Muscle Lipid Droplets: Cellular Signaling to Exercise Physiology and Beyond

Abstract

Conventionally viewed as energy storage depots, lipid droplets (LDs) play a central role in muscle lipid metabolism and intracellular signaling, as recognized by recent advances in our biological understanding. Specific subpopulations of muscle LDs, defined by location and associated proteins, are responsible for distinct biological functions. In this review, the traditional view of muscle LDs is examined, and the emerging role of LDs in intracellular signaling is highlighted. The effects of chronic and acute exercise on muscle LD metabolism and signaling is discussed. In conclusion, future directions for muscle LD research are identified. The primary focus will be on human studies, with inclusion of select animal/cellular/non-muscle studies as appropriate, to provide the underlying mechanisms driving the observed findings.

Background: The Muscle Lipid Droplet Environment

Intramyocellular lipid (IMCL) refers to the generalized accumulation of muscle lipid droplets (LDs). Early studies characterizing IMCL, employed lipid extraction methods from muscle biopsies [1], lipid staining from muscle biopsies [2,3], or magnetic resonance spectroscopy [4]. These studies demonstrate an inverse relationship between IMCL and insulin sensitivity in sedentary humans [1,4]. Interestingly, endurance athletes have high IMCL [2,5], which is similar if not higher than patients with type 2 diabetes mellitus (T2DM), while maintaining high insulin sensitivity [2]. This phenomenon is described as the ‘athlete’s paradox’ [2]. Subsequent studies have used electron microscopy to study IMCL. These studies permitted a more nuanced assessment of the LDs that make up IMCL, including size, density, and localization, which are heavily influenced by metabolic state and exercise training [6–9]. In general, healthy individuals exhibit a median diameter of muscle LDs of approximately 500 nm (range 200–1400 nm) [8], with increased LD density, rather than size, driving the higher IMCL in Type 1 fibers rather than Type 2 fibers [8]. Additionally, Type 1 fibers have greater volume of LDs, mitochondria, and LD-mitochondria contacts than Type 2 fibers [9]. Returning to the ‘athlete’s paradox’, when muscle biopsies with matched IMCL are compared between trained individuals and patients with T2DM, the trained individuals have higher numbers of smaller LD in Type 1 fibers, whereas the patients with T2DM have larger LDs in Type 2 fibers [10].

Within a specific muscle fiber, LDs are distributed between myofibrils or ‘intramyofibrillar’ (IMF), or just below the surface membrane or ‘subsarcolemmal’ (SS) and are intimately associated with mitochondria [9,11,12]. The IMF LDs are the presumed fuel source for the IMF mitochondria, which provides energy for muscle contraction [13]. By contrast, the SS LDs presumably supply SS mitochondria, which provide energy for membrane related processes [13]. Previously, Ferreira et al., used centrifugation techniques to separate SS and IMF mitochondria, and conducted mass spectrometry to identify proteomic differences and respiratory chain activity between the two mitochondrial fractions [14]. Compared with SS mitochondria, IMF mitochondria had increased levels of oxidative phosphorylation proteins, and increased activity of the respiratory chain complex [14]. These findings suggest that IMF mitochondria are highly specialized towards energy production for contractile function, and are consistent with the observation that IMF LDs are preferentially used during energy demand [6,15]. Generally, the overall lipid content is higher in the SS region (2–20-fold higher) than the IMF region [8,9]. Even if training did not significantly alter overall SS or IMF volume within the muscle, training generally increases LD density, and significantly reduces LD size in the SS region [8]. In summary, technological advancements have permitted the field to move beyond IMCL, and highlight LDs as critical players in muscle lipid metabolism.

Traditional View of Muscle LD: Storage Depot

Each LD consists of a hydrophobic core, containing primarily sterol esters and triacylglycerol, surrounded by a phospholipid monolayer [16]. LDs are ostensibly derived from the endoplasmic reticulum, and numerous theories exist on the mechanisms driving LD growth [16]. Importantly, LDs are covered with hundreds of proteins that influence their structural and signaling properties. Of the LD proteins, the perilipins (PLIN) have been the most extensively characterized [17]. PLIN2 and PLIN5 have been the LDs most characterized in muscle [17–19]. PLIN2 is involved in LD growth [19] and blunting lipolysis [20]. Specifically, PLIN2 overexpression increases LD accumulation, whereas knockdown reduces LD formation and increases fat oxidation in cultured myotubes [19]. In contrast to PLIN2, PLIN5 appears to facilitate the interaction between LDs and mitochondria [18]. PLIN5 is highly expressed in oxidative tissue, such as heart, brown adipose tissue, liver, and muscle [21]. Although the role of PLIN5 may appear paradoxical, such as increasing fatty acid (FA) oxidation [22] yet decreasing lipolysis [21], the role of PLIN5 depends on its location (cytosol, nuclear, LD) and environment (lipid-loaded, fasting, β-adrenergic stimulated) [18]. During fasting or exercise, PLIN5 becomes phosphorylated, enhancing FA oxidation and mitochondrial biogenesis, while reducing basal lipolysis and coordinating mitochondrial LD interactions during the rested or fed state [18]. The significance of PLIN5 phosphorylation has been highlighted by recent work, where PLIN5 phospho-mimetic (S155E) and phosphor-dead mutants (S155A) were generated, and the necessity of this phosphorylation modification for PLIN5 nuclear translocation, as well as PGC-1α mediated mitochondrial biogenesis in response to cAMP/fasting conditions was demonstrated [23]. Recently, Gemmink et al., used super-resolution microscopy to observe PLIN5 at the interface between LDs and mitochondria [24], supporting a role for PLIN5-mediated contact between the two organelles.

Although relatively less studied than PLIN2 or PLIN5 within the muscle, PLIN3 and PLIN4 are also involved in muscle lipid metabolism. PLIN3 appears to be involved in muscle lipid oxidation as acute exercise increases PLIN3 protein content, which was positively associated with the increase in ex vivo muscle palmitate oxidation (obtained from muscle biopsies pre- and post-exercise), and whole body fat oxidation [25]. In vitro stimulation of myotubes by epinephrine, or a lipolytic cocktail, increased PLIN3 protein levels, whereas PLIN3 knockdown reduced lipid oxidation [25]. It has been previously shown that lipid infusion in trained participants increased the number of PLIN-positive LDs in Type I fibers, whereas the sedentary participants had an increased number of PLIN3-negative LD [26]. PLIN4 mRNA is associated with SS LD size and chronic exercise training decreased both SS lipid content and PLIN4 mRNA abundance, suggesting that PLIN4 expression is related to SS LD size [27]. In cardiac muscle, PLIN5 expression is linked to PLIN4 [28]. Knockdown or deletion of PLIN4 reduces PLIN5 mRNA and protein levels, resulting in a loss of cardiac lipid accumulation during fasting or high fat feeding [28]. Whether similar findings also exist in skeletal muscle remains unknown.

Muscle LDs are traditionally perceived as energy storage depots, releasing nonesterified fatty acids (NEFAs) during energetic demand via lipolysis. Skeletal muscle relies on either glucose or NEFAs for fuel, depending on substrate availability [29]. Acute NEFA exposure reduces muscle glucose uptake [29,30] as well as oxidative [31] and nonoxidative glucose disposal [30]. Although there are several fates for NEFAs entering muscle, LDs play a central role, as NEFAs entering resting muscle are first partitioned to LDs following uptake before they reach the mitochondria for oxidation [32]. Since 55–85% of muscle LDs are in contact with mitochondria [9], muscle LDs were traditionally thought to function as an important mitochondrial fuel source, especially during nutrient deprivation [33]. However, recent work in brown adipose tissue distinguishing between cytoplasmic and peridroplet mitochondrial subpopulations, elucidated LD-mitochondrial interactions are actually sites for LD expansion [34], as discussed later.

Lipolysis is driven by three major enzymes [35,36]: (i) adipose triglyceride lipase (ATGL); (ii) hormone sensitive lipase (HSL); and (iii) monoacylglycerol (MAG) lipase. These enzymes act sequentially by producing diacylglycerol (DAG), MAG, and glycerol, along with a NEFA at each step [36]. Multiple factors stimulate lipolysis, including acute exercise [37], muscle contraction [37–39], and β-adrenergic stimulation [39,40], and it is generally accepted to occur more at smaller LDs, due to a greater relative surface area for lipase action [16]. The acute exercise-induced increase in lipolysis consequently reduces IMCL, and this is recapitulated with in vitro muscle contraction [37–39] with initial effects reducing LD size, and prolonged contraction decreasing LD size and number, as revealed by in vitro contraction of rat soleus [37]. Approximately 98% of muscle triglyceride lipase activity has been attributed to the combined effect of ATGL and HSL, with ATGL continuing to play a significant role when HSL is pharmacologically inhibited, or genetically knocked out in mouse skeletal muscle [37]. Yet, muscle-specific ATGL knock-out (KO) mice [41] do not have significantly altered peak or submaximal exercise performance [42], possibly due to increased HSL compensation [42].

Emerging View of the LD Signaling Node: Intracellular Signaling between the LD and the Nucleus

Recent studies have identified additional roles of LDs beyond fuel storage and fuel provision. Given its crucial role in cellular bioenergetics, it is unsurprising that environmental stimuli and energy status heavily influence LD interactions with other organelles, such as the mitochondria, nucleus, lipid metabolites, and associated proteins [12,34]. The responsiveness of LDs to acute homeostatic challenges, such as nutrient starvation, lipid loading, cold exposure, and exercise, highlights the dynamic role of LDs, and their potential to serve as signaling nodes. Much work remains in clarifying the mechanisms by which different LD subpopulations are regulated, and how they influence specific cellular pathways. Nevertheless, LDs are increasingly recognized in playing key roles in intracellular signaling. In this section, the interplay between LDs and the nucleus is discussed. Much of what is known about LD signaling to the nucleus pertains to transcriptional regulation of mitochondrial biogenesis and FA oxidation. Due to the novelty of this area, data is presented from several tissue models (heart, muscle, and liver) to examine the potential relationship between LD signaling and the nucleus (Figure 1, Key Figure).

Figure 1. Key Figure. LD Signaling to the Nucleus.

Stimulation of the β-adrenergic receptor subsequently triggers AC-mediated production of cAMP and concomitant activation of PKA. Downstream phosphorylation of PLIN5 results in enhanced binding of lipolytically-derived monounsaturated fatty acids and nuclear translocation. In the nucleus, PLIN5 interacts with and promotes SIRT1-mediated deacetylation of select target genes including PGC-1α. These nuclear events ultimately upregulate PPARα/PGC-1α target gene expression, and eventually increase mitochondrial biogenesis and FA oxidation. Whether the IMF, SS, or both LD pools are involved in nuclear signaling remains to be determined. The black solid arrows indicate direct actions carried out by a protein/enzyme, the red broken arrow indicates upregulated downstream pathways, and the blue broken arrow indicates nuclear translocation. Abbreviations: AC, adenylate cyclase; ATGL, adipose triglyceride lipase; ATP, adenosine triphosphate; FA, fatty acid; IMF, intramyofibrillar; LD, lipid droplet; PGC-1α, peroxisome proliferator-activated receptor-γ coactivator 1α; PKA, protein kinase A; PLIN5, perilipin 5; PPARα, peroxisome proliferator-activated receptor α; SIRT1, sirtuin 1; SS, subsarcolemmal.

Although ATGL is necessary in mobilizing adipose-derived NEFAs during fasting for the muscle [43], ATGL also plays a critical role in regulating peroxisome proliferator-activated receptor α (PPARα), and peroxisome proliferator activated receptor-gamma coactivator-1α (PGC-1α) signaling, to promote intracellular fat oxidation and mitochondrial biogenesis [23,44,45].The lipolytic activity of ATGL at the periphery of LDs influences PPARα activation, which was traditionally attributed to the generation of ligands for PPARα, as administration of PPARα agonists reduced lipid accumulation in the heart and improved cardiac function in ATGL-deficient mice [44]. However, PPARα agonism with a pharmacologic PPAR-α agonist in ATGL-null mice could not completely restore FA utilization as assessed by respiratory quotients [44]. Moreover, fenofibrate-induced stimulation of PPARα, did not normalize PPARα target gene expression in liver with ATGL knockdown [46].

The NAD-dependent deacetylase sirtuin 1 (SIRT1) was also identified as a key player through its deacetylation of PGC-1α, a coactivator for PPARα signaling, in response to low glucose conditions, ultimately driving mitochondrial biogenesis and FA oxidation in C2C12 cells [47]. Activation of the cAMP and cAMP-dependent protein kinase (PKA) signaling cascade, via β-adrenergic stimulation or oleic acid exposure, also triggers SIRT1-mediated deacetylation of PGC-1α in a NAD-independent manner in mouse skeletal muscle [47]. Studies in murine hepatic cells later revealed the effects of β-adrenergic signaling on SIRT1 activation are actually mediated by ATGL [45].

In addition to its involvement with mitochondria-LD associations, PLIN5 is also implicated in LD-nuclear signaling. In both muscle and brown adipose tissue, fasting or catecholamines stimulate PKA-mediated phosphorylation of PLIN5, resulting in PLIN5 nuclear translocation where it interacts with SIRT1 and PGC-1α to ultimately drive a transcriptional program promoting FA catabolism [48]. Additionally, it was recently established that PLIN5 actually bridges ATGL activity with downstream SIRT1-PGC-1α-PPARα signaling, as lipolytically-derived monounsaturated FAs bind to and are trafficked via PLIN5, to allosterically activate SIRT1 in the nucleus (Figure 1) [23]. Collectively, the molecular mechanisms involved in FA oxidation and mitochondrial function involve key LD-associated proteins, underscoring the importance of the LD as a critical signaling node in lipid metabolism.

The Effect of Chronic Exercise on Muscle Fat Metabolism

In sedentary humans, there is an established inverse relationship between IMCL and insulin sensitivity [1,4], with ‘lipotoxicity’ impairing insulin signaling to cause insulin resistance [49]. Presumably, ‘lipotoxicity’ arises when increased levels of IMCL enhance production of lipotoxic metabolites, such as DAG [50], ceramides [51], long chain acyl-CoAs [52], and oxidative stress [53], which blunts muscle insulin signaling [49,53], and glucose uptake [49].

To explain the ‘athlete’s paradox’, subsequent investigations have identified several potential mechanisms including: (i) differences in IMCL localization within the muscle [7]; (ii) reduced, albeit inconsistently observed, accumulation of lipotoxic metabolites in athletes [54,55]; and (iii) increased muscle lipid turnover in athletes [55,56]. In terms of IMCL, the ‘athlete’s paradox’ can be partially explained by recent microscopy findings examining the distribution across Type 1/Type 2 fiber types and SS/IMF localization. Endurance athletes have more Type 1 fibers [5,57], which have higher IMCL [5,8] and less lipid content in the SS region [8]. Sedentary humans have more Type 2 fibers [57], with obesity [6] and T2DM [8] having greater lipid content in the SS region (Figure 2). As previously noted when IMCL was matched, patients with T2DM have larger LD size in the SS region of Type 2 fibers, whereas trained humans have higher LD density in the IMF region of Type 1 fibers [10]. This distribution is relevant, as Type 2 fibers are less oxidative than Type 1 fibers, and SS LDs presumably are the primary energy source for SS mitochondria, which are involved in membrane related processes, rather than muscle contraction [13].

Figure 2. Effects of Chronic Exercise Training on Muscle LD Distribution.

This is a schematic of LD distribution within the skeletal muscle of sedentary (A), or trained (B) humans. We demonstrate the distribution of LD and mitochondria within the SS space (designated by blue background) and IMF space (blue arrows). Sedentary behavior (A) is associated with higher levels of SS LDs and fewer mitochondria than chronic exercise training (B). In contrast to sedentary behavior, chronic aerobic training is associated with larger myofibrils, increased LD density, greater LD-mitochondria contact, higher IMF LDs, and fewer SS LDs. Abbreviations: LD, lipid droplet; IMF, intramyofibrillar; SS, subsarcolemmal.

The training duration and the participant’s activity status prior to training, likely influences the IMCL/LD adaptation to chronic aerobic training. Thus, changes in IMCL content (or lack of) needs to be considered within this context. Chronic aerobic exercise generally increases IMCL [5,12], which appears to be due to increasing LD density rather than LD size [5,12,58]. This may be mechanistically relevant as smaller LDs have a higher surface area to volume ratio than larger LDs, which enhances lipid mobilization and turnover [59]. Interestingly, aerobic training-related increases in IMCL are not necessarily observed [7,9], which may be due to differential effects of exercise on SS versus IMF LDs, as aerobic training can increase [10] or not alter IMF LDs [7], yet reduce SS LDs [7,9]. Chronic aerobic exercise may also increase LD contact with mitochondria [12], and alter localization of lipid metabolites [60] and LD-related proteins [58,61], to enhance lipid storage and future utilization. In contrast to chronic aerobic exercise, the effects of chronic resistance exercise on IMCL remains sparsely investigated, with the literature reporting mixed findings of increasing [62], not altering [63], or decreasing IMCL levels [64].

The Effect of Acute Exercise on Muscle Fat Metabolism

To appreciate the role of the LD as a signaling node, it is important to understand the response of LDs to acute perturbation, with acute exercise as a prototypical example. The effect of acute resistance exercise on muscle fat metabolism remains inconsistent, with few studies reported in the literature [65,66]. Table 1 focuses on the effects of acute aerobic exercise on muscle fat metabolism.

Table 1.

Acute Effects of Exercise on Muscle Lipid Metabolism^a

Acute exercise effects on IMCL
Exercise	Study design	Main outcome	Refs
Eight endurance trained male cyclists Cycling at 60%VO2max for 2 h with 2-h post-exercise recovery	Muscle biopsy pre-, immediately post-, and 2 h post-acute exercise Stable isotope tracers to measure fat and glucose oxidation	NEFA oxidation increased during exercise ~62% decline in IMCL only in Type 1 fibers due to reduced LD size and density No change in IMCL observed during 2-h recovery period	[70]
Nine Endurance trained male cyclists Cycling at 55% VO2max for 3 hours	MRS pre-, immediately post-, 24 h post-, and 48 h post-acute exercise Muscle biopsy at 48 h post-acute exercise Performed twice with 2 days post-exercise normal diet (39% energy from fat) and low-fat diet (24% energy from fat)	Acute exercise reduced MRS-measured IMCL by ~20% Increased repletion of IMCL (~142%) with post-exercise normal diet than low-fat diet	[71]
Nine active men Cycling at 75% VO2max for 2 h	MRS pre- and, 0.5 h post, and 1 h post-acute exercise Muscle biopsy pre- and post-acute exercise	IMCL measured by ORO reduced by 31% IMCL measured by MRS declined by 47% IMCL measured by biochemical analysis did not significantly change – concern about fiber type variability between samples affecting results	[69]
17M/19F - recreationally active to endurance trained Cycling at 63% VO2max for 90 min	Muscle biopsy pre- and post-acute exercise	IMCL measured by EM decreased (~14–18%) IMCL decrease due to reducing LD density. LD size did not change	[74]
Eight non-obese, premenopausal women Cross over study with one session of 90 min exercise (45 min treadmill + 45 min cycle ergometer) at 65% VO2max vs no exercise	Acute exercise/no exercise was followed by 15-h lipid infusion Muscle biopsy performed after 13 h lipid infusion	Lipid infusion during acute exercise increased IMCL (50% greater) than lipid infusion during no exercise Increased levels of proteins involved in muscle triglyceride synthesis [mGPAT (35%), DGAT1 (25%) and SCD1(35%)] observed with acute exercise	[76]
Seven men (untrained) Cycling at 65% VO2max for 60 min	Muscle biopsy pre- and post-acute exercise	IMCL measured by ORO reduced (~50%) in Type 1 fibers only IMCL reduction due to decreased LD density (~50% decrease) and LD size (~11%) No change in IMCL in Type 2 fibers IMCL reduction primarily seen in PLIN2+ LD	[68]
16 sedentary men tested before and after 6-week training program [sprint interval training (n = 8) or endurance training (n = 8)] Cycling at 65% VO2max for 60 min	Muscle biopsy before and after acute exercise Acute exercise testing before and after 6 weeks of training	With acute exercise, IMCL reduced by 15–17% in Type 1 fibers (pretraining program), due to reduced LD density With acute exercise, IMCL reduced by 27–43% in Type 1 fibers (post 6-week training program) due to reduced LD density Chronic training increased IMCL, due to increased IMCL density. IMCL size did not change.	[58]
Ten trained men 20 km cross-country skiing time trial (~57 min)	Muscle biopsy pre- and post-time trial from arm (triceps brachii) and leg (vastus lateralis)	IMCL measured by EM Acute exercise did not alter LD size but reduced LD density Arms: IMCL declined (53%) with acute exercise (reduced IMF LD density~51 −58%) Legs: IMCL did not change with acute exercise	[15]
21 males (recreationally active, seven young lean, seven old lean, seven old overweight) Cycling at 50% VO2max for 60 min	Muscle biopsy before and after acute exercise	IMCL measured by EM showed acute exercise reducing IMCL (~20–30%) primarily due to reduced IMF LD droplet density (20–30%) In the old/overweight group, exercise increased LD size in the SS (~25%) and IMF (~7%) region	[6]
34 participants [14 (9M/5F) obese, sedentary controls, 15 (11M/4F) with T2DM, 15 (11M/4F) endurance trained Cycling at 50% of VO2max for 90 min	Muscle biopsy pre- and post-acute exercise (immediate and after 2 h recovery) Stable isotope tracer to measure IMTG synthesis	Acute exercise decreased IMTG in trained participants only (25%), which remained low after 2 h recovery Acute exercise did not alter IMCL in the obese or T2DM groups IMTG fractional synthesis rate was increased with acute exercise (T2DM group), with trend to increase (obese group) and significantly decreased during recovery	[56]
Acute exercise effects on IMCL microenvironment
Exercise	Study design	Main outcome	Refs
17M/19F – recreationally active to endurance trained Cycling at 63% VO2max for 90 min	Muscle biopsy pre- and post-exercise EM evaluation of biopsy samples	IMCL effects previously described. Only women increased contact (~10%) between LD and mitochondria post exercise	[74]
Seven men (untrained) Cycling at 65% VO2max for 60 min	Muscle biopsy before and after exercise	IMCL effects previously described Reduction primarily seen in PLIN2+ LD	[68]
16 sedentary men tested before and after 6-week training program [sprint interval training (n = 8) or endurance training (n = 8)] Cycling at 65% VO2max for 60 min	Muscle biopsy before and after acute exercise Acute exercise testing before and after 6 weeks of training	IMCL effects previously described Preferential loss of IMCL from PLIN2+ LD and PLIN5+ LD	[58]
Ten trained men 20 km cross-country skiing time trial (~57 min)	Muscle biopsy pre- and post-time trial from arm (triceps brachii) and leg (vastus lateralis)	IMCL effects previously described Exercise increased LD-mitochondria contact (+4%) in IMF region of Type 1 fibers in the leg	[15]
Eight trained men Cycling at 59% VO2max for 60 min	Muscle biopsy before and after acute exercise	Acute exercise increased colocalization of HSL with LD (+19%), but not ATGL Acute exercise increased colocalization of HSL preferentially to PLIN5+ LD (+53%) than PLIN5- LD (+34%)	[75]

^aAbbreviations: DGAT, diacylglycerol acyltransferase; EM, electron microscopy; mGPAT, mitochondrial glycerol-3-phosphate acyltransferase; MRS, magnetic resonance spectroscopy; ORO, Oil-Red-O; SCD1, sterol CoA desaturase 1.

IMCL is an important source of energy during exercise [67]. Acute exercise reduces IMCL as documented by multiple modalities, including oil-red-o staining of muscle biopsy samples [68–70], electron microscopy [15], and magnetic resonance spectroscopy [69,71]. This reduction is due to increased oxidation [72], with higher fat intake post-exercise resulting in elevated IMCL repletion post-exercise [71]. In general, acute moderate exercise decreases IMCL by 20–70% [56,68–70]. Yet, this reduction is not always observed, and may be metabolic-status dependent, as acute exercise reduces IMCL in young lean adults, but not in older obese adults [6]. Similarly, acute cycling reduced intramuscular triglyceride (IMTG) in trained athletes, but not obese adults or T2DM patients [56]. This apparent paradox could be potentially explained by increased adipose tissue lipolysis, resulting in elevated circulating NEFAs that can supply IMCL [6]. An alternative explanation for this observation in patients with obesity/T2DM, could be reduced muscle mitochondrial content/mitochondrial oxidative capacity, although this topic remains debatable [73].

Exercise-induced IMCL reduction likely depends on the LD environment. This reduction in IMCL predominantly occurs in Type 1 fibers [58,70] and is primarily due to reducing LD density [6,15,58,74], although modest reductions in LD size have also been reported [68,70]. The acute exercise-induced decrease in IMCL also depends on PLIN proteins, as the decline in LD is particularly prominent in PLIN2+ LD and PLIN5+ LD within Type 1 fibers [58]. Subsequent work demonstrates HSL preferentially colocalizes with PLIN5+ LD after acute exercise [75], supporting a role for PLIN related LD breakdown. Within the muscle fiber, acute exercise differentially alters IMCL distribution within the IMF and SS regions. Generally, acute exercise reduces IMF lipid levels, whereas SS lipid content remains unchanged [6,15]. However, exercise-induced alterations to the IMF lipid compartment may depend on clinical phenotypes, as acute exercise reduced IMF lipid content in young lean individuals, and increased IMF and SS lipid levels in old, obese subjects, which was attributed to increased circulating NEFA levels arising from elevated lipolysis [6]. Additionally, several studies have shown increased contact between LD and mitochondria after acute exercise [12,15,74]. This increased proximity may be physiologically important in facilitating LD expansion/repletion [34] during the post exercise period.

In addition to enhancing IMCL breakdown, acute exercise may also enhance IMCL synthesis. In humans, one session of acute exercise with post-exercise lipid and heparin infusion, enhanced IMTG accumulation in muscle and expression of enzymes associated with IMTG synthesis (mitochondrial glycerol-3-phosphate, DAG acyltransferase 1, and stearoyl-CoA desaturase 1) [76]. Interestingly, lipid-induced insulin resistance, accumulation of FA metabolites (DAG), and activation of proinflammatory pathways (pJNK, IKK/NF-KB), were reduced despite the exercise-induced lipid accretion [76]. Bergman et al., infused U¹³C palmitate, to examine muscle lipid metabolism in response to acute exercise across a range of clinical phenotypes, and found that acute exercise significantly increased IMTG fractional synthesis rate in T2DM individuals, showed a trend (P = 0.08) in obese participants, and did not alter IMTG fractional synthesis rates in trained athletes [56].

In summary, the literature suggests acute exercise primarily reduces IMCL by reducing IMF LD density in Type 1 fibers [15,58,74]. Additionally, acute exercise may also augment LD contact with mitochondria [15,74], and enhance muscle lipid synthesis [56,76].

Concluding Remarks and Future Perspectives

The recent literature highlights several important avenues for future research (see Outstanding Questions). The first avenue involves subpopulations of LDs and mitochondria. Interactions between mitochondria and LDs are heavily involved in lipid metabolism, and recent studies in brown adipose tissue have revealed major bioenergetic differences between peridroplet and cytoplasmic mitochondrial populations [34]. Surprisingly, peridroplet mitochondria prefer pyruvate utilization to fuel enhanced oxidative phosphorylation, driving FA esterification into triglycerides and LD expansion. This challenges the traditional dogma of LD-mitochondrial interactions promoting efficient FA partitioning toward β-oxidation, especially during times of energetic demand [33,77]. Additionally, these metabolic differences between the two mitochondrial subpopulations, are accompanied by alterations in markers associated with mitochondrial fusion and fission activity [34]. Thus, distinguishing the peridroplet and cytoplasmic mitochondrial populations will be important in future studies investigating their roles in skeletal muscle.

Outstanding Questions.

What are the factors that determine formation of LD in the SS and IMF compartments, and can they be modified or influenced pharmacologically?

Are the IMF, SS, or both LD pools, involved in nuclear signaling in skeletal muscle?

Are other signaling pathways influenced by LD-derived lipids?

What are the molecular mechanisms regulating the alterations to mitochondrial dynamics in the peridroplet mitochondria?

Can intersubpopulation transitions occur between cytoplasmic and peridroplet mitochondria, with concomitant bioenergetic alterations, (i.e., can cytoplasmic mitochondria, which emphasize oxidative metabolism, associate with LDs and undergo bioenergetic alterations to support LD expansion)?

How does LD composition and distinct lipolytically-derived FAs alter epigenetic signatures and, conversely, how do epigenetically programmed phenotypes influence LD composition and metabolism?

Similarly, there remains a need to differentiate between SS and IMF LDs, given the clear proteomic and biochemical differences between IMF and SS mitochondria [14]. Differential regulation of PLIN5, which is found at the interface of LDs and mitochondria, provides direct linkage between the organelles [24,78], and may influence the differences in LD subpopulations in skeletal muscle. Although PLIN5 is present in both the SS and IMF mitochondria, PLIN5 protein content is positively correlated to the protein abundance of complexes involved in oxidative phosphorylation and mitochondrial fat oxidation [79]. Moreover, exercise redistributes HSL preferentially to PLIN5+LDs than PLIN2+LDs, suggesting PLIN5+LDs are more involved in lipolytic activity [75]. LD proteins may also affect LD-mitochondrial localization. SNAP23 is a protein involved in the LD-mitochondrial interface [80]. In non-muscle cells, SNAP23 knockdown (3T3 fibroblasts), reduced the localization PLIN2+LD with mitochondria [80], whereas another study reported that SNAP23 knockdown (AML12 cells) did not affect localization of PLIN5+LD with mitochondria [78]. In human skeletal muscle, SNAP23, as measured by immunofluorescence microscopy, colocalizes with the plasma membrane, LDs, and mitochondria [81]; however, the role of SNAP23 in muscle remains to be determined. Therefore, an important area for future investigation will include the differential effect of the SS/IMF compartmentalization on regulation of LD-related proteins and mitochondria function, while considering the influence of the muscle microenvironment (muscle group and fiber type) and metabolic status (resting/acute stimulation and sedentary/trained).

Another important research direction will be characterizing the role of LD-derived FA on nuclear signaling. Lipolytically-derived FAs play a pivotal role in regulating signaling cascades and transcriptional programs pertaining to lipid metabolism and mitochondrial function [23], underscoring the growing understanding of LD interactions with the nucleus. FAs can influence gene transcription through histone modifications, as lipids provide most acetyl groups via FA oxidation [82], and short chain FAs are a source for acylation, such as crotonylation [83]. Moreover, in vitro and in vivo studies have investigated the effects of FAs on DNA methylation and histone modifications to alter metabolic phenotypes [84]. Future studies exploring the interdependence of LDs and the epigenome will likely provide key insights into LD-nuclear signaling mechanisms.

In conclusion, skeletal muscle LDs are heavily influenced by multiple factors, including muscle tissue environment (fiber type and subcellular locations), metabolic status, and stimulus type. However, the understanding of the regulatory mechanisms conferring the context-dependent differences of lipid deposition and utilization in skeletal muscle remains elementary. Further characterization of organellar interactions, and cellular signaling networks involving the LD as a signaling node, will provide critical new insights for skeletal muscle lipid metabolism and its relationship to metabolic dysfunction and flexibility.

Highlights.

In a skeletal muscle context, microenvironments comprised of fiber type and region within the myofiber, result in lipid droplet (LD) compartmentalization.

Acute and chronic exercise can differentially alter muscle LD stores within compartments, and promote mitochondrial-LD interactions, but the exercise-induced effects may depend on metabolic status.

Traditionally viewed as an energy storage organelle, the LD also functions as a critical signaling node, involved in regulating transcriptional programs affiliated with mitochondrial biogenesis, and fatty acid (FA) oxidation.