubMed]
244.
Negre-Salvayre, A.; Auge, N.; Ayala, V.; Basaga, H.; Boada, J.; Brenke, R.; Chapple, S.; Cohen, G.; Feher, J.; Grune, T.; et al.
Pathological aspects of lipid peroxidation. Free Radic. Res. 2010,44, 1125–1171. [CrossRef] [PubMed]
245.
Negre-Salvayre, A.; Coatrieux, C.; Ingueneau, C.; Salvayre, R. Advanced lipid peroxidation end products in oxidative damage to
proteins. Potential role in diseases and therapeutic prospects for the inhibitors. Br. J. Pharmacol. 2008,153, 6–20. [CrossRef]
246.
Johnson, D.C.; Dean, D.R.; Smith, A.D.; Johnson, M.K. Structure, function, and formation of biological iron-sulfur clusters. Annu.
Rev. Biochem. 2005,74, 247–281. [CrossRef]
247.
Boal, A.K.; Yavin, E.; Barton, J.K. DNA repair glycosylases with a [4Fe-4S] cluster: A redox cofactor for DNA-mediated charge
transport? J. Inorg. Biochem. 2007,101, 1913–1921. [CrossRef]
248.
Rouault, T.A. Mammalian iron-sulphur proteins: Novel insights into biogenesis and function. Nat. Rev. Mol. Cell Biol.
2015
,
16, 45–55. [CrossRef] [PubMed]
249.
Crooks, D.R.; Maio, N.; Lane, A.N.; Jarnik, M.; Higashi, R.M.; Haller, R.G.; Yang, Y.; Fan, T.W.; Linehan, W.M.; Rouault, T.A. Acute
loss of iron-sulfur clusters results in metabolic reprogramming and generation of lipid droplets in mammalian cells. J. Biol. Chem.
2018,293, 8297–8311. [CrossRef] [PubMed]
250.
Koeppen, A.H. Friedreich’s ataxia: Pathology, pathogenesis, and molecular genetics. J. Neurol. Sci.
2011
,303, 1–12. [CrossRef]
[PubMed]
251.
Martelli, A.; Napierala, M.; Puccio, H. Understanding the genetic and molecular pathogenesis of Friedreich’s ataxia through
animal and cellular models. Dis. Model. Mech. 2012,5, 165–176. [CrossRef]
252.
Puccio, H.; Simon, D.; Cossée, M.; Criqui-Filipe, P.; Tiziano, F.; Melki, J.; Hindelang, C.; Matyas, R.; Rustin, P.; Koenig, M.
Mouse models for Friedreich ataxia exhibit cardiomyopathy, sensory nerve defect and Fe-S enzyme deficiency followed by
intramitochondrial iron deposits. Nat. Genet. 2001,27, 181–186. [CrossRef]
253.
Navarro, J.A.; Ohmann, E.; Sanchez, D.; Botella, J.A.; Liebisch, G.; Moltó, M.D.; Ganfornina, M.D.; Schmitz, G.; Schneuwly, S.
Altered lipid metabolism in a Drosophila model of Friedreich’s ataxia. Hum. Mol. Genet. 2010,19, 2828–2840. [CrossRef]
254.
Bresgen, N.; Eckl, P.M. Oxidative stress and the homeodynamics of iron metabolism. Biomolecules
2015
,5, 808–847. [CrossRef]
[PubMed]
255.
Halliwell, B.; Gutteridge, J.M. Biologically relevant metal ion-dependent hydroxyl radical generation. An update. FEBS Lett.
1992,307, 108–112. [CrossRef]
256.
Schaur, R.J.; Siems, W.; Bresgen, N.; Eckl, P.M. 4-Hydroxy-nonenal-A Bioactive Lipid Peroxidation Product. Biomolecules
2015
,
5, 2247–2337. [CrossRef]
257.
Girard, V.; Goubard, V.; Querenet, M.; Seugnet, L.; Pays, L.; Nataf, S.; Dufourd, E.; Cluet, D.; Mollereau, B.; Davoust, N. Spen
modulates lipid droplet content in adult Drosophila glial cells and protects against paraquat toxicity. Sci. Rep.
2020
,10, 20023.
[CrossRef] [PubMed]
258. Long, M.; McWilliams, T.G. Lipid droplets promote efficient mitophagy. Autophagy 2022,19, 724–725. [CrossRef] [PubMed]
259.
Andersen, D.S.; Colombani, J.; Palmerini, V.; Chakrabandhu, K.; Boone, E.; Röthlisberger, M.; Toggweiler, J.; Basler, K.; Mapelli,
M.; Hueber, A.O.; et al. The Drosophila TNF receptor Grindelwald couples loss of cell polarity and neoplastic growth. Nature
2015
,
522, 482–486. [CrossRef]

Biomolecules 2023,13, 912 48 of 56
260.
Muliyil, S.; Levet, C.; Düsterhöft, S.; Dulloo, I.; Cowley, S.A.; Freeman, M. ADAM17-triggered TNF signalling protects the ageing
Drosophila retina from lipid droplet-mediated degeneration. Embo J. 2020,39, e104415. [CrossRef] [PubMed]
261.
Lagrutta, L.C.; Layerenza, J.P.; Bronsoms, S.; Trejo, S.A.; Ves-Losada, A. Nuclear-lipid-droplet proteome: Carboxylesterase as a
nuclear lipase involved in lipid-droplet homeostasis. Heliyon 2021,7, e06539. [CrossRef]
262.
Lian, J.; Nelson, R.; Lehner, R. Carboxylesterases in lipid metabolism: From mouse to human. Protein Cell
2018
,9, 178–195.
[CrossRef] [PubMed]
263.
Wang, H.; Wei, E.; Quiroga, A.D.; Sun, X.; Touret, N.; Lehner, R. Altered lipid droplet dynamics in hepatocytes lacking
triacylglycerol hydrolase expression. Mol. Biol. Cell 2010,21, 1991–2000. [CrossRef]
264.
Lian, J.H.; Wei, E.H.; Wang, S.P.; Quiroga, A.D.; Li, L.N.; Di Pardo, A.; van der Veen, J.; Sipione, S.; Mitchell, G.A.; Lehner, R. Liver
Specific Inactivation of Carboxylesterase 3/Triacylglycerol Hydrolase Decreases Blood Lipids Without Causing Severe Steatosis
in Mice. Hepatology 2012,56, 2154–2162. [CrossRef]
265.
Fujimoto, T. Nuclear lipid droplets—How are they different from their cytoplasmic siblings? J. Cell Sci.
2022
,135, jcs259253.
[CrossRef]
266.
McPhee, M.J.; Salsman, J.; Foster, J.; Thompson, J.; Mathavarajah, S.; Dellaire, G.; Ridgway, N.D. Running ‘LAPS’ around nLD:
Nuclear Lipid Droplet Form and Function. Front. Cell Dev. Biol. 2022,10, 837406. [CrossRef]
267.
Chen, H.; Zheng, X.; Xiao, D.; Zheng, Y. Age-associated de-repression of retrotransposons in the Drosophila fat body, its potential
cause and consequence. Aging Cell 2016,15, 542–552. [CrossRef]
268.
Wood, J.G.; Jones, B.C.; Jiang, N.; Chang, C.; Hosier, S.; Wickremesinghe, P.; Garcia, M.; Hartnett, D.A.; Burhenn, L.;
Neretti, N.; et al. Chromatin-modifying genetic interventions suppress age-associated transposable element activation and extend
life span in Drosophila.Proc. Natl. Acad. Sci. USA 2016,113, 11277–11282. [CrossRef] [PubMed]
269. Czech, B.; Hannon, G.J. Small RNA sorting: Matchmaking for Argonautes. Nat. Rev. Genet. 2011,12, 19–31. [CrossRef]
270.
Li, W.; Prazak, L.; Chatterjee, N.; Grüninger, S.; Krug, L.; Theodorou, D.; Dubnau, J. Activation of transposable elements during
aging and neuronal decline in Drosophila.Nat. Neurosci. 2013,16, 529–531. [CrossRef] [PubMed]
271.
Li, Z.; Johnson, M.R.; Ke, Z.; Chen, L.; Welte, M.A. Drosophila lipid droplets buffer the H2Av supply to protect early embryonic
development. Curr. Biol. 2014,24, 1485–1491. [CrossRef]
272.
Loyola, A.C.; Zhang, L.; Shang, R.; Dutta, P.; Li, J.; Li, W.X. Identification of methotrexate as a heterochromatin-promoting drug.
Sci. Rep. 2019,9, 11673. [CrossRef] [PubMed]
273.
Wood, J.G.; Hillenmeyer, S.; Lawrence, C.; Chang, C.; Hosier, S.; Lightfoot, W.; Mukherjee, E.; Jiang, N.; Schorl, C.;
Brodsky, A.S.; et al. Chromatin remodeling in the aging genome of Drosophila.Aging Cell 2010,9, 971–978. [CrossRef]
274.
Yang, L.; Ma, Z.; Wang, H.; Niu, K.; Cao, Y.; Sun, L.; Geng, Y.; Yang, B.; Gao, F.; Chen, Z.; et al. Ubiquitylome study identifies
increased histone 2A ubiquitylation as an evolutionarily conserved aging biomarker. Nat. Commun. 2019,10, 2191. [CrossRef]
275.
Maleszewska, M.; Mawer, J.S.P.; Tessarz, P. Histone Modifications in Ageing and Lifespan Regulation. Curr. Mol. Biol. Rep.
2016
,
2, 26–35. [CrossRef]
276.
Feser, J.; Truong, D.; Das, C.; Carson, J.J.; Kieft, J.; Harkness, T.; Tyler, J.K. Elevated histone expression promotes life span extension.
Mol. Cell 2010,39, 724–735. [CrossRef]
277.
Benayoun, B.A.; Pollina, E.A.; Brunet, A. Epigenetic regulation of ageing: Linking environmental inputs to genomic stability. Nat.
Rev. Mol. Cell Biol. 2015,16, 593–610. [CrossRef] [PubMed]
278. Muñoz-Najar, U.; Sedivy, J.M. Epigenetic control of aging. Antioxid. Redox Signal. 2011,14, 241–259. [CrossRef] [PubMed]
279.
Shah, P.P.; Donahue, G.; Otte, G.L.; Capell, B.C.; Nelson, D.M.; Cao, K.; Aggarwala, V.; Cruickshanks, H.A.; Rai, T.S.;
McBryan, T.; et al. Lamin B1 depletion in senescent cells triggers large-scale changes in gene expression and the chromatin
landscape. Genes Dev. 2013,27, 1787–1799. [CrossRef]
280.
Toiber, D.; Erdel, F.; Bouazoune, K.; Silberman, D.M.; Zhong, L.; Mulligan, P.; Sebastian, C.; Cosentino, C.; Martinez-Pastor, B.;
Giacosa, S.; et al. SIRT6 Recruits SNF2H to DNA Break Sites, Preventing Genomic Instability through Chromatin Remodeling.
Mol. Cell 2013,51, 454–468. [CrossRef] [PubMed]
281.
Kim, H.S.; Xiao, C.Y.; Wang, R.H.; Lahusen, T.; Xu, X.L.; Vassilopoulos, A.; Vazquez-Ortiz, G.; Jeong, W.I.; Park, O.; Ki, S.H.; et al.
Hepatic-Specific Disruption of SIRT6 in Mice Results in Fatty Liver Formation Due to Enhanced Glycolysis and Triglyceride
Synthesis. Cell Metab. 2010,12, 224–236. [CrossRef]
282.
Penrose, H.; Heller, S.; Cable, C.; Makboul, R.; Chadalawada, G.; Chen, Y.; Crawford, S.E.; Savkovic, S.D. Epidermal growth
factor receptor mediated proliferation depends on increased lipid droplet density regulated via a negative regulatory loop with
FOXO3/Sirtuin6. Biochem. Biophys. Res. Commun. 2016,469, 370–376. [CrossRef]
283. Eckl, P.M.; Bresgen, N. Genotoxicity of lipid oxidation compounds. Free Radic. Biol. Med. 2017,111, 244–252. [CrossRef]
284.
Shubeita, G.T.; Tran, S.L.; Xu, J.; Vershinin, M.; Cermelli, S.; Cotton, S.L.; Welte, M.A.; Gross, S.P. Consequences of motor copy
number on the intracellular transport of kinesin-1-driven lipid droplets. Cell 2008,135, 1098–1107. [CrossRef]
285. Welte, M.A. Proteins under new management: Lipid droplets deliver. Trends Cell Biol. 2007,17, 363–369. [CrossRef]
286.
Welte, M.A.; Gross, S.P.; Postner, M.; Block, S.M.; Wieschaus, E.F. Developmental regulation of vesicle transport in Drosophila
embryos: Forces and kinetics. Cell 1998,92, 547–557. [CrossRef]
287.
Bartsch, T.F.; Longoria, R.A.; Florin, E.L.; Shubeita, G.T. Lipid droplets purified from Drosophila embryos as an endogenous handle
for precise motor transport measurements. Biophys. J. 2013,105, 1182–1191. [CrossRef]

Biomolecules 2023,13, 912 49 of 56
288.
Larsen, K.S.; Xu, J.; Cermelli, S.; Shu, Z.; Gross, S.P. BicaudalD actively regulates microtubule motor activity in lipid droplet
transport. PLoS ONE 2008,3, e3763. [CrossRef] [PubMed]
289.
Welte, M.A.; Cermelli, S.; Griner, J.; Viera, A.; Guo, Y.; Kim, D.H.; Gindhart, J.G.; Gross, S.P. Regulation of lipid-droplet transport
by the perilipin homolog LSD2. Curr. Biol. 2005,15, 1266–1275. [CrossRef] [PubMed]
290. Jäckle, H.; Jahn, R. Vesicle transport: Klarsicht clears up the matter. Curr. Biol. 1998,8, R542–R544. [CrossRef]
291.
Myat, M.M.; Andrew, D.J. Epithelial tube morphology is determined by the polarized growth and delivery of apical membrane.
Cell 2002,111, 879–891. [CrossRef]
292.
Yu, Y.V.; Li, Z.; Rizzo, N.P.; Einstein, J.; Welte, M.A. Targeting the motor regulator Klar to lipid droplets. BMC Cell Biol.
2011
,12, 9.
[CrossRef] [PubMed]
293.
Guo, Y.; Jangi, S.; Welte, M.A. Organelle-specific control of intracellular transport: Distinctly targeted isoforms of the regulator
Klar. Mol. Biol. Cell 2005,16, 1406–1416. [CrossRef]
294.
Ingle, L.; Wood, T.R.; Banta, A.M. A study of longevity, growth, reproduction and heart rate in Daphnia longispina as influenced
by limitations in quantity of food. J. Exp. Zool. 1937,76, 325–352. [CrossRef]
295. Fanestil, D.D.; Barrows, C.H., Jr. Aging in the rotifer. J. Gerontol. 1965,20, 462–469.
296.
Comfort, A. Effect of Delayed and resumed growth on the longevity of a fish (Lebistes reticulatus, peters) in captivity. Gerontologia
1963,49, 150–155. [CrossRef]
297.
Dilan, C.-B.; Begun, E.; Ahmet Tugrul, O.; Hulusi, K.; Michelle, A. Zebrafish Aging Models and Possible Interventions. In Recent
Advances in Zebrafish Researches; Yusuf, B., Ed.; IntechOpen: Rijeka, Croatia, 2018; pp. 3–26.
298.
Rudzinska, M.A. The influence of amount of food on the reproduction rate and longevity of a sectarian. (Tokophyra infusionum).
Science 1951,113, 10–11. [CrossRef]
299.
Sutphin, G.L.; Kaeberlein, M. Dietary restriction by bacterial deprivation increases life span in wild-derived nematodes. Exp.
Gerontol. 2008,43, 130–135. [CrossRef]
300.
Schulz, T.J.; Zarse, K.; Voigt, A.; Urban, N.; Birringer, M.; Ristow, M. Glucose Restriction Extends Caenorhabditis elegans Life Span
by Inducing Mitochondrial Respiration and Increasing Oxidative Stress. Cell Metab. 2007,6, 280–293. [CrossRef]
301.
Suckow, B.K.; Suckow, M.A. Lifespan extension by the antioxidant curcumin in Drosophila melanogaster. Int. J. Biomed. Sci.
2006
,
2, 402–405.
302.
Mohammed, I.; Hollenberg, M.D.; Ding, H.; Triggle, C.R. A Critical Review of the Evidence That Metformin Is a Putative
Anti-Aging Drug That Enhances Healthspan and Extends Lifespan. Front. Endocrinol. 2021,12, 718942. [CrossRef] [PubMed]
303.
Belenky, P.; Racette, F.G.; Bogan, K.L.; McClure, J.M.; Smith, J.S.; Brenner, C. Nicotinamide riboside promotes Sir2 silencing and
extends lifespan via Nrk and Urh1/Pnp1/Meu1 pathways to NAD+. Cell 2007,129, 473–484. [CrossRef]
304.
Hofer, S.J.; Davinelli, S.; Bergmann, M.; Scapagnini, G.; Madeo, F. Caloric Restriction Mimetics in Nutrition and Clinical Trials.
Front. Nutr. 2021,8, 717343. [CrossRef]
305. Ross, M.H. Length of life and nutrition in the rat. J. Nutr. 1961,75, 197–210. [CrossRef]
306.
Weindruch, R.; Walford, R.L. Dietary restriction in mice beginning at 1 year of age: Effect on life-span and spontaneous cancer
incidence. Science 1982,215, 1415–1418. [CrossRef]
307.
Nedergaard, J.; Bengtsson, T.; Cannon, B. Unexpected evidence for active brown adipose tissue in adult humans. Am. J.
Physiol.-Endocrinol. Metab. 2007,293, E444–E452. [CrossRef] [PubMed]
308.
Darcy, J.; Tseng, Y.-H. ComBATing aging—Does increased brown adipose tissue activity confer longevity? GeroScience
2019
,
41, 285–296. [CrossRef]
309.
Zoico, E.; Rubele, S.; De Caro, A.; Nori, N.; Mazzali, G.; Fantin, F.; Rossi, A.; Zamboni, M. Brown and Beige Adipose Tissue and
Aging. Front. Endocrinol. 2019,10, 368. [CrossRef]
310.
Fabbiano, S.; Suárez-Zamorano, N.; Rigo, D.; Veyrat-Durebex, C.; Stevanovic Dokic, A.; Colin, D.J.; Trajkovski, M. Caloric
Restriction Leads to Browning of White Adipose Tissue through Type 2 Immune Signaling. Cell Metab.
2016
,24, 434–446.
[CrossRef] [PubMed]
311.
Baumeier, C.; Kaiser, D.; Heeren, J.; Scheja, L.; John, C.; Weise, C.; Eravci, M.; Lagerpusch, M.; Schulze, G.; Joost, H.-G.; et al.
Caloric restriction and intermittent fasting alter hepatic lipid droplet proteome and diacylglycerol species and prevent diabetes in
NZO mice. Biochim. Biophys. Acta (BBA)—Mol. Cell Biol. Lipids 2015,1851, 566–576. [CrossRef] [PubMed]
312.
Franceschi, C.; Garagnani, P.; Morsiani, C.; Conte, M.; Santoro, A.; Grignolio, A.; Monti, D.; Capri, M.; Salvioli, S. The Continuum
of Aging and Age-Related Diseases: Common Mechanisms but Different Rates. Front. Med. 2018,5, 61. [CrossRef]
313. van Deursen, J.M. The role of senescent cells in ageing. Nature 2014,509, 439–446. [CrossRef] [PubMed]
314.
Lizardo, D.Y.; Lin, Y.L.; Gokcumen, O.; Atilla-Gokcumen, G.E. Regulation of lipids is central to replicative senescence. Mol.
Biosyst. 2017,13, 498–509. [CrossRef]
315.
Bartke, A.; Darcy, J. GH and ageing: Pitfalls and new insights. Best Pract. Res. Clin. Endocrinol. Metab.
2017
,31, 113–125. [CrossRef]
316.
Vitale, G.; Salvioli, S.; Franceschi, C. Oxidative stress and the ageing endocrine system. Nat. Rev. Endocrinol.
2013
,9, 228–240.
[CrossRef]
317.
Vitale, G.; Cesari, M.; Mari, D. Aging of the endocrine system and its potential impact on sarcopenia. Eur. J. Intern. Med.
2016
,
35, 10–15. [CrossRef]
318.
Sadria, M.; Layton, A.T. Interactions among mTORC, AMPK and SIRT: A computational model for cell energy balance and
metabolism. Cell Commun. Signal. 2021,19, 57. [CrossRef] [PubMed]

Biomolecules 2023,13, 912 50 of 56
319.
Ralhan, I.; Chang, C.L.; Lippincott-Schwartz, J.; Ioannou, M.S. Lipid droplets in the nervous system. J. Cell Biol.
2021
,
220, e202102136. [CrossRef]
320.
Yang, D.S.; Stavrides, P.; Saito, M.; Kumar, A.; Rodriguez-Navarro, J.A.; Pawlik, M.; Huo, C.; Walkley, S.U.; Saito, M.;
Cuervo, A.M.; et al. Defective macroautophagic turnover of brain lipids in the TgCRND8 Alzheimer mouse model: Prevention by
correcting lysosomal proteolytic deficits. Brain 2014,137, 3300–3318. [CrossRef]
321.
Derk, J.; Bermudez Hernandez, K.; Rodriguez, M.; He, M.; Koh, H.; Abedini, A.; Li, H.; Fenyo, D.; Schmidt, A.M. Diaphanous
1 (DIAPH1) is Highly Expressed in the Aged Human Medial Temporal Cortex and Upregulated in Myeloid Cells during
Alzheimer’s Disease. J. Alzheimers Dis. 2018,64, 995–1007. [CrossRef] [PubMed]
322.
Hamilton, L.K.; Dufresne, M.; Joppe, S.E.; Petryszyn, S.; Aumont, A.; Calon, F.; Barnabe-Heider, F.; Furtos, A.; Parent, M.;
Chaurand, P.; et al. Aberrant Lipid Metabolism in the Forebrain Niche Suppresses Adult Neural Stem Cell Proliferation in an
Animal Model of Alzheimer’s Disease. Cell Stem Cell 2015,17, 397–411. [CrossRef] [PubMed]
323.
Qi, G.; Mi, Y.; Shi, X.; Gu, H.; Brinton, R.D.; Yin, F. ApoE4 Impairs Neuron-Astrocyte Coupling of Fatty Acid Metabolism. Cell Rep.
2021,34, 108572. [CrossRef]
324.
Brekk, O.R.; Honey, J.R.; Lee, S.; Hallett, P.J.; Isacson, O. Cell type-specific lipid storage changes in Parkinson’s disease patient
brains are recapitulated by experimental glycolipid disturbance. Proc. Natl. Acad. Sci. USA 2020,117, 27646–27654. [CrossRef]
325.
Cole, N.B.; Murphy, D.D.; Grider, T.; Rueter, S.; Brasaemle, D.; Nussbaum, R.L. Lipid droplet binding and oligomerization
properties of the Parkinson’s disease protein alpha-synuclein. J. Biol. Chem. 2002,277, 6344–6352. [CrossRef] [PubMed]
326. Yako, T.; Otsu, W.; Nakamura, S.; Shimazawa, M.; Hara, H. Lipid Droplet Accumulation Promotes RPE Dysfunction. Int. J. Mol.
Sci. 2022,23, 1790. [CrossRef]
327.
Arbaizar-Rovirosa, M.; Gallizioli, M.; Pedragosa, J.; Lozano, J.J.; Casal, C.; Pol, A.; Planas, A.M. Age-dependent lipid droplet-rich
microglia worsen stroke outcome in old mice. bioRxiv 2022, 2022.2003.2014.484305. [CrossRef]
328.
Plakkal Ayyappan, J.; Paul, A.; Goo, Y.H. Lipid droplet-associated proteins in atherosclerosis (Review). Mol. Med. Rep.
2016
,
13, 4527–4534. [CrossRef]
329.
Sukhorukov, V.N.; Khotina, V.A.; Chegodaev, Y.S.; Ivanova, E.; Sobenin, I.A.; Orekhov, A.N. Lipid Metabolism in Macrophages:
Focus on Atherosclerosis. Biomedicines 2020,8, 262. [CrossRef]
330.
Goldberg, I.J.; Reue, K.; Abumrad, N.A.; Bickel, P.E.; Cohen, S.; Fisher, E.A.; Galis, Z.S.; Granneman, J.G.; Lewandowski, E.D.;
Murphy, R.; et al. Deciphering the Role of Lipid Droplets in Cardiovascular Disease: A Report from the 2017 National Heart,
Lung, and Blood Institute Workshop. Circulation 2018,138, 305–315. [CrossRef] [PubMed]
331.
Huang, W.; Gao, F.; Zhang, Y.; Chen, T.; Xu, C. Lipid Droplet-Associated Proteins in Cardiomyopathy. Ann. Nutr. Metab.
2022
,
78, 1–13. [CrossRef]
332. Al Saedi, A.; Debruin, D.A.; Hayes, A.; Hamrick, M. Lipid metabolism in sarcopenia. Bone 2022,164, 116539. [CrossRef]
333.
Conte, M.; Vasuri, F.; Trisolino, G.; Bellavista, E.; Santoro, A.; Degiovanni, A.; Martucci, E.; D’Errico-Grigioni, A.; Caporossi, D.;
Capri, M.; et al. Increased Plin2 expression in human skeletal muscle is associated with sarcopenia and muscle weakness. PLoS
ONE 2013,8, e73709. [CrossRef]
334.
Weyand, C.M.; Wu, B.; Goronzy, J.J. The metabolic signature of T cells in rheumatoid arthritis. Curr. Opin. Rheumatol.
2020
,
32, 159–167. [CrossRef] [PubMed]
335.
Naiff, P.F.; Kuckelhaus, S.A.S.; Corazza, D.; Leite, L.M.; Couto, S.; deOliveira, M.S.; Santiago, L.M.; Silva, L.F.; Oliveira, L.A.;
Grisi, D.C.; et al. Quantification of lipid bodies in monocytes from patients with periodontitis. Clin. Exp. Dent. Res.
2021
,
7, 93–100. [CrossRef] [PubMed]
336.
Rho, J.H.; Kim, H.J.; Joo, J.Y.; Lee, J.Y.; Lee, J.H.; Park, H.R. Periodontal Pathogens Promote Foam Cell Formation by Blocking
Lipid Efflux. J. Dent. Res. 2021,100, 1367–1377. [CrossRef]
337.
McGee-Lawrence, M.E.; Carpio, L.R.; Schulze, R.J.; Pierce, J.L.; McNiven, M.A.; Farr, J.N.; Khosla, S.; Oursler, M.J.; Westendorf, J.J.
Hdac3 Deficiency Increases Marrow Adiposity and Induces Lipid Storage and Glucocorticoid Metabolism in Osteochondropro-
genitor Cells. J. Bone Miner. Res. 2016,31, 116–128. [CrossRef] [PubMed]
338.
Rendina-Ruedy, E.; Rosen, C.J. Lipids in the Bone Marrow: An Evolving Perspective. Cell Metab.
2020
,31, 219–231. [CrossRef]
[PubMed]
339.
Lippiello, L.; Walsh, T.; Fienhold, M. The association of lipid abnormalities with tissue pathology in human osteoarthritic articular
cartilage. Metabolism 1991,40, 571–576. [CrossRef]
340.
Lee, S.W.; Rho, J.H.; Lee, S.Y.; Chung, W.T.; Oh, Y.J.; Kim, J.H.; Yoo, S.H.; Kwon, W.Y.; Bae, J.Y.; Seo, S.Y.; et al. Dietary fat-associated
osteoarthritic chondrocytes gain resistance to lipotoxicity through PKCK2/STAMP2/FSP27. Bone Res.
2018
,6, 20. [CrossRef]
[PubMed]
341.
Mustonen, A.M.; Nieminen, P. Fatty Acids and Oxylipins in Osteoarthritis and Rheumatoid Arthritis-a Complex Field with
Significant Potential for Future Treatments. Curr. Rheumatol. Rep. 2021,23, 41. [CrossRef]
342.
Tong, X.; Liu, S.; Stein, R.; Imai, Y. Lipid Droplets’ Role in the Regulation of beta-Cell Function and beta-Cell Demise in Type 2
Diabetes. Endocrinology 2022,163, bqac007. [CrossRef]
343.
Tong, X.; Stein, R. Lipid Droplets Protect Human beta-Cells From Lipotoxicity-Induced Stress and Cell Identity Changes. Diabetes
2021,70, 2595–2607. [CrossRef]
344.
He, Y.; Su, Y.; Duan, C.; Wang, S.; He, W.; Zhang, Y.; An, X.; He, M. Emerging role of aging in the progression of NAFLD to HCC.
Ageing Res. Rev. 2023,84, 101833. [CrossRef]

Biomolecules 2023,13, 912 51 of 56
345.
Stephens, N.A.; Skipworth, R.J.; Macdonald, A.J.; Greig, C.A.; Ross, J.A.; Fearon, K.C. Intramyocellular lipid droplets increase
with progression of cachexia in cancer patients. J. Cachexia Sarcopenia Muscle 2011,2, 111–117. [CrossRef] [PubMed]
346.
Nardi, F.; Fitchev, P.; Brooks, K.M.; Franco, O.E.; Cheng, K.; Hayward, S.W.; Welte, M.A.; Crawford, S.E. Lipid droplet velocity is a
microenvironmental sensor of aggressive tumors regulated by V-ATPase and PEDF. Lab. Investig.
2019
,99, 1822–1834. [CrossRef]
[PubMed]
347. Hamsanathan, S.; Gurkar, A.U. Lipids as Regulators of Cellular Senescence. Front. Physiol. 2022,13, 796850. [CrossRef]
348.
Ogrodnik, M.; Zhu, Y.; Langhi, L.G.P.; Tchkonia, T.; Kruger, P.; Fielder, E.; Victorelli, S.; Ruswhandi, R.A.; Giorgadze, N.;
Pirtskhalava, T.; et al. Obesity-Induced Cellular Senescence Drives Anxiety and Impairs Neurogenesis. Cell Metab.
2019
,29,
1061–1077.e1068. [CrossRef]
349.
Chee, W.Y.; Kurahashi, Y.; Kim, J.; Miura, K.; Okuzaki, D.; Ishitani, T.; Kajiwara, K.; Nada, S.; Okano, H.; Okada, M. beta-
catenin-promoted cholesterol metabolism protects against cellular senescence in naked mole-rat cells. Commun. Biol.
2021
,4, 357.
[CrossRef]
350.
Justesen, J.; Stenderup, K.; Ebbesen, E.N.; Mosekilde, L.; Steiniche, T.; Kassem, M. Adipocyte tissue volume in bone marrow is
increased with aging and in patients with osteoporosis. Biogerontology 2001,2, 165–171. [CrossRef]
351.
Verma, S.; Rajaratnam, J.H.; Denton, J.; Hoyland, J.A.; Byers, R.J. Adipocytic proportion of bone marrow is inversely related to
bone formation in osteoporosis. J. Clin. Pathol. 2002,55, 693–698. [CrossRef] [PubMed]
352.
Rosen, C.J.; Bouxsein, M.L. Mechanisms of disease: Is osteoporosis the obesity of bone? Nat. Clin. Pract. Rheumatol.
2006
,2, 35–43.
[CrossRef]
353.
Rendina-Ruedy, E.; Guntur, A.R.; Rosen, C.J. Intracellular lipid droplets support osteoblast function. Adipocyte
2017
,6, 250–258.
[CrossRef]
354.
Diascro, D.D.; Vogel, R.L.; Johnson, T.E.; Witherup, K.M.; Pitzenberger, S.M.; Rutledge, S.J.; Prescott, D.J.; Rodan, G.A.; Schmidt,
A. High fatty acid content in rabbit serum is responsible for the differentiation of osteoblasts into adipocyte-like cells. J. Bone
Miner. Res. 1998,13, 96–106. [CrossRef] [PubMed]
355.
Sebo, Z.L.; Rendina-Ruedy, E.; Ables, G.P.; Lindskog, D.M.; Rodeheffer, M.S.; Fazeli, P.K.; Horowitz, M.C. Bone Marrow Adiposity:
Basic and Clinical Implications. Endocr. Rev. 2019,40, 1187–1206. [CrossRef]
356.
Maurin, A.C.; Chavassieux, P.M.; Frappart, L.; Delmas, P.D.; Serre, C.M.; Meunier, P.J. Influence of mature adipocytes on osteoblast
proliferation in human primary cocultures. Bone 2000,26, 485–489. [CrossRef]
357.
Elbaz, A.; Wu, X.; Rivas, D.; Gimble, J.M.; Duque, G. Inhibition of fatty acid biosynthesis prevents adipocyte lipotoxicity on
human osteoblasts in vitro. J. Cell. Mol. Med. 2010,14, 982–991. [CrossRef]
358.
Backesjo, C.M.; Li, Y.; Lindgren, U.; Haldosen, L.A. Activation of Sirt1 decreases adipocyte formation during osteoblast
differentiation of mesenchymal stem cells. Cells Tissues Organs 2009,189, 93–97. [CrossRef]
359.
Najt, C.P.; Khan, S.A.; Heden, T.D.; Witthuhn, B.A.; Perez, M.; Heier, J.L.; Mead, L.E.; Franklin, M.P.; Karanja, K.K.;
Graham, M.J.; et al. Lipid Droplet-Derived Monounsaturated Fatty Acids Traffic via PLIN5 to Allosterically Activate SIRT1. Mol.
Cell 2020,77, 810–824 e818. [CrossRef]
360.
Han, L.; Zhou, R.; Niu, J.; McNutt, M.A.; Wang, P.; Tong, T. SIRT1 is regulated by a PPARgamma-SIRT1 negative feedback loop
associated with senescence. Nucleic Acids Res. 2010,38, 7458–7471. [CrossRef]
361.
Moerman, E.J.; Teng, K.; Lipschitz, D.A.; Lecka-Czernik, B. Aging activates adipogenic and suppresses osteogenic programs in
mesenchymal marrow stroma/stem cells: The role of PPAR-gamma2 transcription factor and TGF-beta/BMP signaling pathways.
Aging Cell 2004,3, 379–389. [CrossRef]
362.
Gong, J.; Sun, Z.; Wu, L.; Xu, W.; Schieber, N.; Xu, D.; Shui, G.; Yang, H.; Parton, R.G.; Li, P. Fsp27 promotes lipid droplet growth
by lipid exchange and transfer at lipid droplet contact sites. J. Cell Biol. 2011,195, 953–963. [CrossRef]
363.
Wang, F.; Ren, S.Y.; Chen, J.F.; Liu, K.; Li, R.X.; Li, Z.F.; Hu, B.; Niu, J.Q.; Xiao, L.; Chan, J.R.; et al. Myelin degeneration and
diminished myelin renewal contribute to age-related deficits in memory. Nat. Neurosci. 2020,23, 481–486. [CrossRef]
364.
Farokhian, F.; Yang, C.; Beheshti, I.; Matsuda, H.; Wu, S. Age-Related Gray and White Matter Changes in Normal Adult Brains.
Aging Dis. 2017,8, 899–909. [CrossRef]
365.
Capilla-Gonzalez, V.; Cebrian-Silla, A.; Guerrero-Cazares, H.; Garcia-Verdugo, J.M.; Quiñones-Hinojosa, A. Age-related changes
in astrocytic and ependymal cells of the subventricular zone. Glia 2014,62, 790–803. [CrossRef] [PubMed]
366.
Kabaso, D.; Coskren, P.J.; Henry, B.I.; Hof, P.R.; Wearne, S.L. The electrotonic structure of pyramidal neurons contributing to
prefrontal cortical circuits in macaque monkeys is significantly altered in aging. Cereb. Cortex 2009,19, 2248–2268. [CrossRef]
367.
O’Brien, J.S.; Sampson, E.L. Lipid composition of the normal human brain: Gray matter, white matter, and myelin. J. Lipid Res.
1965,6, 537–544. [CrossRef]
368.
Yang, C.; Wang, X.; Wang, J.; Wang, X.; Chen, W.; Lu, N.; Siniossoglou, S.; Yao, Z.; Liu, K. Rewiring Neuronal Glycerolipid
Metabolism Determines the Extent of Axon Regeneration. Neuron 2020,105, 276–292.e275. [CrossRef]
369.
Kaushik, S.; Rodriguez-Navarro, J.A.; Arias, E.; Kiffin, R.; Sahu, S.; Schwartz, G.J.; Cuervo, A.M.; Singh, R. Autophagy in
hypothalamic AgRP neurons regulates food intake and energy balance. Cell Metab. 2011,14, 173–183. [CrossRef] [PubMed]
370.
Ioannou, M.S.; Jackson, J.; Sheu, S.H.; Chang, C.L.; Weigel, A.V.; Liu, H.; Pasolli, H.A.; Xu, C.S.; Pang, S.; Matthies, D.; et al.
Neuron-Astrocyte Metabolic Coupling Protects against Activity-Induced Fatty Acid Toxicity. Cell
2019
,177, 1522–1535.e1514.
[CrossRef]

Biomolecules 2023,13, 912 52 of 56
371.
Martinez-Vicente, M.; Talloczy, Z.; Wong, E.; Tang, G.; Koga, H.; Kaushik, S.; de Vries, R.; Arias, E.; Harris, S.; Sulzer, D.; et al.
Cargo recognition failure is responsible for inefficient autophagy in Huntington’s disease. Nat. Neurosci.
2010
,13, 567–576.
[CrossRef]
372.
Marschallinger, J.; Iram, T.; Zardeneta, M.; Lee, S.E.; Lehallier, B.; Haney, M.S.; Pluvinage, J.V.; Mathur, V.; Hahn, O.;
Morgens, D.W.; et al. Lipid-droplet-accumulating microglia represent a dysfunctional and proinflammatory state in the aging
brain. Nat. Neurosci. 2020,23, 194–208. [CrossRef]
373.
Shimabukuro, M.K.; Langhi, L.G.; Cordeiro, I.; Brito, J.M.; Batista, C.M.; Mattson, M.P.; Mello Coelho, V. Lipid-laden cells
differentially distributed in the aging brain are functionally active and correspond to distinct phenotypes. Sci. Rep.
2016
,6, 23795.
[CrossRef]
374.
Bresgen, N.; Jaksch, H.; Bauer, H.C.; Eckl, P.; Krizbai, I.; Tempfer, H. Astrocytes are more resistant than cerebral endothelial cells
toward geno- and cytotoxicity mediated by short-term oxidative stress. J. Neurosci. Res.
2006
,84, 1821–1828. [CrossRef] [PubMed]
375.
Schroeter, M.L.; Mertsch, K.; Giese, H.; Muller, S.; Sporbert, A.; Hickel, B.; Blasig, I.E. Astrocytes enhance radical defence in
capillary endothelial cells constituting the blood-brain barrier. FEBS Lett. 1999,449, 241–244. [CrossRef]
376.
Husain, M.A.; Laurent, B.; Plourde, M. APOE and Alzheimer’s Disease: From Lipid Transport to Physiopathology and Therapeu-
tics. Front. Neurosci. 2021,15, 630502. [CrossRef]
377.
Corder, E.H.; Saunders, A.M.; Strittmatter, W.J.; Schmechel, D.E.; Gaskell, P.C.; Small, G.W.; Roses, A.D.; Haines, J.L.; Pericak-
Vance, M.A. Gene dose of apolipoprotein E type 4 allele and the risk of Alzheimer’s disease in late onset families. Science
1993
,
261, 921–923. [CrossRef]
378.
Liu, L.; MacKenzie, K.R.; Putluri, N.; Maletic-Savatic, M.; Bellen, H.J. The Glia-Neuron Lactate Shuttle and Elevated ROS Promote
Lipid Synthesis in Neurons and Lipid Droplet Accumulation in Glia via APOE/D. Cell Metab.
2017
,26, 719–737.e716. [CrossRef]
379.
Xu, L.; Pu, J. Alpha-Synuclein in Parkinson’s Disease: From Pathogenetic Dysfunction to Potential Clinical Application. Park. Dis.
2016,2016, 1720621. [CrossRef]
380.
Polymeropoulos, M.H.; Lavedan, C.; Leroy, E.; Ide, S.E.; Dehejia, A.; Dutra, A.; Pike, B.; Root, H.; Rubenstein, J.; Boyer, R.; et al.
Mutation in the alpha-synuclein gene identified in families with Parkinson’s disease. Science 1997,276, 2045–2047. [CrossRef]
381.
Loov, C.; Scherzer, C.R.; Hyman, B.T.; Breakefield, X.O.; Ingelsson, M. alpha-Synuclein in Extracellular Vesicles: Functional
Implications and Diagnostic Opportunities. Cell. Mol. Neurobiol. 2016,36, 437–448. [CrossRef]
382. Gasser, T.; Hardy, J.; Mizuno, Y. Milestones in PD genetics. Mov. Disord. 2011,26, 1042–1048. [CrossRef]
383.
Islimye, E.; Girard, V.; Gould, A.P. Functions of Stress-Induced Lipid Droplets in the Nervous System. Front. Cell Dev. Biol.
2022
,
10, 863907. [CrossRef]
384.
Taran, A.S.; Shuvalova, L.D.; Lagarkova, M.A.; Alieva, I.B. Huntington’s Disease-An Outlook on the Interplay of the HTT Protein,
Microtubules and Actin Cytoskeletal Components. Cells 2020,9, 1514. [CrossRef]
385.
Churkina Taran, A.S.; Shakhov, A.S.; Kotlobay, A.A.; Alieva, I.B. Huntingtin and Other Neurodegeneration-Associated Proteins
in the Development of Intracellular Pathologies: Potential Target Search for Therapeutic Intervention. Int. J. Mol. Sci.
2022
,
23, 5533. [CrossRef]
386.
Gruber, A.; Hornburg, D.; Antonin, M.; Krahmer, N.; Collado, J.; Schaffer, M.; Zubaite, G.; Lüchtenborg, C.; Sachsenheimer,
T.; Brügger, B.; et al. Molecular and structural architecture of polyQ aggregates in yeast. Proc. Natl. Acad. Sci. USA
2018
,
115, E3446–E3453. [CrossRef]
387.
Cyske, Z.; Gaffke, L.; Pierzynowska, K.; W˛egrzyn, G. Tubulin Cytoskeleton in Neurodegenerative Diseases-not Only Primary
Tubulinopathies. Cell. Mol. Neurobiol. 2022.Online ahead of print. [CrossRef]
388.
Ketut, S.; Pande, D.; Made Siswadi, S.; Kuswardhani, R.A.T. Age is an Important Risk Factor for Type 2 Diabetes Mellitus and
Cardiovascular Diseases. In Glucose Tolerance; Sureka, C., Ed.; IntechOpen: Rijeka, Croatia, 2012; p. Ch. 5.
389.
Han, S.K.; Baik, S.K.; Kim, M.Y. Non-alcoholic fatty liver disease: Definition and subtypes. Clin. Mol. Hepatol.
2022
,29, S5–S16.
[CrossRef]
390.
Guo, X.; Yin, X.; Liu, Z.; Wang, J. Non-Alcoholic Fatty Liver Disease (NAFLD) Pathogenesis and Natural Products for Prevention
and Treatment. Int. J. Mol. Sci. 2022,23, 5489. [CrossRef]
391.
Buzzetti, E.; Pinzani, M.; Tsochatzis, E.A. The multiple-hit pathogenesis of non-alcoholic fatty liver disease (NAFLD). Metabolism
2016,65, 1038–1048. [CrossRef]
392.
Ogrodnik, M.; Miwa, S.; Tchkonia, T.; Tiniakos, D.; Wilson, C.L.; Lahat, A.; Day, C.P.; Burt, A.; Palmer, A.; Anstee, Q.M.; et al.
Cellular senescence drives age-dependent hepatic steatosis. Nat. Commun. 2017,8, 15691. [CrossRef]
393. Lau, L.F. CCN1/CYR61: The very model of a modern matricellular protein. Cell. Mol. Life Sci. 2011,68, 3149–3163. [CrossRef]
394.
Kim, K.H.; Won, J.H.; Cheng, N.; Lau, L.F. The matricellular protein CCN1 in tissue injury repair. J. Cell Commun. Signal.
2018
,
12, 273–279. [CrossRef] [PubMed]
395.
Ju, L.; Sun, Y.; Xue, H.; Chen, L.; Gu, C.; Shao, J.; Lu, R.; Luo, X.; Wei, J.; Ma, X.; et al. CCN1 promotes hepatic steatosis and
inflammation in non-alcoholic steatohepatitis. Sci. Rep. 2020,10, 3201. [CrossRef]
396.
Kim, K.H.; Chen, C.C.; Monzon, R.I.; Lau, L.F. Matricellular protein CCN1 promotes regression of liver fibrosis through induction
of cellular senescence in hepatic myofibroblasts. Mol. Cell. Biol. 2013,33, 2078–2090. [CrossRef]
397.
Quan, T.; Qin, Z.; Voorhees, J.J.; Fisher, G.J. Cysteine-rich protein 61 (CCN1) mediates replicative senescence-associated aberrant
collagen homeostasis in human skin fibroblasts. J. Cell Biochem. 2012,113, 3011–3018. [CrossRef]

Biomolecules 2023,13, 912 53 of 56
398.
Qin, Z.; Robichaud, P.; He, T.; Fisher, G.J.; Voorhees, J.J.; Quan, T. Oxidant exposure induces cysteine-rich protein 61 (CCN1) via
c-Jun/AP-1 to reduce collagen expression in human dermal fibroblasts. PLoS ONE 2014,9, e115402. [CrossRef]
399.
Borkham-Kamphorst, E.; Steffen, B.T.; Van de Leur, E.; Haas, U.; Tihaa, L.; Friedman, S.L.; Weiskirchen, R. CCN1/CYR61
overexpression in hepatic stellate cells induces ER stress-related apoptosis. Cell. Signal. 2016,28, 34–42. [CrossRef]
400.
Cheng, N.Y.; Kim, K.H.; Lau, L.F. Senescent hepatic stellate cells promote liver regeneration through IL-6 and ligands of CXCR2.
JCI Insight 2022,7, e158207. [CrossRef]
401.
Han, D.; Ybanez, M.D.; Ahmadi, S.; Yeh, K.; Kaplowitz, N. Redox regulation of tumor necrosis factor signaling. Antioxid. Redox
Signal. 2009,11, 2245–2263. [CrossRef]
402.
Duvigneau, J.C.; Luis, A.; Gorman, A.M.; Samali, A.; Kaltenecker, D.; Moriggl, R.; Kozlov, A.V. Crosstalk between inflammatory
mediators and endoplasmic reticulum stress in liver diseases. Cytokine 2019,124, 154577. [CrossRef]
403.
Long, Z.; Cao, M.; Su, S.H.; Wu, G.Y.; Meng, F.S.; Wu, H.; Liu, J.Z.; Yu, W.H.; Atabai, K.; Wang, X. Inhibition of hepatocyte
nuclear factor 1b induces hepatic steatosis through DPP4/NOX1-mediated regulation of superoxide. Free Radic. Biol. Med.
2017
,
113, 71–83. [CrossRef]
404.
Bellanné-Chantelot, C.; Clauin, S.; Chauveau, D.; Collin, P.; Daumont, M.; Douillard, C.; Dubois-Laforgue, D.; Dusselier, L.;
Gautier, J.F.; Jadoul, M.; et al. Large genomic rearrangements in the hepatocyte nuclear factor-1beta (TCF2) gene are the most
frequent cause of maturity-onset diabetes of the young type 5. Diabetes 2005,54, 3126–3132. [CrossRef]
405.
Bonnycastle, L.L.; Willer, C.J.; Conneely, K.N.; Jackson, A.U.; Burrill, C.P.; Watanabe, R.M.; Chines, P.S.; Narisu, N.; Scott, L.J.;
Enloe, S.T.; et al. Common variants in maturity-onset diabetes of the young genes contribute to risk of type 2 diabetes in Finns.
Diabetes 2006,55, 2534–2540. [CrossRef]
406.
Barak, P.; Rai, A.; Rai, P.; Mallik, R. Quantitative optical trapping on single organelles in cell extract. Nat. Methods
2013
,10, 68–70.
[CrossRef]
407.
Rai, P.; Kumar, M.; Sharma, G.; Barak, P.; Das, S.; Kamat, S.S.; Mallik, R. Kinesin-dependent mechanism for controlling triglyceride
secretion from the liver. Proc. Natl. Acad. Sci. USA 2017,114, 12958–12963. [CrossRef]
408.
Thiam, A.R.; Antonny, B.; Wang, J.; Delacotte, J.; Wilfling, F.; Walther, T.C.; Beck, R.; Rothman, J.E.; Pincet, F. COPI buds 60-nm
lipid droplets from reconstituted water-phospholipid-triacylglyceride interfaces, suggesting a tension clamp function. Proc. Natl.
Acad. Sci. USA 2013,110, 13244–13249. [CrossRef]
409.
Kumar, M.; Ojha, S.; Rai, P.; Joshi, A.; Kamat, S.S.; Mallik, R. Insulin activates intracellular transport of lipid droplets to release
triglycerides from the liver. J. Cell Biol. 2019,218, 3697–3713. [CrossRef]
410.
Singh, R.; Kaushik, S.; Wang, Y.; Xiang, Y.; Novak, I.; Komatsu, M.; Tanaka, K.; Cuervo, A.M.; Czaja, M.J. Autophagy regulates
lipid metabolism. Nature 2009,458, 1131–1135. [CrossRef]
411.
Singh, R.; Cuervo, A.M. Lipophagy: Connecting autophagy and lipid metabolism. Int. J. Cell Biol.
2012
,2012, 282041. [CrossRef]
412.
Korovila, I.; Höhn, A.; Jung, T.; Grune, T.; Ott, C. Reduced Liver Autophagy in High-Fat Diet Induced Liver Steatosis in New
Zealand Obese Mice. Antioxidants 2021,10, 501. [CrossRef]
413.
Greenberg, A.S.; Coleman, R.A.; Kraemer, F.B.; McManaman, J.L.; Obin, M.S.; Puri, V.; Yan, Q.W.; Miyoshi, H.; Mashek, D.G. The
role of lipid droplets in metabolic disease in rodents and humans. J. Clin. Investig. 2011,121, 2102–2110. [CrossRef]
414. Unger, R.H.; Orci, L. Lipoapoptosis: Its mechanism and its diseases. Biochim. Biophys. Acta 2002,1585, 202–212. [CrossRef]
415.
Akazawa, Y.; Nakao, K. Lipotoxicity pathways intersect in hepatocytes: Endoplasmic reticulum stress, c-Jun N-terminal kinase-1,
and death receptors. Hepatol. Res. 2016,46, 977–984. [CrossRef]
416.
Gaggini, M.; Ndreu, R.; Michelucci, E.; Rocchiccioli, S.; Vassalle, C. Ceramides as Mediators of Oxidative Stress and Inflammation
in Cardiometabolic Disease. Int. J. Mol. Sci. 2022,23, 2719. [CrossRef]
417.
Poli, G.; Schaur, R.J.; Siems, W.G.; Leonarduzzi, G. 4-hydroxynonenal: A membrane lipid oxidation product of medicinal interest.
Med. Res. Rev. 2008,28, 569–631. [CrossRef]
418.
Gueraud, F.; Atalay, M.; Bresgen, N.; Cipak, A.; Eckl, P.M.; Huc, L.; Jouanin, I.; Siems, W.; Uchida, K. Chemistry and biochemistry
of lipid peroxidation products. Free Radic. Res. 2010,44, 1098–1124. [CrossRef]
419.
Dixon, S.J.; Lemberg, K.M.; Lamprecht, M.R.; Skouta, R.; Zaitsev, E.M.; Gleason, C.E.; Patel, D.N.; Bauer, A.J.; Cantley, A.M.;
Yang, W.S.; et al. Ferroptosis: An iron-dependent form of nonapoptotic cell death. Cell
2012
,149, 1060–1072. [CrossRef] [PubMed]
420.
Xiong, F.; Zhou, Q.; Huang, X.; Cao, P.; Wang, Y. Ferroptosis plays a novel role in nonalcoholic steatohepatitis pathogenesis. Front.
Pharmacol. 2022,13, 1055793. [CrossRef]
421.
Kazankov, K.; Jørgensen, S.M.D.; Thomsen, K.L.; Møller, H.J.; Vilstrup, H.; George, J.; Schuppan, D.; Grønbæk, H. The role
of macrophages in nonalcoholic fatty liver disease and nonalcoholic steatohepatitis. Nat. Rev. Gastroenterol. Hepatol.
2019
,
16, 145–159. [CrossRef]
422.
Tiniakos, D.G.; Vos, M.B.; Brunt, E.M. Nonalcoholic fatty liver disease: Pathology and pathogenesis. Annu. Rev. Pathol.
2010
,
5, 145–171. [CrossRef]
423.
Takahashi, Y.; Fukusato, T. Histopathology of nonalcoholic fatty liver disease/nonalcoholic steatohepatitis. World J. Gastroenterol.
2014,20, 15539–15548. [CrossRef]
424.
Matsumoto, A.; Naito, M.; Itakura, H.; Ikemoto, S.; Asaoka, H.; Hayakawa, I.; Kanamori, H.; Aburatani, H.; Takaku, F.;
Suzuki, H.; et al. Human macrophage scavenger receptors: Primary structure, expression, and localization in atherosclerotic
lesions. Proc. Natl. Acad. Sci. USA 1990,87, 9133–9137. [CrossRef]

Biomolecules 2023,13, 912 54 of 56
425.
Moore, K.J.; Kunjathoor, V.V.; Koehn, S.L.; Manning, J.J.; Tseng, A.A.; Silver, J.M.; McKee, M.; Freeman, M.W. Loss of receptor-
mediated lipid uptake via scavenger receptor A or CD36 pathways does not ameliorate atherosclerosis in hyperlipidemic mice.
J. Clin. Investig. 2005,115, 2192–2201. [CrossRef]
426.
Gerrity, R.G. The role of the monocyte in atherogenesis: I. Transition of blood-borne monocytes into foam cells in fatty lesions.
Am. J. Pathol. 1981,103, 181–190. [PubMed]
427. Tabas, I. Nonoxidative modifications of lipoproteins in atherogenesis. Annu. Rev. Nutr. 1999,19, 123–139. [CrossRef]
428.
Pohl, A.; Devaux, P.F.; Herrmann, A. Function of prokaryotic and eukaryotic ABC proteins in lipid transport. Biochim. Biophys.
Acta 2005,1733, 29–52. [CrossRef] [PubMed]
429.
Cuchel, M.; Rader, D.J. Macrophage reverse cholesterol transport: Key to the regression of atherosclerosis? Circulation
2006
,
113, 2548–2555. [CrossRef]
430.
Chang, T.Y.; Chang, C.C.; Lin, S.; Yu, C.; Li, B.L.; Miyazaki, A. Roles of acyl-coenzyme A:cholesterol acyltransferase-1 and -2. Curr.
Opin. Lipidol. 2001,12, 289–296. [CrossRef]
431.
Yano, H.; Fujiwara, Y.; Horlad, H.; Pan, C.; Kai, K.; Niino, D.; Ohsawa, K.; Higashi, M.; Nosaka, K.; Okuno, Y.; et al. Blocking
cholesterol efflux mechanism is a potential target for antilymphoma therapy. Cancer Sci. 2022,113, 2129–2143. [CrossRef]
432.
Hynynen, R.; Suchanek, M.; Spandl, J.; Bäck, N.; Thiele, C.; Olkkonen, V.M. OSBP-related protein 2 is a sterol receptor on lipid
droplets that regulates the metabolism of neutral lipids. J. Lipid Res. 2009,50, 1305–1315. [CrossRef]
433.
Guyard, V.; Monteiro-Cardoso, V.F.; Omrane, M.; Sauvanet, C.; Houcine, A.; Boulogne, C.; Ben Mbarek, K.; Vitale, N.; Faklaris, O.;
El Khallouki, N.; et al. ORP5 and ORP8 orchestrate lipid droplet biogenesis and maintenance at ER-mitochondria contact sites.
J. Cell Biol. 2022,221, 2107. [CrossRef]
434.
Anderson, A.; Campo, A.; Fulton, E.; Corwin, A.; Jerome, W.G., 3rd; O’Connor, M.S. 7-Ketocholesterol in disease and aging. Redox
Biol. 2020,29, 101380. [CrossRef]
435.
Ghzaiel, I.; Nury, T.; Zarrouk, A.; Vejux, A.; Lizard, G. Oxiapoptophagy in Age-Related Diseases. Comment on Ouyang et al.
7-Ketocholesterol Induces Oxiapoptophagy and Inhibits Osteogenic Differentiation in MC3T3-E1 Cells. Cells
2022
,11, 2882.
[CrossRef]
436.
Monier, S.; Samadi, M.; Prunet, C.; Denance, M.; Laubriet, A.; Athias, A.; Berthier, A.; Steinmetz, E.; Jürgens, G.;
Nègre-Salvayre, A.; et al. Impairment of the cytotoxic and oxidative activities of 7 beta-hydroxycholesterol and 7-ketocholesterol
by esterification with oleate. Biochem. Biophys. Res. Commun. 2003,303, 814–824. [CrossRef]
437.
Nury, T.; Zarrouk, A.; Yammine, A.; Mackrill, J.J.; Vejux, A.; Lizard, G. Oxiapoptophagy: A type of cell death induced by some
oxysterols. Br. J. Pharmacol. 2021,178, 3115–3123. [CrossRef]
438.
Ouyang, J.; Xiao, Y.; Ren, Q.; Huang, J.; Zhou, Q.; Zhang, S.; Li, L.; Shi, W.; Chen, Z.; Wu, L. 7-Ketocholesterol Induces
Oxiapoptophagy and Inhibits Osteogenic Differentiation in MC3T3-E1 Cells. Cells 2022,11, 2882. [CrossRef]
439.
Samadi, A.; Sabuncuoglu, S.; Samadi, M.; Isikhan, S.Y.; Chirumbolo, S.; Peana, M.; Lay, I.; Yalcinkaya, A.; Bjørklund, G. A
Comprehensive Review on Oxysterols and Related Diseases. Curr. Med. Chem. 2021,28, 110–136. [CrossRef]
440.
Lee-Rueckert, M.; Lappalainen, J.; Kovanen, P.T.; Escola-Gil, J.C. Lipid-Laden Macrophages and Inflammation in Atherosclerosis
and Cancer: An Integrative View. Front. Cardiovasc. Med. 2022,9, 777822. [CrossRef]
441. Odegaard, J.I.; Chawla, A. Alternative macrophage activation and metabolism. Annu. Rev. Pathol. 2011,6, 275–297. [CrossRef]
442.
Lathe, R.; Sapronova, A.; Kotelevtsev, Y. Atherosclerosis and Alzheimer–diseases with a common cause? Inflammation, oxysterols,
vasculature. BMC Geriatr. 2014,14, 36. [CrossRef]
443.
Razani, B.; Feng, C.; Coleman, T.; Emanuel, R.; Wen, H.; Hwang, S.; Ting, J.P.; Virgin, H.W.; Kastan, M.B.; Semenkovich, C.F.
Autophagy links inflammasomes to atherosclerotic progression. Cell Metab. 2012,15, 534–544. [CrossRef]
444.
Robichaud, S.; Rasheed, A.; Pietrangelo, A.; Kim, A.D.; Boucher, D.M.; Emerton, C.; Vijithakumar, V.; Gharibeh, L.; Fairman, G.;
Mak, E.; et al. Autophagy Is Differentially Regulated in Leukocyte and Nonleukocyte Foam Cells during Atherosclerosis. Circ.
Res. 2022,130, 831–847. [CrossRef]
445.
Feil, S.; Fehrenbacher, B.; Lukowski, R.; Essmann, F.; Schulze-Osthoff, K.; Schaller, M.; Feil, R. Transdifferentiation of vascular
smooth muscle cells to macrophage-like cells during atherogenesis. Circ. Res. 2014,115, 662–667. [CrossRef]
446.
Banke, N.H.; Wende, A.R.; Leone, T.C.; O’Donnell, J.M.; Abel, E.D.; Kelly, D.P.; Lewandowski, E.D. Preferential oxidation of
triacylglyceride-derived fatty acids in heart is augmented by the nuclear receptor PPARalpha. Circ. Res.
2010
,107, 233–241.
[CrossRef]
447.
Barger, P.M.; Brandt, J.M.; Leone, T.C.; Weinheimer, C.J.; Kelly, D.P. Deactivation of peroxisome proliferator-activated receptor-
alpha during cardiac hypertrophic growth. J. Clin. Investig. 2000,105, 1723–1730. [CrossRef]
448.
Ding, G.; Fu, M.; Qin, Q.; Lewis, W.; Kim, H.W.; Fukai, T.; Bacanamwo, M.; Chen, Y.E.; Schneider, M.D.; Mangelsdorf, D.J.; et al.
Cardiac peroxisome proliferator-activated receptor gamma is essential in protecting cardiomyocytes from oxidative damage.
Cardiovasc. Res. 2007,76, 269–279. [CrossRef]
449.
Amen, T.; Kaganovich, D. Small Molecule Screen Reveals Joint Regulation of Stress Granule Formation and Lipid Droplet
Biogenesis. Front. Cell Dev. Biol. 2020,8, 606111. [CrossRef]
450. Sharma, S.; Adrogue, J.V.; Golfman, L.; Uray, I.; Lemm, J.; Youker, K.; Noon, G.P.; Frazier, O.H.; Taegtmeyer, H. Intramyocardial
lipid accumulation in the failing human heart resembles the lipotoxic rat heart. FASEB J. 2004,18, 1692–1700. [CrossRef]
451.
Cui, X.; Wang, J.; Zhang, Y.; Wei, J.; Wang, Y. Plin5, a New Target in Diabetic Cardiomyopathy. Oxidative Med. Cell. Longev.
2022
,
2022, 2122856. [CrossRef]

Biomolecules 2023,13, 912 55 of 56
452.
Zhang, X.; Xu, W.; Xu, R.; Wang, Z.; Zhang, X.; Wang, P.; Peng, K.; Li, M.; Li, J.; Tan, Y.; et al. Plin5 Bidirectionally Regulates Lipid
Metabolism in Oxidative Tissues. Oxidative Med. Cell. Longev. 2022,2022, 4594956. [CrossRef]
453.
Wang, H.; Sreenivasan, U.; Hu, H.; Saladino, A.; Polster, B.M.; Lund, L.M.; Gong, D.W.; Stanley, W.C.; Sztalryd, C. Perilipin 5,
a lipid droplet-associated protein, provides physical and metabolic linkage to mitochondria. J. Lipid Res.
2011
,52, 2159–2168.
[CrossRef]
454.
Holzem, K.M.; Vinnakota, K.C.; Ravikumar, V.K.; Madden, E.J.; Ewald, G.A.; Dikranian, K.; Beard, D.A.; Efimov, I.R. Mitochondrial
structure and function are not different between nonfailing donor and end-stage failing human hearts. Faseb J.
2016
,30, 2698–2707.
[CrossRef]
455.
Pollak, N.M.; Schweiger, M.; Jaeger, D.; Kolb, D.; Kumari, M.; Schreiber, R.; Kolleritsch, S.; Markolin, P.; Grabner, G.F.;
Heier, C.; et al. Cardiac-specific overexpression of perilipin 5 provokes severe cardiac steatosis via the formation of a lipolytic
barrier. J. Lipid Res. 2013,54, 1092–1102. [CrossRef]
456.
Wang, H.; Sreenivasan, U.; Gong, D.W.; O’Connell, K.A.; Dabkowski, E.R.; Hecker, P.A.; Ionica, N.; Konig, M.; Mahurkar, A.;
Sun, Y.; et al. Cardiomyocyte-specific perilipin 5 overexpression leads to myocardial steatosis and modest cardiac dysfunction.
J. Lipid Res. 2013,54, 953–965. [CrossRef]
457.
Kuramoto, K.; Okamura, T.; Yamaguchi, T.; Nakamura, T.Y.; Wakabayashi, S.; Morinaga, H.; Nomura, M.; Yanase, T.; Otsu, K.;
Usuda, N.; et al. Perilipin 5, a lipid droplet-binding protein, protects heart from oxidative burden by sequestering fatty acid from
excessive oxidation. J. Biol. Chem. 2012,287, 23852–23863. [CrossRef]
458.
Zhu, Y.; Zhang, X.; Zhang, L.; Zhang, M.; Li, L.; Luo, D.; Zhong, Y. Perilipin5 protects against lipotoxicity and alleviates
endoplasmic reticulum stress in pancreatic beta-cells. Nutr. Metab. 2019,16, 50. [CrossRef]
459.
Cinato, M.; Mardani, I.; Miljanovic, A.; Drevinge, C.; Laudette, M.; Bollano, E.; Henricsson, M.; Tolo, J.; Bauza Thorbrugge, M.;
Levin, M.; et al. Cardiac Plin5 interacts with SERCA2 and promotes calcium handling and cardiomyocyte contractility. Life Sci.
Alliance 2023,6, e202201690. [CrossRef]
460.
Cruz, A.L.S.; Barreto, E.A.; Fazolini, N.P.B.; Viola, J.P.B.; Bozza, P.T. Lipid droplets: Platforms with multiple functions in cancer
hallmarks. Cell Death Dis. 2020,11, 105. [CrossRef]
461.
Ukraintseva, S.V.; Yashin, A.I. Opposite Phenotypes of Cancer and Aging Arise from Alternative Regulation of Common Signaling
Pathways. Ann. N. Y. Acad. Sci. 2003,1010, 489–492. [CrossRef]
462.
Antunes, P.; Cruz, A.; Barbosa, J.; Bonifácio, V.D.B.; Pinto, S.N. Lipid Droplets in Cancer: From Composition and Role to Imaging
and Therapeutics. Molecules 2022,27, 991. [CrossRef]
463. Li, Z.; Liu, H.; Luo, X. Lipid droplet and its implication in cancer progression. Am. J. Cancer Res. 2020,10, 4112–4122.
464.
Lung, J.; Hung, M.-S.; Wang, T.-Y.; Chen, K.-L.; Luo, C.-W.; Jiang, Y.-Y.; Wu, S.-Y.; Lee, L.-W.; Lin, P.-Y.; Chen, F.-F.; et al. Lipid
Droplets in Lung Cancers Are Crucial for the Cell Growth and Starvation Survival. Int. J. Mol. Sci. 2022,23, 12533. [CrossRef]
465.
Shyu, P.; Wong, X.F.A.; Crasta, K.; Thibault, G. Dropping in on lipid droplets: Insights into cellular stress and cancer. Biosci. Rep.
2018,38, BSR20180764. [CrossRef]
466.
Castelli, S.; De Falco, P.; Ciccarone, F.; Desideri, E.; Ciriolo, M.R. Lipid Catabolism and ROS in Cancer: A Bidirectional Liaison.
Cancers 2021,13, 5484. [CrossRef]
467.
Zhang, J.; Liu, Z.; Lian, Z.; Liao, R.; Chen, Y.; Qin, Y.; Wang, J.; Jiang, Q.; Wang, X.; Gong, J. Monoacylglycerol Lipase: A Novel
Potential Therapeutic Target and Prognostic Indicator for Hepatocellular Carcinoma. Sci. Rep. 2016,6, 35784. [CrossRef]
468.
Patel, D.; Salloum, D.; Saqcena, M.; Chatterjee, A.; Mroz, V.; Ohh, M.; Foster, D.A. A Late G1 Lipid Checkpoint That Is
Dysregulated in Clear Cell Renal Carcinoma Cells. J. Biol. Chem. 2017,292, 936–944. [CrossRef]
469. Qi, W.; Weber, C.R.; Wasland, K.; Roy, H.; Wali, R.; Joshi, S.; Savkovic, S.D. Tumor suppressor FOXO3 mediates signals from the
EGF receptor to regulate proliferation of colonic cells. Am. J. Physiol. -Gastrointest. Liver Physiol.
2011
,300, G264–G272. [CrossRef]
[PubMed]
470. Yu, W.; Bozza, P.T.; Tzizik, D.M.; Gray, J.P.; Cassara, J.; Dvorak, A.M.; Weller, P.F. Co-compartmentalization of MAP kinases and
cytosolic phospholipase A2 at cytoplasmic arachidonate-rich lipid bodies. Am. J. Pathol. 1998,152, 759–769. [PubMed]
471.
Yu, W.; Cassara, J.; Weller, P.F. Phosphatidylinositide 3-kinase localizes to cytoplasmic lipid bodies in human polymorphonuclear
leukocytes and other myeloid-derived cells. Blood 2000,95, 1078–1085. [CrossRef]
472.
Mittal, D.; Gubin, M.M.; Schreiber, R.D.; Smyth, M.J. New insights into cancer immunoediting and its three component phases—
Elimination, equilibrium and escape. Curr. Opin. Immunol. 2014,27, 16–25. [CrossRef]
473. Wong, R.S.Y. Apoptosis in cancer: From pathogenesis to treatment. J. Exp. Clin. Cancer Res. 2011,30, 87. [CrossRef]
474.
Bozza, P.T.; Bakker-Abreu, I.; Navarro-Xavier, R.A.; Bandeira-Melo, C. Lipid body function in eicosanoid synthesis: An update.
Prostaglandins Leukot. Essent. Fat. Acids (PLEFA) 2011,85, 205–213. [CrossRef]
475.
Accioly, M.T.; Pacheco, P.; Maya-Monteiro, C.M.; Carrossini, N.; Robbs, B.K.; Oliveira, S.S.; Kaufmann, C.; Morgado-Diaz, J.A.;
Bozza, P.T.; Viola, J.P. Lipid bodies are reservoirs of cyclooxygenase-2 and sites of prostaglandin-E2 synthesis in colon cancer cells.
Cancer Res. 2008,68, 1732–1740. [CrossRef]
476.
Johnson, A.M.; Kleczko, E.K.; Nemenoff, R.A. Eicosanoids in Cancer: New Roles in Immunoregulation. Front. Pharmacol.
2020
,
11, 595498. [CrossRef]
477.
Finetti, F.; Travelli, C.; Ercoli, J.; Colombo, G.; Buoso, E.; Trabalzini, L. Prostaglandin E2 and Cancer: Insight into Tumor
Progression and Immunity. Biology 2020,9, 434. [CrossRef]

Biomolecules 2023,13, 912 56 of 56
478.
Veglia, F.; Tyurin, V.A.; Mohammadyani, D.; Blasi, M.; Duperret, E.K.; Donthireddy, L.; Hashimoto, A.; Kapralov, A.; Amoscato,
A.; Angelini, R.; et al. Lipid bodies containing oxidatively truncated lipids block antigen cross-presentation by dendritic cells in
cancer. Nat. Commun. 2017,8, 2122. [CrossRef]
479.
Hakumaki, J.M.; Kauppinen, R.A. 1H NMR visible lipids in the life and death of cells. Trends Biochem. Sci.
2000
,25, 357–362.
[CrossRef] [PubMed]
480. Blankenberg, F.G. In vivo detection of apoptosis. J. Nucl. Med. 2008,49 (Suppl. S2), 81S–95S. [CrossRef] [PubMed]
481. Henique, C.; Mansouri, A.; Fumey, G.; Lenoir, V.; Girard, J.; Bouillaud, F.; Prip-Buus, C.; Cohen, I. Increased mitochondrial fatty
acid oxidation is sufficient to protect skeletal muscle cells from palmitate-induced apoptosis. J. Biol. Chem.
2010
,285, 36818–36827.
[CrossRef]
482.
Choi, S.E.; Jung, I.R.; Lee, Y.J.; Lee, S.J.; Lee, J.H.; Kim, Y.; Jun, H.S.; Lee, K.W.; Park, C.B.; Kang, Y. Stimulation of lipogenesis as
well as fatty acid oxidation protects against palmitate-induced INS-1 beta-cell death. Endocrinology
2011
,152, 816–827. [CrossRef]
[PubMed]
483.
Boren, J.; Brindle, K.M. Apoptosis-induced mitochondrial dysfunction causes cytoplasmic lipid droplet formation. Cell Death
Differ. 2012,19, 1561–1570. [CrossRef]
484.
Wang, J.-B.; Qi, L.-L.; Zheng, S.-D.; Wu, T.-X. Curcumin induces apoptosis through the mitochondria-mediated apoptotic pathway
in HT-29 cells. J. Zhejiang Univ. Sci. B 2009,10, 93–102. [CrossRef]
485.
Roy, M.; Chakraborty, S.; Siddiqi, M.; Bhattacharya, R.K. Induction of Apoptosis in Tumor Cells by Natural Phenolic Compounds.
Asian Pac. J. Cancer Prev. APJCP 2002,3, 61–67.
486.
Jiang, M.C.; Yang-Yen, H.F.; Yen, J.J.; Lin, J.K. Curcumin induces apoptosis in immortalized NIH 3T3 and malignant cancer cell
lines. Nutr. Cancer 1996,26, 111–120. [CrossRef]
487.
Cao, J.; Liu, Y.; Jia, L.; Zhou, H.M.; Kong, Y.; Yang, G.; Jiang, L.P.; Li, Q.J.; Zhong, L.F. Curcumin induces apoptosis through
mitochondrial hyperpolarization and mtDNA damage in human hepatoma G2 cells. Free Radic. Biol. Med.
2007
,43, 968–975.
[CrossRef]
488.
Zhang, I.; Cui, Y.; Amiri, A.; Ding, Y.; Campbell, R.E.; Maysinger, D. Pharmacological inhibition of lipid droplet formation
enhances the effectiveness of curcumin in glioblastoma. Eur. J. Pharm. Biopharm. 2016,100, 66–76. [CrossRef]
489.
Jin, C.; Yuan, P. Implications of lipid droplets in lung cancer: Associations with drug resistance. Oncol. Lett.
2020
,20, 2091–2104.
[CrossRef] [PubMed]
490. Lossi, L. The concept of intrinsic versus extrinsic apoptosis. Biochem. J. 2022,479, 357–384. [CrossRef]
491.
Song, J.H.; Tse, M.C.L.; Bellail, A.; Phuphanich, S.; Khuri, F.; Kneteman, N.M.; Hao, C. Lipid Rafts and Nonrafts Mediate Tumor
Necrosis Factor–Related Apoptosis-Inducing Ligand–Induced Apoptotic and Nonapoptotic Signals in Non–Small Cell Lung
Carcinoma Cells. Cancer Res. 2007,67, 6946–6955. [CrossRef]
492.
Zembroski, A.S.; Andolino, C.; Buhman, K.K.; Teegarden, D. Proteomic Characterization of Cytoplasmic Lipid Droplets in Human
Metastatic Breast Cancer Cells. Front. Oncol. 2021,11, 576326. [CrossRef] [PubMed]
493.
Sofi, F.; Abbate, R.; Gensini, G.F.; Casini, A. Accruing evidence on benefits of adherence to the Mediterranean diet on health: An
updated systematic review and meta-analysis. Am. J. Clin. Nutr. 2010,92, 1189–1196. [CrossRef] [PubMed]
494.
Russo, M.A.; Sansone, L.; Polletta, L.; Runci, A.; Rashid, M.M.; De Santis, E.; Vernucci, E.; Carnevale, I.; Tafani, M. Sirtuins and
resveratrol-derived compounds: A model for understanding the beneficial effects of the Mediterranean diet. Endocr. Metab.
Immune Disord. Drug Targets 2014,14, 300–308. [CrossRef]
495.
Park, S.J.; Ahmad, F.; Philp, A.; Baar, K.; Williams, T.; Luo, H.; Ke, H.; Rehmann, H.; Taussig, R.; Brown, A.L.; et al. Resveratrol
ameliorates aging-related metabolic phenotypes by inhibiting cAMP phosphodiesterases. Cell 2012,148, 421–433. [CrossRef]
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