Next Article in Journal
Ncx3-Induced Mitochondrial Dysfunction in Midbrain Leads to Neuroinflammation in Striatum of A53t-α-Synuclein Transgenic Old Mice
Next Article in Special Issue
Liquid Biopsy for Cancer Cachexia: Focus on Muscle-Derived microRNAs
Previous Article in Journal
Heme Burden and Ensuing Mechanisms That Protect the Kidney: Insights from Bench and Bedside
Previous Article in Special Issue
Common Muscle Metabolic Signatures Highlight Arginine and Lysine Metabolism as Potential Therapeutic Targets to Combat Unhealthy Aging
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 22
Issue 15
10.3390/ijms22158176
ijms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Phospholipids: Identification and Implication in Muscle Pathophysiology

by
Rezlène Bargui
1,
Audrey Solgadi
2,
Bastien Prost
2,
Mélanie Chester
1,
Ana Ferreiro
1,3,
Jérôme Piquereau
4 and
Maryline Moulin
1,*
1
Basic and Translational Myology Laboratory, CNRS UMR8251, Université de Paris, F-75013 Paris, France
2
UMS-IPSIT-SAMM, Université Paris-Saclay, INSERM, CNRS, Ingénierie et Plateformes au Service de l’Innovation Thérapeutique, F-92296 Châtenay-Malabry, France
3
AP-HP, Reference Centre for Neuromuscular Disorders, Institute of Myology, Pitié-Salpêtrière Hospital, F-75013 Paris, France
4
Signalling and Cardiovascular Pathophysiology, INSERM UMR1180, Université Paris-Saclay, F-92296 Châtenay-Malabry, France
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2021, 22(15), 8176; https://0-doi-org.brum.beds.ac.uk/10.3390/ijms22158176
Submission received: 15 July 2021 / Revised: 24 July 2021 / Accepted: 26 July 2021 / Published: 30 July 2021
(This article belongs to the Special Issue Novel Molecular Approaches to Skeletal Muscle Disease and Disuse)
Browse Figures
Review Reports Versions Notes

Abstract

:
Phospholipids (PLs) are amphiphilic molecules that were essential for life to become cellular. PLs have not only a key role in compartmentation as they are the main components of membrane, but they are also involved in cell signaling, cell metabolism, and even cell pathophysiology. Considered for a long time to simply be structural elements of membranes, phospholipids are increasingly being viewed as sensors of their environment and regulators of many metabolic processes. After presenting their main characteristics, we expose the increasing methods of PL detection and identification that help to understand their key role in life processes. Interest and importance of PL homeostasis is growing as pathogenic variants in genes involved in PL biosynthesis and/or remodeling are linked to human diseases. We here review diseases that involve deregulation of PL homeostasis and present a predominantly muscular phenotype.

1. Introduction

Life is amazing. The organization of ‘only’ four major types of biological macromolecules constitutes the majority of the dry weight of the human body. Specifically, carbohydrates, proteins, nucleic acids, and lipids perform an incredible variety of functions in the organism. Lipids, also previously known as ‘lipin’ and ‘lipoid’, are a huge family without a real consensual definition from a chemical or biological point of view. In 1920, Bloor from the department of Biochemistry and Pharmacology of The University of California, proposed a classification of lipids in three groups, simple lipids with fats, oils and waxes; compound lipids including phospholipids (PLs) and glycolipids; and a last group including derived lipids (fatty acids (FA), steroids, etc.) [1]. Of all the different lipids, PLs were indispensable to the appearance of life since they can delimit cellular compartments [2,3]. After presentation of the basic concepts on PLs, we present how to isolate, separate, and identify these molecules. The importance of PLs in numerous biological functions is highlighted by rare genetic diseases that involved deregulation of PLs homeostasis. We will center this review on diseases with a predominantly muscular phenotype.

2. Basic Concepts on PLs

Phospholipids include phosphatidic acid (PA), phosphatidylglycerol (PG), phosphatiylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol (PI), phosphatidylserine (PS), cardiolipin (CL), and sphingomyelin (SM). Among the PLs, the most abundant are glycerophospholipids (or phosphoglycerides), whose common structure comprises a polar head containing a glycerol to which is attached a phosphate group carrying an additional molecule (which gives its identity to the lipid) and two fatty acids (hydrophobic hydrocarbon chains, sn1 and sn2) [4] (cf. Figure 1). The polar head group can be simply a hydrogene in the case of phosphatidic acid; be an ethanolamine, choline, serine, inositol, or glycerol group; and even a phosphatidylglycerol for cardiolipin. Another abundant type of PL is represented by sphingomyelins, which are phosphosphingolipids (a sub-class of sphingolipids). SM contains a sphingosine (a fatty alcohol containing 18 carbons) bound to phosphocholine and a fatty acid. PLs are specialized lipids, which have multiple names depending on biologist or chemist point of view despite a well-defined but diverse and complex structure. Several classifications of lipids exist according to, for example, their chemical structure or biological origin, but without any real consensus [4]. For example, the fatty acid names depend on the carbon length and the presence of double bonds, but the numbering can be different from the biological or the chemical point of view (cf. Figure 2 and Table 1). Nevertheless, the International Union of Pure and Applied Chemistry (IUPAC) has brought about progress into establishing a chemical nomenclature and terminology for specific scientific fields (https://iupac.org/, accessed on 3 May 2021). Moreover, the database LIPID MAPS (Metabolites and Pathways Strategy—https://www.lipidsmaps.org/, accessed on 3 May 2021) constitutes an important tool for lipidomic analysis, with structures and annotations of bulk lipids [5,6,7]. Importantly, most organic molecules are largely mobile depending on their physical and chemical environment (cf. Figure 2).
The molecular diversity of phospholipids is very large and depends not only on the polar head, the carbon length (short/medium/long/very long), and the number of carbon chains (from one in lyso-PL to four in cardiolipin), but also on the degree of double bonds (responsible for molecular rigidity) and their position (cf. Figure 2). There are also plasmalogens, PL analogs containing an ether or vinyl ether group instead of the usual ester function [8]. The position of a fatty acyl chain in sn1 or sn2 increases the diversity further. Thus, the beautiful and large family of phospholipids already contains more than 40,000 natural molecules [2,9]. Precise and structural identification of lipids in a biological medium has been underway for more than 30 years and is becoming possible thanks to the development of mass spectrometry [10]. Of note, the use of common name of fatty acid should be carefully used unless the position of the double bond and the cis and trans isomeres are fully identified. We advise using the systematic name if no information on double bond position is provided (cf. Table 1).
Ijms 22 08176 g002 550
Figure 2. FA nomenclature. The chemical structure of FA showing the physiological numbering (black), the first double bond being at the third or sixth carbon from the omega start, and the chemical numbering (red) with conventions for the double bond location (E in cis, Z in trans). One of numerous possibilities of 3D representation was obtained thanks to Molview [11]. 3D structure is influenced by local environments such as pH.
Figure 2. FA nomenclature. The chemical structure of FA showing the physiological numbering (black), the first double bond being at the third or sixth carbon from the omega start, and the chemical numbering (red) with conventions for the double bond location (E in cis, Z in trans). One of numerous possibilities of 3D representation was obtained thanks to Molview [11]. 3D structure is influenced by local environments such as pH.
Ijms 22 08176 g002
In order to present the different PLs in more detail, we chose to follow the order of lipid retention following liquid chromatography using a polar stationary phase (polyvinyl alcohol grafted silica column). This allows phospholipids to be separated by polarity, followed by a charged aerosol detector Corona CAD and a high-resolution mass spectrometer [12] (cf. Figure 3 and Table 2). Phosphatidic acid is the simplest PL, one of the least abundant PL present in the plasma membrane, but is central in PL biosynthesis [13]. PA has been involved mainly in membrane biophysical properties and in various signaling pathways such as adipogenesis or autophagy [14]. Three major pathways can generate PA: (i) From the acylation of lyso-PA (LPA) by lysophosphatidic acid acyltransferases (LPAAT) (cf. Figure 4). LPA can be obtained from glycerol-3-phosphate (G3P) thanks to the enzymatic action of glycerol-3-phosphate acyltransferase (GPAT). (ii) From the hydrolysis of phosphatidylcholine by the phospholipase D. (iii) From the phosphorylation of diacylglycerol (DAG) by a diacylglycerol kinase (DGK 10 isoforms in human) [13,15]. Importantly, PA is a major precursor of PL biosynthesis. DAG and CDP-DAG (cytidine diphosphate-DAG) are key intermediates that can be obtained from PA thanks to phosphatidic acid phosphatases (PAP or diacylglycerol-3-phosphate phosphohydrolase as systematic name) and CDP-DAG synthetase (CDS or CTP:phosphatidate cytidylyltransferase as systematic name), respectively [14]. They are two types of PAP, type I encoded by lipin genes and type II mainly found in the plasma membrane known as lipid phosphate phosphatases.
From the liponucleotide CDP-DAG, phosphatidylglycerol and 1,3-bis(sn-3′-phosphatidyl)-sn-glycerol known as cardiolipin and phosphatidylinositol can be synthetized [16]. PG has low abundance, less than 1% of total PLs in mammal cells, and is relatively saturated compared to other PLs [17]. Cardiolipins are unique among the PL family as they are specifically present in mitochondria membranes, especially in inner membrane, and composed of a dimeric phosphatidic acid containing two phosphates and four acyl chains [18] with molecular structure similarity with PG [16,19]. This distinctive four acyl chain composition induces a cone-shaped PL that has specific properties such negative membrane curvature and proton buffering [20]. Moreover, this specific structure leads to an important variety of CL species due to a huge diversity of acyl composition. From the biosynthesized ‘nascent’ CL, there is further remodeling leading to hetero-acylated or homo-acylated CL [16]. Three different enzymes are involved in acyl synthetic transformation: tafazzin (a trans-acylase) and two acyl-transferases MLCAT1 (monolysoCL acyltransferase 1) and ALCAT1 (acylCoA:lysoCL acyltransferase 1) [16]. This lipid dimer with four acyl chains was initially isolated from heart beef as cardiomyocytes are the richest cells in terms of mitochondria [21]. In heart and skeletal muscle, the main cardiolipin specie is composed of four C18:2 acyl chains. The last PL synthesized from CDP-DAG is PI, and the biosynthesis takes place in the ER thanks to PIS (phosphatidylinositol synthase), whereas PG and CL biosynthesis are orchestrated in mitochondria [18]. Despite a relatively minor representation in membrane composition (around 10%), PI is an important PL due to the generation of several phosphorylated derivatives at the inositol ring, which are involved in a plethora of cell signalizations such as membrane trafficking and regulation of protein function [18]. Strikingly, the fatty acyl composition of PI is mainly C18:0/C20:4 in numerous rodent tissues, and the consequences of this specific fatty acyl composition are not well understood [22].
PC and PE, the two most abundant phospholipids, are generated by the Kennedy pathway by which DAG is transferred to CDP-choline and CDP-ethanolamine, respectively (cf. Figure 4). These intermediates are obtained from phospho-choline and phospho-ethanolamine thanks to CTP:phosphocholine cytidylyltransferase (CT) and CTP:phosphoethanolamine PE cytidylyltransferase (ET) [23]. PE can generate PC by methylation reactions by phosphatidylethanolamine N-methyltransferase (PEMT). PE can also be produced specifically in the mitochondria by a phosphatidylserine decarboxylase (PSD) from PS. Two enzyme phosphatidylserine synthase 1 and 2 responsible for base-exchange reactions synthesize PS from PC and PE, respectively [24]. PC are also involved in sphingomyelins (which do not have glycerol in their structure) synthesis in the Golgi from ceramides (produced in the ER) by sphingomyelin synthase (SMS), which transfers phosphocholine from a PC [25].
Details in PL biosynthesis pathways are reviewed in [19,26,27]. Multiple pathways can lead to PL synthesis, re-synthesis, or remodeling using various intermediates, and these pathways could happen in different membranes such as plasma membrane, ER membrane, lysosome/peroxisome membrane, or mitochondria membranes. De-acylation and re-acylation reactions due to a large number of phospholipase enzymes add further difficulties in pinpoint the precise PL composition [28]. Each cell has a unique PL fingerprint, making it difficult to unravel the physiological and pathophysiological significance [18]. The development of methods that track PL movement from their place of synthesis to their final organelle destination will help to decipher their role in cells.
Table
Table 2. Main characteristics of phospholipids.
Table 2. Main characteristics of phospholipids.
Phospholipid Group% of Total Lipids in
Eukaryotic Cells [27]		Main
Biosynthesis
Site		Main Characteristics
Phosphatidic acid
PA		1–2ERThe simplest PL
Main precursor for PL synthesis
Inverse cone
Phosphatidylglycerol
PG		<1MitochondriaPrecursor of cardiolipin
Cardiolipin
CL		2–5MitochondriaPresent only in mitochondrial membranes
Contains four fatty acyl chains
Inverse cone
Phosphatidylinositol
PI		10–15ERSource of inositol 1,4,5 triphosphate (IP3)
Cylinder shape
Phosphatidylethanolamine
PE		15–25ER and also
mitochondria from PS		Second most abundant PL
Inverse cone
Phosphatidylserine
PS		5–10ERMostly present in the inner leaflet of the cell membrane, when externalized they became a signal of recognition for cell phagocytosis by macrophages (characteristic of apoptosis)
Cylinder shape
Lysophosphatidylethanolamine
LPE		NDObtained from
phospholipase action		Contains one fatty acyl chain
Hydrolysis of one fatty acid of a PE by a phospholipase
Phosphatidylcholine
PC		45–55ERThe most abundant (outer leaflet of cell membrane)
Constitutes a reserve of choline and methyl groups
Formerly called lecithin
Cylinder shape
Sphingomyelin
SM		5–10GolgiPhosphocholine bound to ceramide
Abundant on the outer leaflet of the membrane
Cylinder shape
Lysophosphatidylcholine
LPC		NDObtained from
phospholipase action		Contains one fatty acyl chain
Hydrolysis of one fatty acid of a PC by a phospholipase

3. Main Methods for PL Analysis

In 2021, more than 200,000 results were obtained through a PubMed search using the keyword “phospholipids”. Evolution of the number of publications is closely linked to technological progress in the lipid research field, particularly with chromatography techniques (first thin layer, then gas and liquid phase) associated with ever more efficient detectors. Since the early 2000s, metabolomic screening approaches using coupled mass spectrometry have been flourishing.
Liquid–liquid extraction (LLE) is the most widely used method for extracting lipids from a biological medium. The first LLE method was developed by Folch in 1957 using a mixture of chloroform, methanol, and water (8:4:3 v/v/v) [29]. Most of the lipids (from a wide range of polarity) are recovered from the sample [30]. However, PL extraction yield can vary from 50 to 90%, some of them remaining in the aqueous phase [31]. The Folch method was subsequently modified by Bligh and Dyer to increase the proportion of polar lipids extracted by using a 1:2:0.8 v/v/v mixture of chloroform, methanol, and water [32]. Since then, many protocols have been developed to optimize extraction efficiency according to the lipids of interest or the nature of biological samples studied and to limit the use of toxic solvents such as chloroform [33].
The characterization of PL molecular species is mainly performed with mass spectrometry (MS) tools generally coupled with chromatographic techniques. Various separation methods have been developed, depending on the lipid mixture and the questions to answer [34,35]. Gas chromatography (GC), often used for fatty acid analysis, is not suitable for whole PL analysis as PLs are too polar and not very volatile. The overall FA composition can be obtained by GC/MS after hydrolysis of PLs, but this method does not allow for identification of plasmalogens ether or vinyl ether acyl chain. Liquid chromatography (LC) is much more suitable for PL analysis including plasmalogens. Normal phase HPLC (high-performance LC) has the advantage of separating PLs according to the polar head nature (allowing quantification of PL classes on a reduced retention time). This facilitates the structural identification of molecular species; however, it requires the use of hazardous solvents [36]. Reverse phase HPLC separates PL molecular species on the basis of hydrophobicity of FA chains, depending on carbon length and number of unsaturations; this induces mixing of lipid classes and difficulties for the structural characterization of species [37]. A more environmentally friendly approach is the super critical fluid chromatography (SFC), which has the advantages of normal phase HPLC while using eco-compatible (CO2 in super critical phase, H2O) and less toxic solvents (acetonitrile) [38].
Once they are separated by chromatography, mass spectrometry is the most efficient tool for characterizing PLs, which ionize very well in electrospray. The analysis can be done in a non-targeted (profiling) or targeted (semi-quantitative) way [39]. In lipidomic profiling, high-resolution mass spectrometry allows to obtain the atomic composition of the whole molecule (generally in adduct form). If chromatography does not provide information on PL classes, fragmentation spectra are essential to identify the molecular structure precisely. Positive mode fragmentation confirms the nature of the polar head, while negative mode fragmentation gives access to FA composition [40]. Although MS is a powerful tool for structural characterization, it provides limited information on positioning of unsaturation on FA chains [38]. Specific methods have been developed with this aim, using alkaline adducts or ozone as reactant gas combined with targeted fragmentation. However, implementation of theses analyses and spectra interpretation are still highly challenging [35]. In the same way, assignment of sn-positions of PL acyl chains is only possible through validated assays based on monitoring the ratio between fragment ions of positional isomers.
Mass spectrometry coupled with chromatographic techniques are lengthy forms of analysis (several dozen minutes) and consume important volumes of solvents. MS can also be used alone, without prior separation, in shotgun MS analysis [41]. This technique is fast and simple to implement, giving access to a global phospholipidomic profile. Very high spectral resolution is required to identify molecular species with similar masses, and separation of isomers is not possible with shotgun analysis. Simultaneous ionization of hundreds of molecules, with different concentrations and polarities leads to ionization suppression, limiting the visibility of low-concentration and/or poorly ionizable species [42].
Quantification of lipid species remains challenging as molecular standards of each species are not available [38]. FA composition will strongly influence ionization, making analogy with different molecular species not possible [43]. In general, the quantification of PLs by species is semi-quantitative.
Phospholipidomics provides large amount of valuable information that implies statistical analysis like other ‘omics’ approaches. Large-scale profiling gives access to the phospholipidome of cells, tissues, or various samples at a specific time. This PL fingerprint is an important step towards understanding PL metabolism and disturbances.

4. Phospholipids in Biological Membranes

The fluid-mosaic membrane model was proposed from the 1970s by Singer and Nicolson and revisited in 2014 by Nicolson [44,45]. Membranes, interacting with the cytoskeletal network and extracellular matrix, are dynamic structures composed not only of lipids, but also of membrane proteins. Dynamic and reversible micro and even nano-size structures, such as lipid rafts, can be present in the biological membranes [45,46,47]. Membranes are integrated elements within cells, tissues, and organisms; they are sensitive and can react to variation of their environment [45]. We will focus on PLs, the main lipid present in the asymmetrical cell membranes [46]. Their inner leaflet is composed of PS, PA, and PI, which induce a strong negative charge compared to the relatively uncharged outer membrane with a majority of PC and SM [48]. As mentioned previously, the main site of phospholipid biosynthesis is the endoplasmic reticulum membrane, followed by mitochondrial membranes; these are two key organelles for cellular homeostasis, controlling among other things calcium content and ATP production [27]. Vance’s research has shown in the beginning of the 1990s that ‘membrane bridges’ between the ER and mitochondria were able to synthesize a large variety of phospholipids [49,50]. These two organelles are in some places very close (around 10 nm apart), and these connecting zones have been named mitochondria-associated membranes (MAM) or mitochondria–ER connect (MERC) [51,52,53,54]. The MAMs allow for the existence of calcium microdomains [55] and energy microdomains [56], which require proximity between mitochondria and ER, but also exchange of PL, although the transporters associated with certain PL have not been clearly identified [52,54]. The distribution of PL species varies among the different organelles [27,57] and can evolve with changes in the cellular environment and with events such as aging, diet, or the circadian clock [58,59,60].
Membranes are dynamic structures. This is especially well known for mitochondria as they can form a filamentous or a fragmented network [59,61,62]. The inner leaflet of mitochondria is very peculiar since it includes only three major PLs: nearly 40% PC, 40% PE, and more than 15% cardiolipin. This composition, rich in PE of conical shape and CL (four fatty acid chains), allows for the formation of numerous invaginations of the inner membrane, the mitochondrial cristae. Regarding the ER, more than half of its PLs are PCs. The ER is composed of domains that perform multiple functions, leading to a heterogeneous protein distribution. A distinction is made between the smooth ribosome-free ER and the rough ribosome-containing ER, which is involved in protein biosynthesis. Muscle cells possess not only the classic ER but also a vast network of specialized membranes associated with myofibrils called the sarcoplasmic reticulum (SR) [51]. The SR allows the intake, storage, and flow of calcium required for muscle contraction. Avoiding calcium leakage from the ER/SR and uncoupling of mitochondria is essential for optimal cellular function. The integrity of the membranes of these organelles is crucial for this, and phospholipids should play an important role [63].
Another important parameter in membrane composition is cholesterol, particularly for plasma membranes [46]. As the molecular ratio is roughly 0.8 cholesterol per phospholipid in plasma membranes [48], their interaction is surely a key parameter for cell homeostasis, cell signaling, and pathophysiology. Two of the key partners for cholesterol are PC and sphingomyelins [45], which are mainly present in the plasma membrane and lysosomes.
The PL composition of the membranes of the numerous organelles and the plasma membrane is adapted to each cell type to meet different physiological needs [64,65]. Depending on the type of carbon chains of each PL, the membranes will be more or less fluid, flexible, compact, and thick, giving them particular properties [64]. The PLs with saturated fatty acids make the membranes rigid, whereas the membranes that contain fatty acids with several unsaturations will be more fluid [66]. The state (fluid vs. rigid) of the membrane can modify cellular processes such as membrane traffic, vesicle fusion, or formation of lipo-protein complexes at the membrane [64].

5. Importance of PLs in Striated Muscle Functions

The local coupling of the sarco-endoplasmic reticulum and mitochondria, which controls muscle contraction and energy production, appears central to skeletal and cardiac muscle function and dysfunction. Integrity of the membrane organelles is crucial to limit calcium leak from the ER/SR and mitochondrial proton and electron escape. Furthermore, skeletal muscles are subjected to an aperiodic and highly variable mechanical force that is transmitted to membranes [67]. Thus, it is important to study more in detail the membrane composition, particularly the amphiphilic molecules that participate in optimal muscle function. Many questions remain partially unresolved; for example, how are they essential to the dialogue between ER and mitochondria? What links exist between phospholipids and skeletal or cardiac muscle diseases? Is PLs dyshomeostasis a cause or a consequence of muscle dysfunction?
Indeed, PL imbalances can disrupt cell and organelle homeostasis and participate in various pathologies [68]. The indisputable evidence of these points comes from rare genetic diseases. In the Orphanet classification, a group of disorders of PLs, sphingolipids, and fatty acids biosynthesis (ORPHA:352301) was created in 2013 [69,70] and is divided in three sub-groups depending on the predominant phenotypical involvement, namely, (i) central nervous system (ORPHA:352306), (ii) peripheral nerves (ORPHA:352309), and (iii) skeletal muscle (ORPHA:352312). The latter highlights a key role of PLs in muscle physiology with five genetic diseases in which the primary defect directly targets PL: Barth syndrome (ORPHA:111, associated with defects of the TAZ gene encoding tafazzin), congenital cataract-hypertrophic cardiomyopathy-mitochondrial myopathy syndrome also known as Sengers syndrome (ORPHA:1369, due to defects of AGK acylglycerol kinase, SLC25A4 solute carrier family 25 member 4), genetic recurrent myoglobinuria (ORPHA:99845, associated with the Lipin1 gene LPIN1, MTCO1, or MTCO2 mitochondria DNA-encoded cytochrome C oxidase), megaconial congenital muscular dystrophy (ORPHA:280671—CHKB, choline kinase beta), and neutral lipid storage disease (ORPHA:165—ABHD5, abhydrolase domain containing 5, PNPLA2/ATGL patatin-like phospholipase domain-containing 2) (cf. Table 3).
Pathogenic variants in the TAZ gene, encoding a transacylase necessary for cardiolipin remodeling, are responsible for Barth’s syndrome (BTHS), a devastating disease affecting the neuromuscular and metabolic systems, characterized by dilated cardiomyopathy and skeletal myopathy in infants [128]. Despite normal content of PE and PC, decreased levels of long polyunsaturated chains (20:4 and 22:6) and increased of 18:2 (probably linoleic acid) has been described in children with this syndrome [129]. Mitochondria in BTHS have important alterations of the cristae ultrastructure (fold and invagination of the inner mitochondrial membrane) leading to defective ATP production [130,131]. Studies involving patients with Barth syndrome are extremely valuable to highlight the key roles played by CLs in mitochondrial respiratory function. Thus, the alteration of CL maturation in these patients has been shown to lead to an instability of respiratory chain supercomplexes that affects complex I activity, thereby demonstrating the importance of CLs for complex I-containing supercomplex formation and function [132].
Sengers syndrome, due to pathogenic variants in the AGK gene, shares strong similarities with Barth syndrome, including skeletal myopathy and cardiomyopathy (hypertrophic), being lethal within the first year of life in around half of the patients [133]. The AGK kinase is involved in phosphorylation of MAG (monoacylglycerol) to LPA and DAG to PA, which are largely involved in PL biosynthesis. The depletion by siRNA of AGK in PC-3 prostate cancer cells induce a reduction of LPA and PA in mitochondria [134], whereas in HEK293 (human embryonic kidney), no changes were observed [135]. To the best of our knowledge, no phospholipidomic data are available in muscle cells. Moreover, the protein belongs to the translocase of the inner mitochondrial membrane TIM22 complex [136]. This multi-protein complex is involved in the import and assembly of mitochondrial carrier proteins such as ANT (adenine nucleotide translocator) [136].
Pathogenic variants in the CHKB gene, encoding choline kinase beta, an enzyme involved in the synthesis of phosphatidylcholine, cause megaconial congenital muscular dystrophy in humans (associated with developmental delay and autistic behavior) and to rostrocaudal muscular dystrophy in mouse [86,87,88]. Decreased content of this most abundant phospholipid in the eukaryotic membrane leads to giant mitochondria localized in the periphery of muscle fibers [137]. Mitochondrial membrane potential is decreased in patient-derived cells, and a reduction of mitochondrial respiratory complex I activity is observed in their skeletal muscles [138].
Patients with a genetic defect causing loss of function of LPIN1 (gene coding for LIPIN 1) develop myopathy, affecting both skeletal muscle and the heart [81]. Investigations into the links between lipid balance, more specifically phospholipid balance, and skeletal muscle function have confirmed that modification of certain proteins involved in the biosynthesis of PLs influences PLs levels. Loss of Lpin1 in mice alters PL levels in gastrocnemius muscle, with significant increase in PC, PE, PI, PS, and PG levels, as well as in PA levels. LIPIN 1, a Mg2+-dependent phosphatidate phosphatase, thus appears to be an important enzyme in the homeostasis of phospholipids in muscle [81].
Other genes involved in PL, sphingolipid, and fatty acid biosynthesis leading to a predominantly muscular phenotype when mutated in humans could extend the ORPHA:352312 group (cf. Table 3). For example, pathogenic variants of HACD1 (3-hydroxyacyl-CoA dehydratase 1), an ER resident enzyme involved in the synthesis of very long-chain fatty acids, have been associated with congenital myopathies in humans and in dogs [90,91,139]. In Hacd1-deficient mice, skeletal muscle mitochondria exhibit a significant decrease in CL content in the mitochondrial inner membrane [91]. This is associated with important cristae remodeling and strong alterations of mitochondrial coupling, leading to a higher energy dissipation. An in vitro assay showed that enrichment of mitochondrial membranes with CLs rescued the coupling efficiency of ATP synthase and therefore the mitochondrial electron transfer chain function, highlighting the key role of CLs in the respiratory chain [117,121,122,123].
Alteration of PLs is also present in inherited form of myopathies due to pathogenic variant in genes without a direct link to PL biosynthesis (cf. Table 3). In muscle from DMD (Duchenne muscular dystrophy) patients, increased PC and SM have been reported [119]. In EDL muscle from the mdx mice (the murine model of DMD, devoid of dystrophin), an increase PC containing saturated acyl chain (16:0 and 18:0) and a decrease PE and PC containing long and polyunsaturated acyl chain (22:6) and (20:4) were found. Genetic rescue in mdx mice leading to truncated dystrophin expression was able to restore the PL profiles [121]. Additionally, the increased PC/PE ratio in mdx mice is corrected by the transgenic expression of FAS (fatty acid synthase), stearoyl-CoA desaturase-1 (SCD1), and Lipin1 [140]. In dysferlinopathies, muscle diseases due to pathogenic variants in the gene encoding dysferlin, a transmembrane protein implicated in protein trafficking, lipid droplets accumulate in myofibers [141]. Molecular remodeling of the lipidome has been observed in skeletal muscle of BLAJ mouse (a dysferlinopathy model), with increased content of species of sphingomyelin, PC, LPC, PE, and LPE in quadriceps from BLAJ compared to WT [142].
In skeletal muscle, ER stress and the stress response provided by the UPR (unfolding protein response) system can be activated under different conditions, including inherited myopathies. The UPR system may be triggered by the accumulation of misfolded proteins, altered calcium balance, or the presence of reactive oxygen species (ROS) [143,144]. Since PLs are synthesized in the ER and constitute its membranes, a link can exist between alteration of the phospholipid balance and reticulum damage. Indeed, loss of Lpin1 gene function in mice induced, in addition to altered PL composition, a significant increase of different ER stress markers (Bip, Fgf21, Gdf15, HSP90b1, and Edem1) [81]. This study also showed morphological alteration of the mitochondria associated with ER stress [81], consistent with the fact that the maintenance of the phospholipid balance is also ensured by the exchange of PLs between ER and mitochondria via the MAMs, and that mitochondria are a site for the synthesis of certain PLs such as PG or CL.
Thus, PL alterations in mouse muscles can be associated with changes in other pathways, such as ER stress, mitochondrial dysfunction, and disturbed cell homeostasis. Taken together, these changes are associated with different inherited myopathies. Although all the roles of PLs at the muscle level are not yet clearly defined, these molecules seem to be important to maintain cell homeostasis and organelle integrity, ensuring normal muscle function. Therefore, a so-called ‘normal’ composition of PLs or PL homeostasis, particularly in the ER/RS membranes and mitochondria, key structures in muscle, is important for maintain motor function, contraction, muscle development and regeneration.

6. Conclusions

Phospholipids are a large family of diverse lipids with a wide variety of structures and physico-chemical properties. The role of PLs in the physiology and pathophysiology of skeletal muscles has been underanalyzed, although muscle is one of the largest metabolic organs. More efforts should be made to fully understand the mechanisms of PL dyshomeostasis and their pathophysiological relevance, which may further improve our understanding of muscular diseases. In particular, characterizing the mechanisms by which PLs modulate the endoplasmic reticulum and mitochondria functions could contribute to a better understanding of the pathophysiology of inherited or acquired myopathies. Thanks to the development of new generation of advanced lipid analysis technologies, phospholipidomics can now help elucidate the importance of PLs in muscle function. Comprehensive PL profiling is necessary for the discovery of diagnostic biomarkers and new therapeutic targets, as well as for the development of personalized therapeutic approaches to treat or slow the progression of muscle pathologies.

Author Contributions

Writing and editing: R.B., A.S., B.P., M.C., A.F., J.P., and M.M. Conceptualization and supervision: M.M. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by the University of Paris and the National Center for Scientific Research.

Data Availability Statement

Not applicable.

Acknowledgments

We thank all the Ferreiro’s lab members for their continuous support.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Bloor, W.R. Outline of a classification of the lipids. Proc. Soc. Exp. Biol. Med. Soc. Exp. Biol. Med. 1920, 17, 138–140. [Google Scholar] [CrossRef] [Green Version]
  2. Zumbuehl, A. Artificial Phospholipids and Their Vesicles. Langmuir 2019, 35, 10223–10232. [Google Scholar] [CrossRef]
  3. Robertson, J.L. The lipid bilayer membrane and its protein constituents. J. Gen. Physiol. 2018, 150, 1472–1483. [Google Scholar] [CrossRef] [Green Version]
  4. Christie, W.W.; Han, X. Lipid Analysis—Isolation, Separation, Identification and Lipidomic Analysis, 4th ed.; Elsevier: Amsterdam, The Netherlands, 2010; p. 446. [Google Scholar]
  5. Fahy, E.; Subramaniam, S.; Murphy, R.C.; Nishijima, M.; Raetz, C.R.; Shimizu, T.; Spener, F.; van Meer, G.; Wakelam, M.J.; Dennis, E.A. Update of the LIPID MAPS comprehensive classification system for lipids. J. Lipid Res. 2009, 50, S9–S14. [Google Scholar] [CrossRef] [Green Version]
  6. Liebisch, G.; Fahy, E.; Aoki, J.; Dennis, E.A.; Durand, T.; Ejsing, C.S.; Fedorova, M.; Feussner, I.; Griffiths, W.J.; Kofeler, H.; et al. Update on LIPID MAPS classification, nomenclature, and shorthand notation for MS-derived lipid structures. J. Lipid Res. 2020, 61, 1539–1555. [Google Scholar] [CrossRef]
  7. Sud, M.; Fahy, E.; Cotter, D.; Brown, A.; Dennis, E.A.; Glass, C.K.; Merrill, A.H., Jr.; Murphy, R.C.; Raetz, C.R.; Russell, D.W.; et al. LMSD: LIPID MAPS structure database. Nucleic Acids Res. 2007, 35, D527–D532. [Google Scholar] [CrossRef] [Green Version]
  8. Dean, J.M.; Lodhi, I.J. Structural and functional roles of ether lipids. Protein Cell 2018, 9, 196–206. [Google Scholar] [CrossRef] [PubMed]
  9. Kimura, T.; Jennings, W.; Epand, R.M. Roles of specific lipid species in the cell and their molecular mechanism. Prog. Lipid Res. 2016, 62, 75–92. [Google Scholar] [CrossRef]
  10. Wenk, M.R. Lipidomics: New tools and applications. Cell 2010, 143, 888–895. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  11. Smith, T.J. MolView: A program for analyzing and displaying atomic structures on the Macintosh personal computer. J. Mol. Graph. 1995, 13, 122–125, 115. [Google Scholar] [CrossRef]
  12. Moulin, M.; Solgadi, A.; Veksler, V.; Garnier, A.; Ventura-Clapier, R.; Chaminade, P. Sex-specific cardiac cardiolipin remodelling after doxorubicin treatment. Biol. Sex Differ. 2015, 6, 20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Zhukovsky, M.A.; Filograna, A.; Luini, A.; Corda, D.; Valente, C. Phosphatidic acid in membrane rearrangements. FEBS Lett. 2019, 593, 2428–2451. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  14. Lutkewitte, A.J.; Finck, B.N. Regulation of Signaling and Metabolism by Lipin-mediated Phosphatidic Acid Phosphohydrolase Activity. Biomolecules 2020, 10, 1386. [Google Scholar] [CrossRef] [PubMed]
  15. Massart, J.; Zierath, J.R. Role of Diacylglycerol Kinases in Glucose and Energy Homeostasis. Trends Endocrinol. Metab. 2019, 30, 603–617. [Google Scholar] [CrossRef]
  16. Maguire, J.J.; Tyurina, Y.Y.; Mohammadyani, D.; Kapralov, A.A.; Anthonymuthu, T.S.; Qu, F.; Amoscato, A.A.; Sparvero, L.J.; Tyurin, V.A.; Planas-Iglesias, J.; et al. Known unknowns of cardiolipin signaling: The best is yet to come. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2017, 1862, 8–24. [Google Scholar] [CrossRef] [Green Version]
  17. Furse, S. Is phosphatidylglycerol essential for terrestrial life? J. Chem. Biol. 2017, 10, 1–9. [Google Scholar] [CrossRef]
  18. Blunsom, N.J.; Cockcroft, S. Phosphatidylinositol synthesis at the endoplasmic reticulum. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2020, 1865, 158471. [Google Scholar] [CrossRef]
  19. Blunsom, N.J.; Cockcroft, S. CDP-Diacylglycerol Synthases (CDS): Gateway to Phosphatidylinositol and Cardiolipin Synthesis. Front. Cell Dev. Biol. 2020, 8, 63. [Google Scholar] [CrossRef] [Green Version]
  20. Pennington, E.R.; Funai, K.; Brown, D.A.; Shaikh, S.R. The role of cardiolipin concentration and acyl chain composition on mitochondrial inner membrane molecular organization and function. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2019, 1864, 1039–1052. [Google Scholar] [CrossRef]
  21. Pangborn, M.C. Isolation and purification of a serologically acitve phospholipid from beef heart. J. Biol. Chem. 1942, 143, 247–256. [Google Scholar] [CrossRef]
  22. Traynor-Kaplan, A.; Kruse, M.; Dickson, E.J.; Dai, G.; Vivas, O.; Yu, H.; Whittington, D.; Hille, B. Fatty-acyl chain profiles of cellular phosphoinositides. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2017, 1862, 513–522. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. van der Veen, J.N.; Kennelly, J.P.; Wan, S.; Vance, J.E.; Vance, D.E.; Jacobs, R.L. The critical role of phosphatidylcholine and phosphatidylethanolamine metabolism in health and disease. Biochim. Biophys. Acta Biomembr. 2017, 1859 Pt B, 1558–1572. [Google Scholar] [CrossRef]
  24. Kimura, A.K.; Kimura, T. Phosphatidylserine biosynthesis pathways in lipid homeostasis: Toward resolution of the pending central issue for decades. FASEB J. 2021, 35, e21177. [Google Scholar] [CrossRef] [PubMed]
  25. Futerman, A.H.; Riezman, H. The ins and outs of sphingolipid synthesis. Trends Cell Biol. 2005, 15, 312–318. [Google Scholar] [CrossRef]
  26. Funai, K.; Summers, S.A.; Rutter, J. Reign in the membrane: How common lipids govern mitochondrial function. Curr. Opin. Cell Biol. 2020, 63, 162–173. [Google Scholar] [CrossRef] [PubMed]
  27. Vance, J.E. Phospholipid synthesis and transport in mammalian cells. Traffic 2015, 16, 1–18. [Google Scholar] [CrossRef]
  28. Aloulou, A.; Rahier, R.; Arhab, Y.; Noiriel, A.; Abousalham, A. Phospholipases: An Overview. Methods Mol. Biol. 2018, 1835, 69–105. [Google Scholar] [PubMed]
  29. Folch, J.; Lees, M.; Sloane Stanley, G.H. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 1957, 226, 497–509. [Google Scholar] [CrossRef]
  30. Lofgren, L.; Forsberg, G.B.; Stahlman, M. The BUME method: A new rapid and simple chloroform-free method for total lipid extraction of animal tissue. Sci. Rep. 2016, 6, 27688. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  31. Mirzaian, M.; Kramer, G.; Poorthuis, B.J. Quantification of sulfatides and lysosulfatides in tissues and body fluids by liquid chromatography-tandem mass spectrometry. J. Lipid Res. 2015, 56, 936–943. [Google Scholar] [CrossRef] [Green Version]
  32. Bligh, E.G.; Dyer, W.J. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 1959, 37, 911–917. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Zhang, T.; Chen, S.; Syed, I.; Stahlman, M.; Kolar, M.J.; Homan, E.A.; Chu, Q.; Smith, U.; Boren, J.; Kahn, B.B.; et al. A LC-MS-based workflow for measurement of branched fatty acid esters of hydroxy fatty acids. Nat. Protoc. 2016, 11, 747–763. [Google Scholar] [CrossRef] [Green Version]
  34. Cajka, T.; Fiehn, O. Comprehensive analysis of lipids in biological systems by liquid chromatography-mass spectrometry. Trends Anal. Chem. 2014, 61, 192–206. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Hu, T.; Zhang, J.L. Mass-spectrometry-based lipidomics. J. Sep. Sci. 2018, 41, 351–372. [Google Scholar] [CrossRef]
  36. Essaid, D.; Rosilio, V.; Daghildjian, K.; Solgadi, A.; Vergnaud, J.; Kasselouri, A.; Chaminade, P. Artificial plasma membrane models based on lipidomic profiling. Biochim. Biophys. Acta 2016, 1858, 2725–2736. [Google Scholar] [CrossRef]
  37. Schlotterbeck, J.; Chatterjee, M.; Gawaz, M.; Lammerhofer, M. Comprehensive MS/MS profiling by UHPLC-ESI-QTOF-MS/MS using SWATH data-independent acquisition for the study of platelet lipidomes in coronary artery disease. Anal. Chim. Acta 2019, 1046, 1–15. [Google Scholar] [CrossRef]
  38. Holcapek, M.; Liebisch, G.; Ekroos, K. Lipidomic Analysis. Anal. Chem. 2018, 90, 4249–4257. [Google Scholar] [CrossRef] [Green Version]
  39. Lee, H.C.; Yokomizo, T. Applications of mass spectrometry-based targeted and non-targeted lipidomics. Biochem. Biophys. Res. Commun. 2018, 504, 576–581. [Google Scholar] [CrossRef]
  40. Ekroos, K.; Ejsing, C.S.; Bahr, U.; Karas, M.; Simons, K.; Shevchenko, A. Charting molecular composition of phosphatidylcholines by fatty acid scanning and ion trap MS3 fragmentation. J. Lipid Res. 2003, 44, 2181–2192. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  41. Ejsing, C.S.; Sampaio, J.L.; Surendranath, V.; Duchoslav, E.; Ekroos, K.; Klemm, R.W.; Simons, K.; Shevchenko, A. Global analysis of the yeast lipidome by quantitative shotgun mass spectrometry. Proc. Natl. Acad. Sci. USA 2009, 106, 2136–2141. [Google Scholar] [CrossRef] [Green Version]
  42. Yang, K.; Han, X. Lipidomics: Techniques, Applications, and Outcomes Related to Biomedical Sciences. Trends Biochem. Sci. 2016, 41, 954–969. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Khoury, S.; El Banna, N.; Tfaili, S.; Chaminade, P. A study of inter-species ion suppression in electrospray ionization-mass spectrometry of some phospholipid classes. Anal. Bioanal. Chem. 2016, 408, 1453–1465. [Google Scholar] [CrossRef] [PubMed]
  44. Singer, S.J.; Nicolson, G.L. The fluid mosaic model of the structure of cell membranes. Science 1972, 175, 720–731. [Google Scholar] [CrossRef]
  45. Nicolson, G.L. The Fluid-Mosaic Model of Membrane Structure: Still relevant to understanding the structure, function and dynamics of biological membranes after more than 40 years. Biochim. Biophys. Acta 2014, 1838, 1451–1466. [Google Scholar] [CrossRef] [Green Version]
  46. Doktorova, M.; Symons, J.L.; Levental, I. Structural and functional consequences of reversible lipid asymmetry in living membranes. Nat. Chem. Biol. 2020, 16, 1321–1330. [Google Scholar] [CrossRef] [PubMed]
  47. Doktorova, M. On the Long and Winding Road to a Perfect Membrane Model. Biophys. J. 2020, 118, 273–275. [Google Scholar] [CrossRef]
  48. Steck, T.L.; Lange, Y. Transverse distribution of plasma membrane bilayer cholesterol: Picking sides. Traffic 2018, 19, 750–760. [Google Scholar] [CrossRef] [Green Version]
  49. Vance, J.E. Phospholipid synthesis in a membrane fraction associated with mitochondria. J. Biol. Chem. 1990, 265, 7248–7256. [Google Scholar] [CrossRef]
  50. Vance, J.E. Newly made phosphatidylserine and phosphatidylethanolamine are preferentially translocated between rat liver mitochondria and endoplasmic reticulum. J. Biol. Chem. 1991, 266, 89–97. [Google Scholar] [CrossRef]
  51. Boncompagni, S.; Pozzer, D.; Viscomi, C.; Ferreiro, A.; Zito, E. Physical and Functional Cross Talk Between Endo-Sarcoplasmic Reticulum and Mitochondria in Skeletal Muscle. Antioxid. Redox Signal. 2019, 32, 873–883. [Google Scholar] [CrossRef]
  52. Giacomello, M.; Pellegrini, L. The coming of age of the mitochondria-ER contact: A matter of thickness. Cell Death Differ. 2016, 23, 1417–1427. [Google Scholar] [CrossRef]
  53. Heden, T.D.; Neufer, P.D.; Funai, K. Looking Beyond Structure: Membrane Phospholipids of Skeletal Muscle Mitochondria. Trends Endocrinol. Metab. 2016, 27, 553–562. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Vance, J.E. MAM (mitochondria-associated membranes) in mammalian cells: Lipids and beyond. Biochim. Biophys. Acta 2014, 1841, 595–609. [Google Scholar] [CrossRef] [PubMed]
  55. Dorn, G.W., 2nd; Maack, C. SR and mitochondria: Calcium cross-talk between kissing cousins. J. Mol. Cell. Cardiol. 2013, 55, 42–49. [Google Scholar] [CrossRef]
  56. Piquereau, J.; Novotova, M.; Fortin, D.; Garnier, A.; Ventura-Clapier, R.; Veksler, V.; Joubert, F. Postnatal development of mouse heart: Formation of energetic microdomains. J. Physiol. 2010, 588 Pt 13, 2443–2454. [Google Scholar] [CrossRef]
  57. Yang, Y.; Lee, M.; Fairn, G.D. Phospholipid subcellular localization and dynamics. J. Biol. Chem. 2018, 293, 6230–6240. [Google Scholar] [CrossRef] [Green Version]
  58. Adhyapak, P.; Kapoor, S. Membrane Dynamics in Health and Disease: Impact on Cellular Signalling. J. Membr. Biol. 2019, 252, 213–226. [Google Scholar] [CrossRef] [PubMed]
  59. Giacomello, E.; Crea, E.; Torelli, L.; Bergamo, A.; Reggiani, C.; Sava, G.; Toniolo, L. Age Dependent Modification of the Metabolic Profile of the Tibialis Anterior Muscle Fibers in C57BL/6J Mice. Int. J. Mol. Sci. 2020, 21, 3923. [Google Scholar] [CrossRef] [PubMed]
  60. Nohara, K.; Kim, E.; Wirianto, M.; Mileykovskaya, E.; Dowhan, W.; Chen, Z.; Yoo, S.H. Cardiolipin Synthesis in Skeletal Muscle Is Rhythmic and Modifiable by Age and Diet. Oxidative Med. Cell. Longev. 2020, 2020, 5304768. [Google Scholar] [CrossRef]
  61. Janikiewicz, J.; Szymanski, J.; Malinska, D.; Patalas-Krawczyk, P.; Michalska, B.; Duszynski, J.; Giorgi, C.; Bonora, M.; Dobrzyn, A.; Wieckowski, M.R. Mitochondria-associated membranes in aging and senescence: Structure, function, and dynamics. Cell Death Dis. 2018, 9, 332. [Google Scholar] [CrossRef]
  62. Yaffe, M.P. Dynamic mitochondria. Nat. Cell Biol. 1999, 1, E149–E150. [Google Scholar] [CrossRef]
  63. Bozdaganyan, M.E.; Lokhmatikov, A.V.; Voskoboynikova, N.; Cherepanov, D.A.; Steinhoff, H.J.; Shaitan, K.V.; Mulkidjanian, A.Y. Proton leakage across lipid bilayers: Oxygen atoms of phospholipid ester linkers align water molecules into transmembrane water wires. Biochim. Biophys. Acta Bioenerg. 2019, 1860, 439–451. [Google Scholar] [CrossRef] [PubMed]
  64. Hishikawa, D.; Valentine, W.J.; Iizuka-Hishikawa, Y.; Shindou, H.; Shimizu, T. Metabolism and functions of docosahexaenoic acid-containing membrane glycerophospholipids. FEBS Lett. 2017, 591, 2730–2744. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. van Meer, G.; de Kroon, A.I. Lipid map of the mammalian cell. J. Cell Sci. 2011, 124 Pt 1, 5–8. [Google Scholar] [CrossRef] [Green Version]
  66. Tyler, A.I.I.; Greenfield, J.L.; Seddon, J.M.; Brooks, N.J.; Purushothaman, S. Coupling Phase Behavior of Fatty Acid Containing Membranes to Membrane Bio-Mechanics. Front. Cell Dev. Biol. 2019, 7, 187. [Google Scholar] [CrossRef] [PubMed]
  67. McNeil, P.L.; Steinhardt, R.A. Plasma membrane disruption: Repair, prevention, adaptation. Annu. Rev. Cell Dev. Biol. 2003, 19, 697–731. [Google Scholar] [CrossRef] [PubMed]
  68. Patel, D.; Witt, S.N. Ethanolamine and Phosphatidylethanolamine: Partners in Health and Disease. Oxidative Med. Cell. Longev. 2017, 2017, 4829180. [Google Scholar] [CrossRef] [Green Version]
  69. Lamari, F.; Mochel, F.; Sedel, F.; Saudubray, J.M. Disorders of phospholipids, sphingolipids and fatty acids biosynthesis: Toward a new category of inherited metabolic diseases. J. Inherit. Metab. Dis. 2013, 36, 411–425. [Google Scholar] [CrossRef]
  70. Wortmann, S.B.; Espeel, M.; Almeida, L.; Reimer, A.; Bosboom, D.; Roels, F.; de Brouwer, A.P.; Wevers, R.A. Inborn errors of metabolism in the biosynthesis and remodelling of phospholipids. J. Inherit. Metab. Dis. 2015, 38, 99–110. [Google Scholar] [CrossRef] [PubMed]
  71. Barth, P.G.; Valianpour, F.; Bowen, V.M.; Lam, J.; Duran, M.; Vaz, F.M.; Wanders, R.J. X-linked cardioskeletal myopathy and neutropenia (Barth syndrome): An update. Am. J. Med. Genet. A 2004, 126A, 349–354. [Google Scholar] [CrossRef]
  72. Damschroder, D.; Reynolds, C.; Wessells, R. Drosophila tafazzin mutants have impaired exercise capacity. Physiol. Rep. 2018, 6, e13604. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Zegallai, H.M.; Hatch, G.M. Barth syndrome: Cardiolipin, cellular pathophysiology, management, and novel therapeutic targets. Mol. Cell. Biochem. 2021, 476, 1605–1629. [Google Scholar] [CrossRef] [PubMed]
  74. Mayr, J.A.; Haack, T.B.; Graf, E.; Zimmermann, F.A.; Wieland, T.; Haberberger, B.; Superti-Furga, A.; Kirschner, J.; Steinmann, B.; Baumgartner, M.R.; et al. Lack of the mitochondrial protein acylglycerol kinase causes Sengers syndrome. Am. J. Hum. Genet. 2012, 90, 314–320. [Google Scholar] [CrossRef] [Green Version]
  75. Sengers, R.C.; Trijbels, J.M.; Willems, J.L.; Daniels, O.; Stadhouders, A.M. Congenital cataract and mitochondrial myopathy of skeletal and heart muscle associated with lactic acidosis after exercise. J. Pediatrics 1975, 86, 873–880. [Google Scholar] [CrossRef]
  76. Jiang, H.; Yu, Z.; Ding, N.; Yang, M.; Zhang, L.; Fan, X.; Zhou, Y.; Zou, Q.; Hou, J.; Zheng, J.; et al. The role of AGK in thrombocytopoiesis and possible therapeutic strategies. Blood 2020, 136, 119–129. [Google Scholar] [CrossRef]
  77. Allali, S.; Dorboz, I.; Samaan, S.; Slama, A.; Rambaud, C.; Boespflug-Tanguy, O.; Sarret, C. Mutation in the AGK gene in two siblings with unusual Sengers syndrome. Metab. Brain Dis. 2017, 32, 2149–2154. [Google Scholar] [CrossRef]
  78. Zeharia, A.; Shaag, A.; Houtkooper, R.H.; Hindi, T.; de Lonlay, P.; Erez, G.; Hubert, L.; Saada, A.; de Keyzer, Y.; Eshel, G.; et al. Mutations in LPIN1 cause recurrent acute myoglobinuria in childhood. Am. J. Hum. Genet. 2008, 83, 489–494. [Google Scholar] [CrossRef] [Green Version]
  79. Donkor, J.; Sariahmetoglu, M.; Dewald, J.; Brindley, D.N.; Reue, K. Three mammalian lipins act as phosphatidate phosphatases with distinct tissue expression patterns. J. Biol. Chem. 2007, 282, 3450–3457. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  80. Csaki, L.S.; Reue, K. Lipins: Multifunctional lipid metabolism proteins. Annu. Rev. Nutr. 2010, 30, 257–272. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  81. Rashid, T.; Nemazanyy, I.; Paolini, C.; Tatsuta, T.; Crespin, P.; de Villeneuve, D.; Brodesser, S.; Benit, P.; Rustin, P.; Baraibar, M.A.; et al. Lipin1 deficiency causes sarcoplasmic reticulum stress and chaperone-responsive myopathy. EMBO J. 2019, 38, e99576. [Google Scholar] [CrossRef]
  82. Reilich, P.; Horvath, R.; Krause, S.; Schramm, N.; Turnbull, D.M.; Trenell, M.; Hollingsworth, K.G.; Gorman, G.S.; Hans, V.H.; Reimann, J. The phenotypic spectrum of neutral lipid storage myopathy due to mutations in the PNPLA2 gene. J. Neurol. 2011, 258, 1987–1997. [Google Scholar] [CrossRef]
  83. Wu, J.W.; Wang, S.P.; Alvarez, F.; Casavant, S.; Gauthier, N.; Abed, L.; Soni, K.G.; Yang, G.; Mitchell, G.A. Deficiency of liver adipose triglyceride lipase in mice causes progressive hepatic steatosis. Hepatology 2011, 54, 122–132. [Google Scholar] [CrossRef] [PubMed]
  84. Rao, M.; Guo, G.; Li, M.; Chen, S.; Chen, K.; Chen, X.; Song, J.; Hu, S. The homozygous variant c.245G > A/p.G82D in PNPLA2 is associated with arrhythmogenic cardiomyopathy phenotypic manifestations. Clin. Genet. 2019, 96, 532–540. [Google Scholar] [CrossRef] [PubMed]
  85. Li, M.; Hirano, K.I.; Ikeda, Y.; Higashi, M.; Hashimoto, C.; Zhang, B.; Kozawa, J.; Sugimura, K.; Miyauchi, H.; Suzuki, A.; et al. Triglyceride deposit cardiomyovasculopathy: A rare cardiovascular disorder. Orphanet J Rare Dis 2019, 14, 134. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Sher, R.B.; Aoyama, C.; Huebsch, K.A.; Ji, S.; Kerner, J.; Yang, Y.; Frankel, W.N.; Hoppel, C.L.; Wood, P.A.; Vance, D.E.; et al. A rostrocaudal muscular dystrophy caused by a defect in choline kinase beta, the first enzyme in phosphatidylcholine biosynthesis. J. Biol. Chem. 2006, 281, 4938–4948. [Google Scholar] [CrossRef] [Green Version]
  87. Mitsuhashi, S.; Hatakeyama, H.; Karahashi, M.; Koumura, T.; Nonaka, I.; Hayashi, Y.K.; Noguchi, S.; Sher, R.B.; Nakagawa, Y.; Manfredi, G.; et al. Muscle choline kinase beta defect causes mitochondrial dysfunction and increased mitophagy. Hum. Mol. Genet. 2011, 20, 3841–3851. [Google Scholar] [CrossRef]
  88. Mitsuhashi, S.; Ohkuma, A.; Talim, B.; Karahashi, M.; Koumura, T.; Aoyama, C.; Kurihara, M.; Quinlivan, R.; Sewry, C.; Mitsuhashi, H.; et al. A congenital muscular dystrophy with mitochondrial structural abnormalities caused by defective de novo phosphatidylcholine biosynthesis. Am. J. Hum. Genet. 2011, 88, 845–851. [Google Scholar] [CrossRef] [Green Version]
  89. Sayed-Zahid, A.A.; Sher, R.B.; Sukoff Rizzo, S.J.; Anderson, L.C.; Patenaude, K.E.; Cox, G.A. Functional rescue in a mouse model of congenital muscular dystrophy with megaconial myopathy. Hum. Mol. Genet. 2019, 28, 2635–2647. [Google Scholar] [CrossRef]
  90. Blondelle, J.; Ohno, Y.; Gache, V.; Guyot, S.; Storck, S.; Blanchard-Gutton, N.; Barthelemy, I.; Walmsley, G.; Rahier, A.; Gadin, S.; et al. HACD1, a regulator of membrane composition and fluidity, promotes myoblast fusion and skeletal muscle growth. J. Mol. Cell Biol. 2015, 7, 429–440. [Google Scholar] [CrossRef] [Green Version]
  91. Prola, A.; Blondelle, J.; Vandestienne, A.; Piquereau, J.; Denis, R.G.P.; Guyot, S.; Chauvin, H.; Mourier, A.; Maurer, M.; Henry, C.; et al. Cardiolipin content controls mitochondrial coupling and energetic efficiency in muscle. Sci. Adv. 2021, 7, eabd6322. [Google Scholar] [CrossRef] [PubMed]
  92. Abbasi-Moheb, L.; Westenberger, A.; Alotaibi, M.; Alghamdi, M.A.; Hertecant, J.L.; Ariamand, A.; Beetz, C.; Rolfs, A.; Bertoli-Avella, A.M.; Bauer, P. Biallelic loss-of-function HACD1 variants are a bona fide cause of congenital myopathy. Clin. Genet. 2021, 99, 513–518. [Google Scholar] [CrossRef]
  93. Gregersen, N. The acyl-CoA dehydrogenation deficiencies. Recent advances in the enzymic characterization and understanding of the metabolic and pathophysiological disturbances in patients with acyl-CoA dehydrogenation deficiencies. Scand. J. Clin. Lab. Investig. Suppl. 1985, 174, 1–60. [Google Scholar]
  94. Strauss, A.W.; Powell, C.K.; Hale, D.E.; Anderson, M.M.; Ahuja, A.; Brackett, J.C.; Sims, H.F. Molecular basis of human mitochondrial very-long-chain acyl-CoA dehydrogenase deficiency causing cardiomyopathy and sudden death in childhood. Proc. Natl. Acad. Sci. USA 1995, 92, 10496–10500. [Google Scholar] [CrossRef] [Green Version]
  95. Tucci, S.; Flogel, U.; Spiekerkoetter, U. Sexual dimorphism of lipid metabolism in very long-chain acyl-CoA dehydrogenase deficient (VLCAD-/-) mice in response to medium-chain triglycerides (MCT). Biochim. Biophys. Acta 2015, 1852, 1442–1450. [Google Scholar] [CrossRef] [Green Version]
  96. Tucci, S.; Alatibi, K.I.; Wehbe, Z. Altered Metabolic Flexibility in Inherited Metabolic Diseases of Mitochondrial Fatty Acid Metabolism. Int. J. Mol. Sci. 2021, 22, 3799. [Google Scholar] [CrossRef]
  97. Vengalil, S.; Preethish-Kumar, V.; Polavarapu, K.; Christopher, R.; Gayathri, N.; Natarajan, A.; Manjunath, M.; Nashi, S.; Prasad, C.; Nalini, A. Fatty acid oxidation defects presenting as primary myopathy and prominent dropped head syndrome. Neuromuscul. Disord. 2017, 27, 986–996. [Google Scholar] [CrossRef] [PubMed]
  98. Stanley, C.A. New genetic defects in mitochondrial fatty acid oxidation and carnitine deficiency. Adv. Pediatrics 1987, 34, 59–88. [Google Scholar]
  99. Tolwani, R.J.; Hamm, D.A.; Tian, L.; Sharer, J.D.; Vockley, J.; Rinaldo, P.; Matern, D.; Schoeb, T.R.; Wood, P.A. Medium-chain acyl-CoA dehydrogenase deficiency in gene-targeted mice. PLoS Genet. 2005, 1, e23. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  100. Lim, S.C.; Tajika, M.; Shimura, M.; Carey, K.T.; Stroud, D.A.; Murayama, K.; Ohtake, A.; McKenzie, M. Loss of the Mitochondrial Fatty Acid beta-Oxidation Protein Medium-Chain Acyl-Coenzyme A Dehydrogenase Disrupts Oxidative Phosphorylation Protein Complex Stability and Function. Sci. Rep. 2018, 8, 153. [Google Scholar] [CrossRef] [Green Version]
  101. Scheuerman, O.; Wanders, R.J.; Waterham, H.R.; Dubnov-Raz, G.; Garty, B.Z. Mitochondrial trifunctional protein deficiency with recurrent rhabdomyolysis. Pediatric Neurol. 2009, 40, 465–467. [Google Scholar] [CrossRef]
  102. Ibdah, J.A.; Paul, H.; Zhao, Y.; Binford, S.; Salleng, K.; Cline, M.; Matern, D.; Bennett, M.J.; Rinaldo, P.; Strauss, A.W. Lack of mitochondrial trifunctional protein in mice causes neonatal hypoglycemia and sudden death. J. Clin. Investig. 2001, 107, 1403–1409. [Google Scholar] [CrossRef] [Green Version]
  103. Fraser, H.; Geppert, J.; Johnson, R.; Johnson, S.; Connock, M.; Clarke, A.; Taylor-Phillips, S.; Stinton, C. Evaluation of earlier versus later dietary management in long-chain 3-hydroxyacyl-CoA dehydrogenase or mitochondrial trifunctional protein deficiency: A systematic review. Orphanet J. Rare Dis. 2019, 14, 258. [Google Scholar] [CrossRef] [Green Version]
  104. Dorfman, M.L.; Hershko, C.; Eisenberg, S.; Sagher, F. Ichthyosiform dermatosis with systemic lipidosis. Arch. Dermatol. 1974, 110, 261–266. [Google Scholar] [CrossRef] [PubMed]
  105. Lefevre, C.; Jobard, F.; Caux, F.; Bouadjar, B.; Karaduman, A.; Heilig, R.; Lakhdar, H.; Wollenberg, A.; Verret, J.L.; Weissenbach, J.; et al. Mutations in CGI-58, the gene encoding a new protein of the esterase/lipase/thioesterase subfamily, in Chanarin-Dorfman syndrome. Am. J. Hum. Genet. 2001, 69, 1002–1012. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  106. Oberer, M.; Boeszoermenyi, A.; Nagy, H.M.; Zechner, R. Recent insights into the structure and function of comparative gene identification-58. Curr. Opin. Lipidol. 2011, 22, 149–158. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  107. Jebessa, Z.H.; Shanmukha, K.D.; Dewenter, M.; Lehmann, L.H.; Xu, C.; Schreiter, F.; Siede, D.; Gong, X.M.; Worst, B.C.; Federico, G.; et al. The lipid droplet-associated protein ABHD5 protects the heart through proteolysis of HDAC4. Nat. Metab. 2019, 1, 1157–1167. [Google Scholar] [CrossRef] [PubMed]
  108. Radner, F.P.; Streith, I.E.; Schoiswohl, G.; Schweiger, M.; Kumari, M.; Eichmann, T.O.; Rechberger, G.; Koefeler, H.C.; Eder, S.; Schauer, S.; et al. Growth retardation, impaired triacylglycerol catabolism, hepatic steatosis, and lethal skin barrier defect in mice lacking comparative gene identification-58 (CGI-58). J. Biol. Chem. 2010, 285, 7300–7311. [Google Scholar] [CrossRef] [Green Version]
  109. North, K.N.; Hoppel, C.L.; De Girolami, U.; Kozakewich, H.P.; Korson, M.S. Lethal neonatal deficiency of carnitine palmitoyltransferase II associated with dysgenesis of the brain and kidneys. J. Pediatrics 1995, 127, 414–420. [Google Scholar] [CrossRef]
  110. Corti, S.; Bordoni, A.; Ronchi, D.; Musumeci, O.; Aguennouz, M.; Toscano, A.; Lamperti, C.; Bresolin, N.; Comi, G.P. Clinical features and new molecular findings in Carnitine Palmitoyltransferase II (CPT II) deficiency. J. Neurol. Sci. 2008, 266, 97–103. [Google Scholar] [CrossRef]
  111. Pereyra, A.S.; Hasek, L.Y.; Harris, K.L.; Berman, A.G.; Damen, F.W.; Goergen, C.J.; Ellis, J.M. Loss of cardiac carnitine palmitoyltransferase 2 results in rapamycin-resistant, acetylation-independent hypertrophy. J. Biol. Chem. 2017, 292, 18443–18456. [Google Scholar] [CrossRef] [Green Version]
  112. Land, J.M.; Mistry, S.; Squier, M.; Hope, P.; Ghadiminejad, I.; Orford, M.; Saggerson, D. Neonatal carnitine palmitoyltransferase-2 deficiency: A case presenting with myopathy. Neuromuscul. Disord. 1995, 5, 129–137. [Google Scholar] [CrossRef]
  113. Begam, M.; Roche, R.; Hass, J.J.; Basel, C.A.; Blackmer, J.M.; Konja, J.T.; Samojedny, A.L.; Collier, A.F.; Galen, S.S.; Roche, J.A. The effects of concentric and eccentric training in murine models of dysferlin-associated muscular dystrophy. Muscle Nerve 2020, 62, 393–403. [Google Scholar] [CrossRef]
  114. Kobayashi, K.; Izawa, T.; Kuwamura, M.; Yamate, J. Dysferlin and animal models for dysferlinopathy. J. Toxicol. Pathol. 2012, 25, 135–147. [Google Scholar] [CrossRef] [Green Version]
  115. Roche, J.A.; Ru, L.W.; Bloch, R.J. Distinct effects of contraction-induced injury in vivo on four different murine models of dysferlinopathy. J. Biomed. Biotechnol. 2012, 2012, 134031. [Google Scholar] [CrossRef] [Green Version]
  116. Tan, S.M.L.; Ong, C.C.; Tan, K.B.; Chin, H.L.; Paliwal, P.R.; Ng, K.W.P.; Lin, W. Subclinical Cardiomyopathy in Miyoshi Myopathy Detected by Late Gadolinium Enhancement Cardiac Magnetic Resonance Imaging. Int. Heart J. 2021, 62, 186–192. [Google Scholar] [CrossRef] [PubMed]
  117. Benabdellah, F.; Yu, H.; Brunelle, A.; Laprevote, O.; De La Porte, S. MALDI reveals membrane lipid profile reversion in MDX mice. Neurobiol. Dis. 2009, 36, 252–258. [Google Scholar] [CrossRef] [PubMed]
  118. Blake, D.J.; Weir, A.; Newey, S.E.; Davies, K.E. Function and genetics of dystrophin and dystrophin-related proteins in muscle. Physiol. Rev. 2002, 82, 291–329. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  119. Dabaj, I.; Ferey, J.; Marguet, F.; Gilard, V.; Basset, C.; Bahri, Y.; Brehin, A.C.; Vanhulle, C.; Leturcq, F.; Marret, S.; et al. Muscle metabolic remodelling patterns in Duchenne muscular dystrophy revealed by ultra-high-resolution mass spectrometry imaging. Sci. Rep. 2021, 11, 1906. [Google Scholar] [CrossRef]
  120. McGreevy, J.W.; Hakim, C.H.; McIntosh, M.A.; Duan, D. Animal models of Duchenne muscular dystrophy: From basic mechanisms to gene therapy. Dis. Models Mech. 2015, 8, 195–213. [Google Scholar] [CrossRef] [Green Version]
  121. Senoo, N.; Miyoshi, N.; Kobayashi, E.; Morita, A.; Tanihata, J.; Takeda, S.; Miura, S. Glycerophospholipid profile alterations are associated with murine muscle-wasting phenotype. Muscle Nerve 2020, 62, 413–418. [Google Scholar] [CrossRef]
  122. Touboul, D.; Piednoel, H.; Voisin, V.; De La Porte, S.; Brunelle, A.; Halgand, F.; Laprevote, O. Changes of phospholipid composition within the dystrophic muscle by matrix-assisted laser desorption/ionization mass spectrometry and mass spectrometry imaging. Eur. J. Mass Spectrom. 2004, 10, 657–664. [Google Scholar] [CrossRef]
  123. Tuazon, M.A.; Henderson, G.C. Fatty acid profile of skeletal muscle phospholipid is altered in mdx mice and is predictive of disease markers. Metabolism 2012, 61, 801–811. [Google Scholar] [CrossRef]
  124. Woodman, S.E.; Park, D.S.; Cohen, A.W.; Cheung, M.W.; Chandra, M.; Shirani, J.; Tang, B.; Jelicks, L.A.; Kitsis, R.N.; Christ, G.J. Caveolin-3 knock-out mice develop a progressive cardiomyopathy and show hyperactivation of the p42/44 MAPK cascade. J. Biol. Chem. 2002, 277, 38988–38997. [Google Scholar] [CrossRef] [Green Version]
  125. Shah, D.S.; Nisr, R.B.; Stretton, C.; Krasteva-Christ, G.; Hundal, H.S. Caveolin-3 deficiency associated with the dystrophy P104L mutation impairs skeletal muscle mitochondrial form and function. J. Cachexia Sarcopenia Muscle 2020, 11, 838–858. [Google Scholar] [CrossRef] [PubMed]
  126. Chen, J.; Zeng, W.; Han, C.; Wu, J.; Zhang, H.; Tong, X. Mutation in the caveolin-3 gene causes asymmetrical distal myopathy. Neuropathology 2016, 36, 485–489. [Google Scholar] [CrossRef]
  127. Pradhan, B.S.; Proszynski, T.J. A Role for Caveolin-3 in the Pathogenesis of Muscular Dystrophies. Int. J. Mol. Sci. 2020, 21, 8736. [Google Scholar] [CrossRef] [PubMed]
  128. Garlid, A.O.; Schaffer, C.T.; Kim, J.; Bhatt, H.; Guevara-Gonzalez, V.; Ping, P. TAZ encodes tafazzin, a transacylase essential for cardiolipin formation and central to the etiology of Barth syndrome. Gene 2020, 726, 144148. [Google Scholar] [CrossRef] [PubMed]
  129. Schlame, M.; Kelley, R.I.; Feigenbaum, A.; Towbin, J.A.; Heerdt, P.M.; Schieble, T.; Wanders, R.J.; DiMauro, S.; Blanck, T.J. Phospholipid abnormalities in children with Barth syndrome. J. Am. Coll. Cardiol. 2003, 42, 1994–1999. [Google Scholar] [CrossRef] [Green Version]
  130. Fajardo, V.A.; Mikhaeil, J.S.; Leveille, C.F.; Saint, C.; LeBlanc, P.J. Cardiolipin content, linoleic acid composition, and tafazzin expression in response to skeletal muscle overload and unload stimuli. Sci. Rep. 2017, 7, 2060. [Google Scholar] [CrossRef] [Green Version]
  131. Ikon, N.; Ryan, R.O. Barth Syndrome: Connecting Cardiolipin to Cardiomyopathy. Lipids 2017, 52, 99–108. [Google Scholar] [CrossRef] [Green Version]
  132. McKenzie, M.; Lazarou, M.; Thorburn, D.R.; Ryan, M.T. Mitochondrial respiratory chain supercomplexes are destabilized in Barth Syndrome patients. J. Mol. Biol. 2006, 361, 462–469. [Google Scholar] [CrossRef]
  133. Wasmus, C.; Dudek, J. Metabolic Alterations Caused by Defective Cardiolipin Remodeling in Inherited Cardiomyopathies. Life 2020, 10, 277. [Google Scholar] [CrossRef]
  134. Bektas, M.; Payne, S.G.; Liu, H.; Goparaju, S.; Milstien, S.; Spiegel, S. A novel acylglycerol kinase that produces lysophosphatidic acid modulates cross talk with EGFR in prostate cancer cells. J. Cell Biol. 2005, 169, 801–811. [Google Scholar] [CrossRef] [Green Version]
  135. Vukotic, M.; Nolte, H.; Konig, T.; Saita, S.; Ananjew, M.; Kruger, M.; Tatsuta, T.; Langer, T. Acylglycerol Kinase Mutated in Sengers Syndrome Is a Subunit of the TIM22 Protein Translocase in Mitochondria. Mol. Cell 2017, 67, 471–483.e7. [Google Scholar] [CrossRef]
  136. Kang, Y.; Stroud, D.A.; Baker, M.J.; De Souza, D.P.; Frazier, A.E.; Liem, M.; Tull, D.; Mathivanan, S.; McConville, M.J.; Thorburn, D.R. Sengers Syndrome-Associated Mitochondrial Acylglycerol Kinase Is a Subunit of the Human TIM22 Protein Import Complex. Mol. Cell 2017, 67, 457–470.e5. [Google Scholar] [CrossRef]
  137. Mitsuhashi, S.; Nishino, I. Phospholipid synthetic defect and mitophagy in muscle disease. Autophagy 2011, 7, 1559–1561. [Google Scholar] [CrossRef]
  138. Marchet, S.; Invernizzi, F.; Blasevich, F.; Bruno, V.; Dusi, S.; Venco, P.; Fiorillo, C.; Baranello, G.; Pallotti, F.; Lamantea, E.; et al. Alteration of mitochondrial membrane inner potential in three Italian patients with megaconial congenital muscular dystrophy carrying new mutations in CHKB gene. Mitochondrion 2019, 47, 24–29. [Google Scholar] [CrossRef]
  139. Muhammad, E.; Reish, O.; Ohno, Y.; Scheetz, T.; Deluca, A.; Searby, C.; Regev, M.; Benyamini, L.; Fellig, Y.; Kihara, A.; et al. Congenital myopathy is caused by mutation of HACD1. Hum. Mol. Genet. 2013, 22, 5229–5236. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  140. Paran, C.W.; Zou, K.; Ferrara, P.J.; Song, H.; Turk, J.; Funai, K. Lipogenesis mitigates dysregulated sarcoplasmic reticulum calcium uptake in muscular dystrophy. Biochim. Et Biophys. Acta 2015, 1851, 1530–1538. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  141. Grounds, M.D.; Terrill, J.R.; Radley-Crabb, H.G.; Robertson, T.; Papadimitriou, J.; Spuler, S.; Shavlakadze, T. Lipid accumulation in dysferlin-deficient muscles. Am. J. Pathol. 2014, 184, 1668–1676. [Google Scholar] [CrossRef] [PubMed]
  142. Haynes, V.R.; Keenan, S.N.; Bayliss, J.; Lloyd, E.M.; Meikle, P.J.; Grounds, M.D.; Watt, M.J. Dysferlin deficiency alters lipid metabolism and remodels the skeletal muscle lipidome in mice. J. Lipid Res. 2019, 60, 1350–1364. [Google Scholar] [CrossRef] [PubMed]
  143. Bohnert, K.R.; McMillan, J.D.; Kumar, A. Emerging roles of ER stress and unfolded protein response pathways in skeletal muscle health and disease. J. Cell. Physiol. 2018, 233, 67–78. [Google Scholar] [CrossRef] [PubMed]
  144. Pitts, M.W.; Hoffmann, P.R. Endoplasmic reticulum-resident selenoproteins as regulators of calcium signaling and homeostasis. Cell Calcium 2018, 70, 76–86. [Google Scholar] [CrossRef] [PubMed]
Ijms 22 08176 g001 550
Figure 1. The great diversity of phospholipids.
Figure 1. The great diversity of phospholipids.
Ijms 22 08176 g001
Ijms 22 08176 g003 550
Figure 3. TIC (total ion chromatogram) by MS ESI-neg profiling and Corona CAD analysis for phospholipid class detection. Standard analysis of 0.3 g/L of PA, PG, CL, PI, PE, PS, LPE, PC, SM, and LPC.
Figure 3. TIC (total ion chromatogram) by MS ESI-neg profiling and Corona CAD analysis for phospholipid class detection. Standard analysis of 0.3 g/L of PA, PG, CL, PI, PE, PS, LPE, PC, SM, and LPC.
Ijms 22 08176 g003
Ijms 22 08176 g004 550
Figure 4. Diacylglycerol (DAG) and CDP-DAG are two key elements in PL biosynthesis. GPAT: glycerol-3-phosphate acyltransferase, LPAAT: lysophosphatidic acid acyltransferases, PAP: phosphatidic acid phosphatases, DGK: diacylglycerol kinase, CDS: CDP-DAG synthetase, DAGAT: diacylglycerol acyl transferase, ALCAT1: AcylCoA:lysoCL acyltransferase 1, MLCAT1: monolysoCL acyltransferase 1, TAZ: tafazzin, EK: ethanolamine kinase, ET: CTP:phosphoethanolamine PE cytidylyltransferase, EPT: ethanolaminephosphotransferase, PSD: phosphatidylserine decarboxylase, PSS2: phosphatidylserine synthase 2, CK: choline kinase, CT: CTP: phosphocholine cytidylyltransferase, CPT: cholinephosphotransferase, PSS1: phosphatidylserine synthase 1, SMS: sphingomyelin synthase.
Figure 4. Diacylglycerol (DAG) and CDP-DAG are two key elements in PL biosynthesis. GPAT: glycerol-3-phosphate acyltransferase, LPAAT: lysophosphatidic acid acyltransferases, PAP: phosphatidic acid phosphatases, DGK: diacylglycerol kinase, CDS: CDP-DAG synthetase, DAGAT: diacylglycerol acyl transferase, ALCAT1: AcylCoA:lysoCL acyltransferase 1, MLCAT1: monolysoCL acyltransferase 1, TAZ: tafazzin, EK: ethanolamine kinase, ET: CTP:phosphoethanolamine PE cytidylyltransferase, EPT: ethanolaminephosphotransferase, PSD: phosphatidylserine decarboxylase, PSS2: phosphatidylserine synthase 2, CK: choline kinase, CT: CTP: phosphocholine cytidylyltransferase, CPT: cholinephosphotransferase, PSS1: phosphatidylserine synthase 1, SMS: sphingomyelin synthase.
Ijms 22 08176 g004
Table
Table 1. The importance of the double bond position.
Table 1. The importance of the double bond position.
Fatty Acid18:118:218:3
Mass and
Formula		282.26 and C18H34O2280.24 and C18H32O2278.22 and C18H30O2
Systemic NameOctadecenoic acidOctadecadienoic acidOctadecatrienoic acid
Number of
Lipid Records		4310163
Common NameDouble Bond
Position		Number of
PubMed
References		Common NameDouble Bond
Position		Number of PubMed
References		Common NameDouble Bond
Position		Number of
PubMed
References
ExampleOleic acid9Z23,694Linoleic acid9Z, 12Z24,310α-Linolenic acid9Z, 12Z, 15Z7036
Trans-
vaccenic acid		11E325Rumenic acid9Z, 11E316γ-Linolenic acid6Z, 9Z, 12Z3145
Cis-vaccenic acid11Z202Linoelaidic acid9E, 12E25Punicic acid9Z, 11E, 13Z111
Petroselinic acid6Z67Sebaleic acid5Z, 8Z9Pinolenic acid5Z, 9Z, 12Z56
Petroselaidic acid6E4Vaccelenic acid11E, 15Z2Jacaric acid8Z, 10E, 12Z13
Formula of the fatty acid can be deduced from its systematic name. Records lipid from https://lipidmaps.org/data/chemdb_lm_text_ontology.php?ABBREV=FA%2018:3 (accessed on 3 May 2021) and records from PubMed https://pubmed.ncbi.nlm.nih.gov/ (accessed on 3 May 2021). (E: trans double bond, Z: cis double bond).
Table
Table 3. PL alterations involved in muscular diseases.
Table 3. PL alterations involved in muscular diseases.
Gene
Mutated		Protein and FunctionDiseaseMuscular
Symptoms		Lipid DisordersAnimal Experimental ModelsRef.
ORPHA 352312: group of disorders of PL, sphingolipids and
FA biosynthesis with skeletal muscle characteristics		TAZ
chromosome X		Tafazzin
Phospholipid-lysophospholipid transacylase
Catalyzes remodeling of immature CL to its mature composition		Barth syndromeDilated cardiomyopathy
Skeletal muscle weakness		Reduced level of CL
Increased level of mono-lysocardiolipin
  • Taz−/− (knock-out) mice:
    most die before birth due to skeletal muscle weakness, survivors develop progressive cardiomyopathy
  • Taz inducible knock-down mice (doxycycline-inducible short hairpin RNA):
    impact on cognitive abilities, brain mitochondrial respiration and the function of hippocampal neurons and glia
  • Tafazzin mutant Drosophila melanogaster:
    motor weakness (reduced flying and climbing abilities)
[71,72,73]
AGK
chromosome 7		AGK
Acylglycerol kinase
Phosphorylates MAG and DAG to form LPA and PA		Sengers syndromeHypertrophic cardiomyopathy
Skeletal myopathy
Congenital cataract		Defects in mitochondria and storage of lipids in muscle (skeletal and heart)
  • Agk−/− mice:
    phenotype similar to Sengers syndrome, develop thrombocytopenia and splenomegaly
[74,75,76,77]
LPIN 1
chromosome 2		Lipin-1
Phosphatidate phosphatase
Catalyzes the conversion of PA to DAG		MyoglobinuriaRhabdomyolysis
Muscle fiber loss		Hyper-triglyceridemia
Accumulation of PL intermediates
  • Spontaneous mutant mice ‘fld’ (fatty liver dystrophy):
    neonatal mice: hypertriglyceridemia andenlarged, triglyceride-rich fatty liver
    after 2 weeks: neuropathy with abnormal myelin formation and hind limb weakness
  • Transgenic mice models with overexpression of Lpin1:
    muscle specific overexpression causes increased body weight
  • Lpin1 floxed/floxed:
    when deleted in muscle: myopathy
[78,79,80,81]
PNPLA2
chromosome 11		ATGL
Adipose triglyceride lipase
Catalyses the hydrolysis of TAG into DAG		Slowly progressive myopathy: neutral lipid storage disease with myopathy (NLSDM)
Arrhythmogenic cardiomyopathy		Distal muscle weakness
Cardiomyopathy		Accumulation of lipid droplets(TAG) in multiple tissues
Altered energy metabolism
Lipidosis of internal organs
  • Atgl−/− mice:
    premature mortality from lipid cardiomyopathy by 16 weeks
    homozygous knockin mouse model: arrhythmias and significant cardiac dysfunction at 12 weeks, sudden death and/or heart failure by 14 weeks
[82,83,84,85]
CHKB
chromosome 22		CHKB
Choline kinase β
Catalyzes the first step in PE biosynthesis		Megaconial congenital muscular dystrophyMitochondrial structural (enlargement) and functional abnormalitiesDecreased level of PC
  • Rostrocaudal muscular dystrophy mice (rmd):
    spontaneous autosomal recessive trait that causes rapidly progressive muscular dystrophy and neonatal forelimb bone deformity
[86,87,88,89]
Disease not listed in ORPHA 352312 despite gene mutation in gene involved in PL biosynthesis, sphingolipids, and FA synthesisHACD1
chromosome 10		HACD1
3-Hydroxyacyl-CoA dehydratase 1
Catalyzes the third of the four reactions of the long-chain FA elongation cycle		Congenital fiber-type disproportion, rare internalized nucleiCongenital myopathy with proximal ± distal muscle weakness, progressively improving over timeIncreased concentrations of ≥ C18 and mono-unsaturated FA
  • Spontaneous Hacd1 pathogenic variant Labrador retriever dogs:
    congenital centronuclear myopathy with fiber size disproportion associated with generalized and progressive muscle weakness
  • Hacd1−/− mice:
    postural and locomotor weakness and reduced weight gain
[90,91,92]
ACADVL
chromosome 17 		VLCAD
Very long chain Acyl-coA dehydrogenase
Involved in the β-oxidation pathway (FA longer than C16)		VLCADD (VLCAD deficiency)Cardiomyopathy
Myopathy with intermittent myalgia and rhabdomyolysis, progressive muscle weakness, often dropped head		Defective long-chain FA oxidation
  • Vlcad−/− mice:
    Cardiac and metabolic dysfunction
    Different phenotypes between male and female: only KO female mice develop a severe clinical phenotype upon medium chain triglycerides supplementation
[93,94,95,96,97]
ACADM
chromosome 1		MCAD
Medium-chain acyl-coenzyme A dehydrogenase
Involved in the β-oxidation pathway (C4 to C12 acyl chain)		Medium-chain acyl-CoA dehydrogenase deficiency (MCADD)Myopathy exertion induced myalgia, progressive proximal limb weakness
Secondary carnitine deficiency		Accumulation of FA intermediates
  • Mcad−/− mice:
    organic aciduria and fatty liver, sporadic cardiac lesions, neonatal mortality
[98,99,100]
HADHA
chromosome 2		MTP
Mitochondrial trifunctional protein
Catalyzes three out of the four steps in beta oxidation		Mitochondrial trifunctional protein deficiency (MTP deficiency or MTPD)Skeletal myopathy, episodic rhabdomyolysis, peripheral neuropathy, cardiomyopathy in severe casesDecreased long-chain FA oxidation
  • Mtpa−/− mice:
    severe phenotype (fetal growth retardation, neonatal hypoglycemia) and early sudden death (necrosis and acute degeneration of cardiac and diaphragmatic myocytes)
[101,102,103]
CGI-58
chromosome 3		ABHD5
Alpha/beta hydrolase domain containing 5
Lipid droplet-associated protein that activates ATGL		Chanarin–Dorfman syndrome
(neutral lipid storage disease)		Moderate myopathy Possible cardiomyopathyAccumulation of TAG droplets
  • Cgi-58 −/− mice:
    various phenotypes including cardiomyopathy, severe hepatic steatosis, skin defects
[104,105,106,107,108]
CPT2
chromosome 1		CPT2
Carnitine palmitoyltransferase 2
Catalyzes the transfer of FA from cytoplasm to mitochondria		Carnitine palmitoyl-transferase II deficiencyEpisodic myalgia, muscle weakness, and rhabdomyolysis with myoglobinuriaReduced the rates of long-chain FA oxidation into carbon dioxide
  • Cpt2floxed/floxed mice:
    cardiac and muscle CPT2 deficiency leads to severe cardiac hypertrophy and ultimately heart failure, with no overt macroscopic muscle phenotype
[109,110,111,112]
Alteration of PL in inherited form of myopathies due to
mutation in genes without a direct link to PL biosynthesis		DYSF
chromosome 2		Dysferlin
Involved in the sarcolemma repair mechanism, interacts with caveolin 3		Dysferlinopathy, including:
limb girdle muscular dystrophy type 2B (LGMD2B)
Miyoshi myopathy		Proximal, distal, or proximo-distal myopathy, predominantly involving lower limbs
Rhabdomyolysis		Early intra-myocellular lipid accumulation (sphingolipids, PLs, cholesterol)
  • various spontaneous murine dysferlin mutants (SJL/J, A/J, BLAJ) with progressive muscular dystrophy
[113,114,115,116]
DMD
chromosome X		Dystrophin
Involved in the connection of the cytoskeleton of a muscle fiber to the extracellular matrix through the cell membrane		Duchenne muscular dystrophy
Becker muscular dystrophy		Cardiomyopathy with
myocardial fibrosis,
muscle weakness and muscle fiber necrosis,
respiratory failure		PL dyshomeostasis
Increased PC and SM
  • Spontaneous pathogenic variant in mice: mdx model
  • Spontaneous pathogenic variant in dog: golden retriever muscular dystrophy
  • Spontaneous pathogenic variant in cat: hypertrophic feline muscular dystrophy
  • Zebrafish model
  • Drosophila model
  • Caenorhabditis model
  • Newly developed rat and pig DMD models
[117,118,119,120,121,122,123]
CAV3
chromosome 3		Caveolin3
Component of caveolae plasma membrane (possible scaffolding role for caveolin-interacting molecules)		Rippling muscle disease
LGMD
Isolated familial hypertrophic cardiomyopathy
HyperCKemia and myalgia		Skeletal muscle weakness
Muscle hyperirritability triggered by stretch		Decreased CL
  • Cav3−/− mice:
    muscular dystrophy phenotype and alteration of the phenotypic behavior of cardiac myocytes
    Altered glucose metabolism and increased myoblast proliferation in muscle cells
[124,125,126,127]
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Bargui, R.; Solgadi, A.; Prost, B.; Chester, M.; Ferreiro, A.; Piquereau, J.; Moulin, M. Phospholipids: Identification and Implication in Muscle Pathophysiology. Int. J. Mol. Sci. 2021, 22, 8176. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms22158176

AMA Style

Bargui R, Solgadi A, Prost B, Chester M, Ferreiro A, Piquereau J, Moulin M. Phospholipids: Identification and Implication in Muscle Pathophysiology. International Journal of Molecular Sciences. 2021; 22(15):8176. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms22158176

Chicago/Turabian Style

Bargui, Rezlène, Audrey Solgadi, Bastien Prost, Mélanie Chester, Ana Ferreiro, Jérôme Piquereau, and Maryline Moulin. 2021. "Phospholipids: Identification and Implication in Muscle Pathophysiology" International Journal of Molecular Sciences 22, no. 15: 8176. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms22158176

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop