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Review

The Swing of Lipids at Peroxisomes and Endolysosomes in T Cell Activation

1
Servicio de Inmunología, Hospital Universitario de la Princesa, Universidad Autónoma de Madrid (UAM), Instituto de Investigación Sanitaria Princesa (IIS-IP), 28006 Madrid, Spain
2
Centro Nacional de Investigaciones Cardiovasculares (CNIC), 28029 Madrid, Spain
3
Centro de Investigación Biomédica en Red de Enfermedades Cardiovasculares (CIBERCV), 28029 Madrid, Spain
*
Authors to whom correspondence should be addressed.
These authors contributed equally to this work.
Int. J. Mol. Sci. 2020, 21(8), 2859; https://0-doi-org.brum.beds.ac.uk/10.3390/ijms21082859
Submission received: 26 February 2020 / Revised: 15 April 2020 / Accepted: 16 April 2020 / Published: 19 April 2020
(This article belongs to the Special Issue Signaling and Organelle Polarization at the Immunological Synapse)

Abstract

:
The immune synapse (IS) is a well-known intercellular communication platform, organized at the interphase between the antigen presenting cell (APC) and the T cell. After T cell receptor (TCR) stimulation, signaling from plasma membrane proteins and lipids is amplified by molecules and downstream pathways for full synapse formation and maintenance. This secondary signaling event relies on intracellular reorganization at the IS, involving the cytoskeleton and components of the secretory/recycling machinery, such as the Golgi apparatus and the endolysosomal system (ELS). T cell activation triggers a metabolic reprogramming that involves the synthesis of lipids, which act as signaling mediators, and an increase of mitochondrial activity. Then, this mitochondrial activity results in elevated reactive oxygen species (ROS) production that may lead to cytotoxicity. The regulation of ROS levels requires the concerted action of mitochondria and peroxisomes. In this review, we analyze this reprogramming and the signaling implications of endolysosomal, mitochondrial, peroxisomal, and lipidic systems in T cell activation.

1. Introduction

Cellular adaptive immunity requires the interaction of antigen presenting cells (APC) with T cells bearing T cell receptors (TCR), which need to be specific for the antigen: major histocompatibility complex (MHC) combination presented by the APC. The intimate contact between the APC and the T cell has come to be known as the immune synapse (IS) [1,2]. The IS is a dynamic, highly organized macromolecular structure initiated by antigen recognition through the TCR. It is worth mentioning that this structure is also formed in other immune events, i.e., between natural killer and infected/tumor cells allowing their cytolytic activity or between macrophages or mast and T cells triggering the inflammatory response. Among other functions, the IS promotes the bidirectional communication between the T cell and the APC [1,3,4,5]. During antigen presentation, the T lymphocyte relocates receptors, cytoskeletal components, and specific organelles to the contact site with the APC. These drive actin polymerization and reorganization, forming a contractile actomyosin ring that shapes the IS [6,7]. These signals also drive de novo polymerization of microtubules (MTs) from the centrosome, which is polarized towards the T cell:APC interface [8]. Once this reorganization is achieved, the cytoskeleton guides the redistribution of the endolysosomal system (ELS) towards the IS, facilitating the subsequent polarized secretion and endocytic recycling of receptors at the T cell-APC contact site [9,10]. Mitochondria also relocate to the IS, where they play a role in the maintenance of Ca2+ fluxes along the T cell activation cycle [11,12,13,14].
To establish a correct communication between the IS-engaged cells, the information travels back and forth between the APC and the T cell. Much attention has been devoted to the activating effects of the APC on the T cell [15]. On the other hand, the engaged T cell also influences APC function using soluble factors (e.g., cytokines and chemokines), exosomes, and other types of secretory vesicles that are released during the APC-T cell contact [16,17,18,19,20]. This crosstalk favors the functional modulation of the APC and T cells that form the IS and, eventually, the optimal priming, activation, and T cell responses against cognate antigens.
Upon IS formation, mitochondria and peroxisomes are major organelles that mediate lipid metabolism changes and, in turn, T cell activation signaling pathways. Mitochondria polarize to the immune synapse and act as major regulators of lipid metabolism, reactive oxygen species (ROS) production and calcium fluxes. In addition, peroxisomes act as collaborators of mitochondria in the regulation of both lipid and ROS levels. The ELS is a dynamic cell compartment with several organelles and the plasma membrane that mediates vesicle and exosome delivery between the APC and the T cell. It comprises two different compartments: early and late endosomes. Late endosomes form intraluminal vesicles (ILVs) by the invagination of their limiting membrane [21], becoming the so-called multivesicular bodies (MVB). These ILV-enriched organelles are directly involved in sorting and recycling of proteins from the plasma membrane to intracellular compartments and vice versa [22]. Furthermore, they are directly related to exosome formation and their sorting towards the APC [23,24,25]. As a result of these transport events the plasma membrane has a heterogeneous profile composed of co-existing domains enriched in different proteins and lipids. These may serve to regulate cell-to-cell communication and activation [26]. T cell activation, in addition, is accompanied by a metabolic reprogramming. This process requires high ATP consumption that results in an increase of lipid and ROS levels. These changes in the T cell modulate plasma membrane composition, protein signaling pathways and cytoskeleton organization. These effects are accompanied by changes in metabolic processes that, in turn, regulate ROS levels. In this review we offer to the reader an overview of the changes in intracellular organelles and lipid metabolism associated to T cell activation through the IS.

2. Endolysosomal System and Multivesicular Bodies

The endolysosomal system is the main regulator of metabolic homeostasis. It allows the recycling of previously synthesized molecules. Endosomes are identified by the presence of Rab GTPase proteins. The ELS uses MVB morphogenetic pathways to reach the lumen of lysosomes, where intracellular and extracellular cargoes are degraded. This process acts in concert with autophagy, in which damaged organelles and aggregated proteins are recycled through lysosomal pathways. Autophagy is induced in IS-forming dendritic cells through ATG16L; it regulates mTORC activity, acting as a negative feedback modulator by destabilizing the synapse. Specific mutations of this protein have been described to lead to hyperstable T cell:APC interactions and possibly contributing to T cell hyperactivity in Crohn’s disease patients [27]. On the T cell side, the intraflagellar transport system (IFT) regulates the biogenesis of lysosomes [28] and the retrograde transport towards intracellular endosomes upon immune synapse formation, favoring CD3 and LAT recruitment and recycling [29]. In addition, there are other non-vesicular and vesicular mechanisms that target selective cargoes to lysosomes, such as chaperone-mediated autophagy and microautophagy [30,31].
Recycling and delivery of molecules at the IS interface is important for a balanced downstream signaling and full T cell activation. For example, both the TCR/CD3 complex and the downstream scaffold protein LAT (Linker for Activation of T Cells) are subjected to a tight regulation along this process [16,17]. Vesicle traffic constitutes a basic component of this machinery, and cytoskeletal components modulate its dynamics [18,19,20]. MVB relocate to the IS and help the establishment of the actin-rich ring through clathrin and associated components, such as Hrs [32]. Hrs (Hepatocyte growth factor-regulated tyrosine kinase substrate) is an endosome-associated regulator for vesicular transport and protein sorting [33].
This process enables the integrin-based adhesive part of the IS (Peripheral Supramolecular Activation Clusters, pSMAC) to seal the intercellular space between the T cell and the APC [1,8]. LBPA (lysobisphosphatidic acid) represents about 15 % of total phospholipids in MVB and is only detected at the IS, which strongly argues for a localized synthesis from a phospholipidic precursor at this location [34]. LBPA appears late in the IS, specifically after MVB fusion [11], which provides the plasma membrane with specific lipids that may regulate other processes. The fusion of MVB at the IS allows the release of their content, including ILVs as exosomes. Exosome membranes are enriched in cholesterol, sphingolipids such as sphingomyelin (SM) and GPI-anchored proteins, as well as tetraspanins and LAMPs (lysosome-associated membrane proteins) [35,36,37,38]. The delivery of exosomes from MVB at the IS interface allows their capture by the APC, which causes changes in the recipient cell [25,39,40,41,42].
Extracellular vesicle biogenesis follows two main routes. One involves the formation of endosomal patches found at the plasma membrane and the fusion of endosome membranes with the plasma membrane [43]. The second is slower because it requires the invagination of endosomal membranes to form the ILVs and their subsequent release through exocytosis. Protein sorting to ILVs is mediated by the ESCRT (endosomal sorting complexes required for transport) machinery after its interaction with K63-ubiquitinated proteins [44,45]. Different components of the ESCRT machinery, such as ESCRT-0 and ESCRT-1 (TSG101), contribute to MHC-II sorting to exosomes and biogenesis of membrane-derived exosomes [46]. On the other hand, ESCRT-III seems to play a negative role, and its inactivation increases ILV production [47]. The cell content of MVB increases with T cell activation. This increase is regulated by diacylglicerol kinase α (DGKα) through a dual role. On one hand, DGKα produces phosphatidic acid from diacylglicerol (DAG), which negatively regulates T cell activation; depletion of this enzyme prevents CD28-dependent T cell anergy [48]. On the other hand, DGKα translocates to MVB upon TCR activation and its inhibition increases mature MVB and exosome secretion [49].

3. Mitochondria and Peroxisomes: Modulators of the Immune Synapse

During differentiation, T cells increase their pool of mitochondria to pay the energetic cost of this process. In addition, these organelles are also involved in lipid synthesis, calcium homeostasis, apoptosis, signaling, and cell progression [50,51]. These processes place mitochondria as important organelles during immune synapse formation and T cell activation. After antigen encounter and receptor stimulation, mitochondria are polarized to the IS [14,52,53]. This is related to the regulation of calcium fluxes and to the production of the energy required for activation. As we describe below, mitochondrial metabolism is critical for T cell activation through production of mitochondrial ROS (mROS) [54,55,56].
Peroxisomes, in collaboration with mitochondria, prevent cell toxicity through ROS elimination. Besides oxidative modulators, these organelles are sites for Fatty Acid β-Oxidation (FAO). They also promote D-amino acid degradation, polyamine oxidation, and synthesis of various cholesterol precursors and plasmalogens [57]. Peroxisome proliferation and degradation are regulated processes and, in contrast to mitochondria, these organelles are unable to fuse with each other. There are two types of peroxisome biogenesis processes: growth and division of pre-existing organelles [58], and de novo formation from the endoplasmic reticulum (ER) [59]. Degradation of these organelles occurs by micro- and macro-autophagy or by 15-lipoxygenase-mediated autolysis [60]. The association between peroxisomes and T cell maturation has remained elusive, although some evidence links PPAR (Peroxisome Proliferator-Activated nuclear Receptor) to T cell development. PPARs are a family of ligand-activated transcription factors that, when dysregulated, affect a variety of physiological processes (lipid metabolism, cellular differentiation, and cancer). For example, alteration of cholesterol metabolism can affect TCR signaling or FAO [61]. In addition, PPARβ overexpression is related to inhibition of peripheral αβ T cell proliferation, while the γδ T cell population remains unaffected [62]. Thus, mitochondria and peroxisomes are key players in T cell activation. Calcium fluxes, ATP production, and ROS levels are processes directly modulated by these organelles, as illustrated in Figure 1.

4. Reactive Oxygen Species as Modulators of T Cell Activation

ROS are chemically reactive free radicals with one unpaired electron in their outer orbit. The most familiar ones are superoxide, hydrogen peroxide, hydroxyl radical, and singlet oxygen. These free radicals are generated as the result of a partial reduction of oxygen, a process that occurs mainly in mitochondria and peroxisomes. This output is directly related with the oxidative and fatty acid metabolism and there is an active interplay between both organelles [63,64]. In fact, they follow common steps in their course, such as dehydrogenation, hydration, and thiolytic cleavage for FAO [65]. In addition, mitochondria and peroxisomes have an equilibrium between oxidant and antioxidant agents to avoid an oxidative stress that could ultimately alter their function. A misregulation of this balance could induce an uncontrolled proliferation, eventually leading to cancer [57,63,64,66,67,68].
ROS have always been related to harmful by-products that cause DNA damage, genomic instability, or protein dysfunction, leading to cell senescence or cell death caused by apoptosis [69,70,71]. However, in the last few years this view has changed. Variations in the oxidative components during different cell processes are able to regulate cell metabolism. In this sense, low to moderate ROS levels contribute to signaling pathways implicated in cell growth, death, and migration [72]. For example, phosphatases, kinases, and transcription factors can be modulated by oxidative transitions. In addition, ROS-related modifications can alter the function of some proteins by modulating their associated signaling pathways [73,74]. T cells are not an exception and variations in ROS levels have a role during their activation and differentiation stages. The main ATP sources of circulating naive T cells (Tn) are oxidative phosphorylation (OXPHOS) and FAO [75,76]. T cell activation causes an abrupt increment of ATP consumption leading to metabolic reprogramming. The catabolic state gives rise to an anabolic metabolism that provides the energy necessary to proliferate, differentiate and become effector T cells (Teff). This is characterized by increased nutrient uptake causing, in consequence, a gain in glycolysis products. This shift activates signaling pathways, transcription factors and effector molecules [77,78,79,80,81]. Additionally, TCR ligation increases the production of ROS by OXPHOS and NADPH oxidases (NOXs), a family of plasma membrane-associated oxidases that mediate proliferation and differentiation [55,82,83]. This is complemented with the crucial role of mROS, which activate members of the NFAT family of transcription factors and Myc [54]. Additionally, they also activate signaling molecules such as mTOR, NF-kB, and AP-1. These effects directly relate mROS with an IL-2 dependent proliferative status and with a metabolic reprogramming of T effector cells [55,56]. After antigen clearance, only a small proportion of T cells survive, becoming memory T cells (Tm). These cells remain primed for a future encounter with the same antigen [84,85]. Tm do not have the same energy demand as Teff, losing those glycolytic requirements and recovering the metabolic profile of Tn. To avoid apoptosis and cell death, this is accompanied by an increment in mitochondrial FAO and the number of mitochondria [76]. In addition, low amounts of metabolic ROS promote increased lifespan and response capacity of these Tm [86].

5. Lipids as a Source of Energy in Metabolic Reprogramming after T Cell Activation

As mentioned above, when T cells are activated, they undergo metabolic reprogramming, acquiring a biosynthetic status. There is a switch from pyruvate oxidation via the tri-carboxylic acid cycle and FAO to aerobic glycolysis, together with induction of the pentose-phosphate pathway, glutaminolysis and de novo fatty acids synthesis (FAS). This is controlled by changes in the metabolic transcriptome and the subsequent induction of the transcription factors Myc and HIF1α [54]. The metabolites resulting from the induction of those pathways are used as the nitrogen and carbon sources necessary for a range of biosynthetic precursors of lipids and polyamines. The reprogramming of activated T cells includes also a marked downregulation of mitochondria-dependent FAO, with a decrease in the levels of products such as carnitine [54]. Another regulator playing an important role in de novo FAS induction is the mTOR1 complex (mTORC1) [87]. Some studies demonstrate that the diminished function of mTORC1 due to the lack of its scaffolding protein Raptor causes an impairment in de novo TCR induced lipid biosynthesis [88]. MTORC1 acts in conjunction with Sterol Regulatory Element Binding Proteins (SREBPs), which are a group of transcription factors acting as essential regulators of cholesterol synthesis. Thus, the mTORC-SREBP pathway activates the lipid biosynthetic program [88].
Although T cells obtain their energy preferably from glucose, the oxidation of fatty acids is also an important source of ATP. FAO occurs in the mitochondrial matrix and requires the entry of the fatty acid attached to carnitine through the mitochondrial membrane with the contribution of carnitine-palmitoyl-CoA transferase-1 (CPT1) [89,90]. Fatty acid entry into mitochondria is regulated by a feedback system: when acetyl-CoA carboxylase is inactivated by an AMPK-induced phosphorylation, the concentration of its product malonyl-CoA decreases, facilitating the activity of CPT1 [91,92]. This process is especially relevant for the development and maintenance of regulatory T cells (Treg) and Tissue-Resident Memory T cells (TRM) [93,94].

6. Lipid second Messengers at the Immune Synapse (IS)

Phosphoinositide molecules have a key function in TCR downstream signaling. Phosphatidylinositol-4,5-bisphosphate (PIP2) plays structural and regulatory roles at the IS, both directly and indirectly through its derivatives. As a result of TCR/CD3 triggering, LAT recruits phospholipase C (PLC) and contributes to its activation. PLC converts PIP2 into inositol-1, 4, 5-trisphosphate (IP3) and diacylglicerol (DAG). PIP2 production at the IS is necessary to replenish the plasma membrane stores, since it represents less than 1% of membrane lipids. Sustained PIP2 turnover at the IS depends on CD28 co-stimulation in a PI5PKI-dependent manner and prevents T cell anergy [95,96,97]. PIP2 activates phospholipase D (PLD), which produces phosphatidic acid and activates PI5PKI, creating a positive feedback loop [98]. PIP2 is also a regulator of actin cytoskeleton assembly and the processes of endocytosis and exocytosis. ERM proteins (ezrin, radixin, and moesin) are found in vesicles. They adopt an open/active conformation upon binding to PIP2, facilitating the binding of a plethora of plasma membrane proteins—including adhesion and signaling receptors and tetraspanin scaffold proteins—to the actin cytoskeleton [99,100]. IP3 allows a transient opening of ER Ca2+ channels. The release of this secondary messenger to the cytosol regulates cell signaling, cytoskeletal reorganization, vesicular traffic, and secretion [8]. In contrast, DAG remains in the plasma membrane, preferentially accumulated at the cSMAC (Central Supramolecular Activation Clusters). Additionally, it is involved in the reorganization of the tubulin cytoskeleton through the control of centrosome polarity [101]. Finally, the joint action of DAG and the released Ca2+, allows the recruitment of protein kinase C (PKC) to the plasma membrane, its activation and, in turn, the beginning of signaling cascades [102].
Fatty acids also play a regulatory role in T cell activation as post-translational labels. For example, palmitoylation or myristoylation of several proteins of TCR downstream signaling leads to their recruitment to the IS. This enables the propagation of the signal inside the cell [103,104,105]. Furthermore, prenylation consists in the covalent attachment of a farnesyl or a geranylgeranyl isoprenoid group, which allows higher hydrophobicity and interaction with target membranes. Small GTPases including Ras, Rac, Rab, and Rho—with roles in signal transduction, cytoskeletal regulation and intracellular vesicle trafficking—undergo prenylation [106]. Acetylation is another post-translational modification important in signaling in both resting and activated T cells [107]. It derives from the key lipid metabolite acetyl-CoA. Acetyl groups modify histones and trigger epigenetic mechanisms which can alter the communication of the T cell with other immune cells, by regulating cytokine gene expression [108]. These changes are related to several chronic diseases [109].

7. Bioactive Sphingolipids and T Cell Activation

Sphingolipids (SL) are one of the most abundant components of the plasma membrane [110]. Bioactive sphingolipids play a role in the regulation of TCR signal transduction and protein sorting during T cell activation. [111]. This type of SL forms membrane domains together with sterols. These domains, together with compartmentalized membrane proteins and their proximal membrane associated signaling components, constitute the so-called lipid microdomains or lipid rafts [112]. The group of bioactive sphingolipids is composed of ceramide (Cer), sphingosine (Sph), ceramide-1-phosphate (C1P), and sphingosine-1-phosphate (S1P) [111]. The dynamics of bioactive sphingolipids and their implication in T cell activation are depicted in Figure 2.
Ceramide has a central function in the metabolism of bioactive sphingolipids; it is the substrate of a variety of enzymes that convert this lipid in the rest of bioactive SLs or other important lipids of the SL metabolism, as SM or glycosphingolipids [111,113]. SM is a key player in T cell activation and an important component of both the plasma membrane and lipid droplets [114]. SM is hydrolyzed at the membrane of diverse cellular components such as lysosomes by the corresponding isoform of the enzyme sphingomyelinase (neutral (nSMase) or acid (aSMase)). This reaction generates the products phosphocholine and Cer [113]. Then, the subsequent release of Cer within the T cell membrane leads to the formation of ceramide-enriched membrane microdomains at the IS [112]. Due to their biophysical properties, these structures compartmentalize receptors and their proximal signaling proteins, regulating TCR signaling. The maintenance of TCR signaling through CD3ζ and ZAP70 polarization to the IS requires nSM2. Furthermore, nSMase plays a role in the dynamics of polarization and stabilization of the microtubular system and the microtubule-organizing center. This is mediated by the recruitment of PKCζ and its downstream substrate Cdc42, which acts as organizer of cell polarity during IS maintenance [114]. Cer released from SM by SMases is used to synthetize de novo the rest of bioactive SL at different regions and compartments. It can also be recirculated to the cell membrane via endosomal pathway [111]. In relation to vesicular transport, there is experimental evidence supporting that nSMase2 triggers the budding of exosomes into MVB in T cells [39,115].
S1P and C1P are formed by the enzymatic action of kinases, which phosphorylate sphingosine and ceramide, respectively. S1P regulates lymphocyte egress into circulation through interaction with its receptor S1PR. S1PR association with CD69 [116,117] downregulates its membrane expression, allowing prolonged lymphocyte retention in inflamed tissues [118]. Due to its involvement in cell trafficking, S1P mediates cancer cell growth, proliferation, survival, and is related to inflammatory and autoimmune diseases such as asthma, atherosclerosis, Crohn’s disease, diabetes, and osteoporosis [119,120,121,122]. In addition, this bioactive SL is able to bind to transcription factors as HDAC1/2 or PPARɣ and modulates T cell metabolism to help differentiation, changing T cell capability of anti-tumor response [119]. C1P, in turn, increases the intracellular Ca2+ concentration in TCR-activated cells, as ceramide does. In contrast to Cer, C1P acts by promoting IP3 production, thereby inducing Ca2+ release from the ER and the opening of store-operated calcium channels (SOCC) at the plasma membrane [123]. Furthermore, C1P is tightly related with eicosanoid biosynthesis during immune response, activating initial rate-limiting enzymes in eicosanoid biosynthesis and inducing arachidonic acid release in the early stages of wound healing [124,125,126].

8. Concluding Remarks

The organization and composition of the cell membrane is a crucial factor during T cell activation. The initial signaling cascade, as well as the cytoskeletal and intracellular traffic, produce the quick polarization of organelles to the IS and the rapid shift in protein and lipid content. Mitochondria, peroxisomes, and the endolysosomal system are key players in lipid and protein homeostasis. In addition, they regulate both vesicle traffic and exosome production and their cargo content. Lipids act as mediators of signaling events, scaffolds for proteins in a structural and/or dynamic way, and as energy suppliers. When energy demands increase, the glycolytic pathway is promoted, causing an ATP increment associated to metabolic reprogramming. At this point, lipids are not further required as energy providers and become players in IS signaling. These molecules act as important modulators altering plasma membrane composition, endolysosomal biogenesis and dynamics, and regulating ROS levels. These roles place lipids as important regulators of the immune synapse and, in turn, of T cell activation. In fact, alterations in lipid homeostasis are related to several diseases of the immune system. Altogether, this information emphasizes the importance of understanding the normal and pathological functions of lipids in relation to T cell signaling.

Author Contributions

Writing—original draft preparation, S.G.D., A.R.-G., N.B.M.-C.; writing—review and editing, S.G.D., A.R.-G., N.B.M.-C.; art graph composition, S.G.D., A.R.-G.; supervision and editing, F.S.-M.; project administration, F.S.-M.; funding acquisition, F.S.-M. All authors have read and agreed to the published version of the manuscript.

Funding

This review was funded by grant SAF2017-82886-R from the Spanish Ministry of Economy and Competitiveness (MINECO), grant S2017/BMD-3671-INFLAMUNE-CM from the Comunidad de Madrid, a grant from the Ramón Areces Foundation “Ciencias de la Vida y la Salud” (CIVP19A5941 XIX Concurso-2018) and a grant from Ayudas Fundación BBVA a Equipos de Investigación Científica (BIOMEDICINA-2018), the Fundació Marató TV3 (grant 122/C/2015) and “La Caixa” Banking Foundation (HR17-00016). BIOIMID (PIE13/041) from Instituto de Salud Carlos III, CIBER Cardiovascular (CB16/11/00272, Fondo de Investigación Sanitaria del Instituto de Salud Carlos III and co-funding by Fondo Europeo de Desarrollo Regional FEDER). The Centro Nacional de Investigaciones Cardiovasculares (CNIC) is supported by the Spanish Ministry of Economy and Competitiveness (MINECO) and the Pro-CNIC Foundation and is a Severo Ochoa Center of Excellence (MINECO award SEV-2015-0505). SGD and ARG are funded by fellowship FPU and FPI programs, from Ministry of Science and Universities, respectively. Authors declare no competing interest. Funding agencies do not have intervened in the design of the studies, with no copyright over the study.

Acknowledgments

Authors thank Manuel Gomez, Miguel Vicente-Manzanares and Danay Cibrian for critical review and editing.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Monks, C.R.; Freiberg, B.A.; Kupfer, H.; Sciaky, N.; Kupfer, A. Three-dimensional segregation of supramolecular activation clusters in T cells. Nature 1998, 395, 82–86. [Google Scholar] [CrossRef] [PubMed]
  2. Fooksman, D.R.; Vardhana, S.; Vasiliver-Shamis, G.; Liese, J.; Blair, D.A.; Waite, J.; Sacristan, C.; Victoria, G.D.; Zanin-Zhorov, A.; Dustin, M.L. Functional anatomy of T cell activation and synapse formation. Annu. Rev. Immunol. 2010, 28, 79–105. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. Dieckmann, N.M.; Frazer, G.L.; Asano, Y.; Stinchcombe, J.C.; Griffiths, G.M. The cytotoxic T lymphocyte immune synapse at a glance. J. Cell Sci. 2016, 129, 2881–2886. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Wülfing, C.; Davis, M.M. A receptor/cytoskeletal movement triggered by costimulation during T cell activation. Science 1998, 282, 2266–2269. [Google Scholar] [CrossRef] [Green Version]
  5. Grakoui, A.; Bromley, S.K.; Sumen, C.; Davis, M.M.; Shaw, A.S.; Allen, P.M.; Dustin, M.L. The immunological synapse: A molecular machine controlling T cell activation. Science 1999, 285, 221–227. [Google Scholar] [CrossRef] [Green Version]
  6. Gomez, T.S.; Billadeau, D.D. T cell activation and the cytoskeleton: You can’t have one without the other. Adv. Immunol. 2008, 97, 1–64. [Google Scholar] [CrossRef]
  7. Ilani, T.; Vasiliver-Shamis, G.; Vardhana, S.; Bretscher, A.; Dustin, M.L. T cell antigen receptor signaling and immunological synapse stability require myosin IIA. Nat. Immunol. 2009, 10, 531–539. [Google Scholar] [CrossRef] [Green Version]
  8. Martin-Cofreces, N.B.; Baixauli, F.; Sanchez-Madrid, F. Immune synapse: Conductor of orchestrated organelle movement. Trends Cell Biol. 2014, 24, 61–72. [Google Scholar] [CrossRef] [Green Version]
  9. Vicente-Manzanares, M.; Sánchez-Madrid, F. Role of the cytoskeleton during leukocyte responses. Nat. Rev. Immunol. 2004, 4, 110–122. [Google Scholar] [CrossRef]
  10. Kupfer, A.; Swain, S.L.; Janeway, C.A.; Singer, S.J. The specific direct interaction of helper T cells and antigen-presenting B cells. Proc. Natl. Acad. Sci. USA 1986, 83, 6080–6083. [Google Scholar] [CrossRef] [Green Version]
  11. Varma, R.; Campi, G.; Yokosuka, T.; Saito, T.; Dustin, M.L. T cell receptor-proximal signals are sustained in peripheral microclusters and terminated in the central supramolecular activation cluster. Immunity 2006, 25, 117–127. [Google Scholar] [CrossRef] [Green Version]
  12. Martin-Cofreces, N.B.; Robles-Valero, J.; Cabrero, J.R.; Mittelbrunn, M.; Gordon-Alonso, M.; Sung, C.H.; Alarcon, B.; Vazquez, J.; Sanchez-Madrid, F. MTOC translocation modulates IS formation and controls sustained T cell signaling. J. Cell Biol. 2008, 182, 951–962. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Onnis, A.; Finetti, F.; Baldari, C.T. Vesicular Trafficking to the Immune Synapse: How to Assemble Receptor-Tailored Pathways from a Basic Building Set. Front. Immunol. 2016, 7, 50. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  14. Quintana, A.; Schwindling, C.; Wenning, A.S.; Becherer, U.; Rettig, J.; Schwarz, E.C.; Hoth, M. T cell activation requires mitochondrial translocation to the immunological synapse. Proc. Natl. Acad. Sci. USA 2007, 104, 14418–14423. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  15. Bromley, S.K.; Burack, W.R.; Johnson, K.G.; Somersalo, K.; Sims, T.N.; Sumen, C.; Davis, M.M.; Shaw, A.S.; Allen, P.M.; Dustin, M.L. The immunological synapse. Annu. Rev. Immunol. 2001, 19, 375–396. [Google Scholar] [CrossRef]
  16. Martin-Cofreces, N.B.; Vicente-Manzanares, M.; Sanchez-Madrid, F. Adhesive Interactions Delineate the Topography of the Immune Synapse. Front. Cell Dev. Biol. 2018, 6, 149. [Google Scholar] [CrossRef]
  17. Martin-Cofreces, N.B.; Sanchez-Madrid, F. Sailing to and Docking at the Immune Synapse: Role of Tubulin Dynamics and Molecular Motors. Front. Immunol. 2018, 9, 1174. [Google Scholar] [CrossRef]
  18. Purbhoo, M.A.; Liu, H.; Oddos, S.; Owen, D.M.; Neil, M.A.; Pageon, S.V.; French, P.M.; Rudd, C.E.; Davis, D.M. Dynamics of subsynaptic vesicles and surface microclusters at the immunological synapse. Sci. Signal. 2010, 3, ra36. [Google Scholar] [CrossRef]
  19. Martin-Cofreces, N.B.; Baixauli, F.; Lopez, M.J.; Gil, D.; Monjas, A.; Alarcon, B.; Sanchez-Madrid, F. End-binding protein 1 controls signal propagation from the T cell receptor. EMBO J. 2012, 31, 4140–4152. [Google Scholar] [CrossRef]
  20. Soares, H.; Henriques, R.; Sachse, M.; Ventimiglia, L.; Alonso, M.A.; Zimmer, C.; Thoulouze, M.I.; Alcover, A. Regulated vesicle fusion generates signaling nanoterritories that control T cell activation at the immunological synapse. J. Exp. Med. 2013, 210, 2415–2433. [Google Scholar] [CrossRef] [Green Version]
  21. Kienzle, C.; von Blume, J. Secretory cargo sorting at the trans-Golgi network. Trends Cell Biol. 2014, 24, 584–593. [Google Scholar] [CrossRef]
  22. Goldenring, J.R. Recycling endosomes. Curr. Opin. Cell Biol. 2015, 35, 117–122. [Google Scholar] [CrossRef] [Green Version]
  23. Mittelbrunn, M.; Sanchez-Madrid, F. Intercellular communication: Diverse structures for exchange of genetic information. Nat. Rev. Mol. Cell Biol. 2012, 13, 328–335. [Google Scholar] [CrossRef] [PubMed]
  24. Villarroya-Beltri, C.; Baixauli, F.; Gutierrez-Vazquez, C.; Sanchez-Madrid, F.; Mittelbrunn, M. Sorting it out: Regulation of exosome loading. Semin. Cancer Biol. 2014, 28, 3–13. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. Torralba, D.; Baixauli, F.; Villarroya-Beltri, C.; Fernández-Delgado, I.; Latorre-Pellicer, A.; Acín-Pérez, R.; Martín-Cófreces, N.B.; Jaso-Tamame, Á.L.; Iborra, S.; Jorge, I.; et al. Priming of dendritic cells by DNA-containing extracellular vesicles from activated T cells through antigen-driven contacts. Nat. Commun. 2018. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  26. Yáñez-Mó, M.; Barreiro, O.; Gordon-Alonso, M.; Sala-Valdés, M.; Sánchez-Madrid, F. Tetraspanin-enriched microdomains: A functional unit in cell plasma membranes. Trends Cell Biol. 2009, 19, 434–446. [Google Scholar] [CrossRef] [PubMed]
  27. Wildenberg, M.E.; Vos, A.C.; Wolfkamp, S.C.; Duijvestein, M.; Verhaar, A.P.; Te Velde, A.A.; van der Brink, G.R.; Hommes, D.W. Autophagy attenuates the adaptive immune response by destabilizing the immunologic synapse. Gastroenterology 2012, 142, 1493–1503. [Google Scholar] [CrossRef]
  28. Finetti, F.; Cassioli, C.; Cianfanelli, V.; Onnis, A.; Paccagnini, E.; Kabanova, A.; Baldari, C.T. The intraflagellar transport protein IFT20 controls lysosome biogenesis by regulating the post-Golgi transport of acid hydrolases. Cell Death Differ. 2020, 27, 310–328. [Google Scholar] [CrossRef]
  29. Finetti, F.; Capitani, N.; Baldari, C.T. Emerging Roles of the Intraflagellar Transport System in the Orchestration of Cellular Degradation Pathways. Front. Cell Dev. Biol. 2019, 7, 292. [Google Scholar] [CrossRef] [Green Version]
  30. Parkinson-Lawrence, E.J.; Shandala, T.; Prodoehl, M.; Plew, R.; Borlace, G.N.; Brooks, D.A. Lysosomal storage disease: Revealing lysosomal function and physiology. Physiol. (Bethesda) 2010, 25, 102–115. [Google Scholar] [CrossRef] [Green Version]
  31. Huber, L.A.; Teis, D. Lysosomal signaling in control of degradation pathways. Curr. Opin. Cell Biol. 2016, 39, 8–14. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Calabia-Linares, C.; Robles-Valero, J.; de la Fuente, H.; Perez-Martinez, M.; Martin-Cofreces, N.; Alfonso-Perez, M.; Gutierrez-Vazquez, C.; Mittelbrunn, M.; Ibiza, S.; Veiga, E.; et al. Endosomal clathrin drives actin accumulation at the immunological synapse. J. Cell Sci. 2011, 124, 820–830. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Pullan, L.; Mullapudi, S.; Huang, Z.; Baldwin, P.R.; Chin, C.; Sun, W.; Tsujimoto, S.; Kolodziej, S.J.; Stoops, J.K.; Lee, J.C.; et al. The endosome-associated protein Hrs is hexameric and controls cargo sorting as a master molecule. Structure 2006, 14, 661–671. [Google Scholar] [CrossRef] [PubMed]
  34. Bissig, C.; Gruenberg, J. Lipid sorting and multivesicular endosome biogenesis. Cold Spring Harb. Perspect. Biol. 2013, 5, a016816. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Thery, C.; Boussac, M.; Veron, P.; Ricciardi-Castagnoli, P.; Raposo, G.; Garin, J.; Amigorena, S. Proteomic analysis of dendritic cell-derived exosomes: A secreted subcellular compartment distinct from apoptotic vesicles. J. Immunol. 2001, 166, 7309–7318. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  36. Escola, J.M.; Kleijmeer, M.J.; Stoorvogel, W.; Griffith, J.M.; Yoshie, O.; Geuze, H.J. Selective enrichment of tetraspan proteins on the internal vesicles of multivesicular endosomes and on exosomes secreted by human B-lymphocytes. J. Biol. Chem. 1998, 273, 20121–20127. [Google Scholar] [CrossRef] [Green Version]
  37. Kalra, H.; Simpson, R.J.; Ji, H.; Aikawa, E.; Altevogt, P.; Askenase, P.; Bond, V.C.; Borras, F.E.; Breakefield, X.; Budnik, V.; et al. Vesiclepedia: A compendium for extracellular vesicles with continuous community annotation. PLoS Biol. 2012, 10, e1001450. [Google Scholar] [CrossRef] [Green Version]
  38. Skotland, T.; Sandvig, K.; Llorente, A. Lipids in exosomes: Current knowledge and the way forward. Prog. Lipid Res. 2017, 66, 30–41. [Google Scholar] [CrossRef]
  39. Mittelbrunn, M.; Gutierrez-Vazquez, C.; Villarroya-Beltri, C.; Gonzalez, S.; Sanchez-Cabo, F.; Gonzalez, M.A.; Bernard, A.; Sanchez-Madrid, F. Unidirectional transfer of microRNA-loaded exosomes from T cells to antigen-presenting cells. Nat. Commun. 2011, 2, 282. [Google Scholar] [CrossRef] [Green Version]
  40. Bobrie, A.; Colombo, M.; Raposo, G.; Thery, C. Exosome secretion: Molecular mechanisms and roles in immune responses. Traffic 2011, 12, 1659–1668. [Google Scholar] [CrossRef]
  41. Stoorvogel, W.; Kleijmeer, M.J.; Geuze, H.J.; Raposo, G. The biogenesis and functions of exosomes. Traffic 2002, 3, 321–330. [Google Scholar] [CrossRef] [PubMed]
  42. Thery, C.; Zitvogel, L.; Amigorena, S. Exosomes: Composition, biogenesis and function. Nat. Rev. Immunol. 2002, 2, 569–579. [Google Scholar] [CrossRef] [PubMed]
  43. Gould, S.J.; Booth, A.M.; Hildreth, J.E. The Trojan exosome hypothesis. Proc. Natl. Acad. Sci. USA 2003, 100, 10592–10597. [Google Scholar] [CrossRef] [Green Version]
  44. Bache, K.G.; Brech, A.; Mehlum, A.; Stenmark, H. Hrs regulates multivesicular body formation via ESCRT recruitment to endosomes. J. Cell Biol. 2003, 162, 435–442. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Hurley, J.H. ESCRT complexes and the biogenesis of multivesicular bodies. Curr. Opin. Cell Biol. 2008, 20, 4–11. [Google Scholar] [CrossRef] [Green Version]
  46. Muralidharan-Chari, V.; Clancy, J.; Plou, C.; Romao, M.; Chavrier, P.; Raposo, G.; D’Souza-Schorey, C. ARF6-regulated shedding of tumor cell-derived plasma membrane microvesicles. Curr. Biol. 2009, 19, 1875–1885. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  47. Colombo, M.; Molita, C.; van Niel, G.; Kowal, J.; Vigneron, J.; Benaroch, P.; Manel, N.; Molita, L.F.; Thery, C.; Raposo, G. Analysis of ESCRT functions in exosome biogenesis, composition and secretion highlights the heterogeneity of extracellular vesicles. J. Cell Sci. 2013, 126, 5553–5565. [Google Scholar] [CrossRef] [Green Version]
  48. Joshi, R.P.; Koretzky, G.A. Diacylglycerol kinases: Regulated controllers of T cell activation, function, and development. Int. J. Mol. Sci. 2013, 14, 6649–6673. [Google Scholar] [CrossRef] [Green Version]
  49. Alonso, R.; Mazzeo, C.; Rodriguez, M.C. Diacylglycerol kinase alpha regulates the formation and polarisation of mature multivesicular bodies involved in the secretion of Fas ligand-containing exosomes in T lymphocytes. Cell Death Differ. 2011, 18, 1161–1173. [Google Scholar] [CrossRef] [Green Version]
  50. Bantug, R.; Galluzzi, L.; Kroemer, G.; Hess, C. The spectrum of T cell metabolism in health and disease. Nat. Rev. Immunol. 2018, 18, 19–34. [Google Scholar] [CrossRef]
  51. Desdín-Micó, G.; Soto-Heredero, G.; Mittelbrunn, M. Mitochondrial activity in T cells. Mitochondrion 2018, 41, 51–57. [Google Scholar] [CrossRef] [PubMed]
  52. Baixauli, F.; Martin-Cofreces, N.B.; Morlino, G.; Carrasco, Y.R.; Calabia-Linares, C.; Veiga, E.; Serrador, J.M.; Sanchez -Madrid, F. The mitochondrial fission factor dynamin-related protein 1 modulates T-cell receptor signalling at the immune synapse. EMBO J. 2011, 30, 1238–1250. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Quintana, A.; Kummerow, C.; Junker, C.; Becherer, U.; Hoth, M. Morphological changes of T cells following formation of the immunological synapse modulate intracellular calcium signals. Cell Calcium 2009, 45, 109–122. [Google Scholar] [CrossRef] [PubMed]
  54. Wang, R.; Dillon, C.P.; Shi, L.Z.; Milasta, S.; Carter, R.; Finkelstein, D.; McCormick, L.L.; Flitzgerald, P.; Chi, H.; Munger, J.; et al. The transcription factor Myc controls metabolic reprogramming upon T lymphocyte activation. Immunity 2011, 35, 871–882. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Kaminski, M.M.; Sauer, S.W.; Klemke, C.D.; Suss, D.; Okun, J.G.; Krammer, P.H.; Gulow, K. Mitochondrial reactive oxygen species control T cell activation by regulating IL-2 and IL-4 expression: Mechanism of ciprofloxacin-mediated immunosuppression. J. Immunol. 2010, 184, 4827–4841. [Google Scholar] [CrossRef] [Green Version]
  56. Sena, L.A.; Li, S.; Jairaman, A.; Prakriya, M.; Ezponda, T.; Hildeman, D.A.; Wang, C.R.; Schumacker, P.T.; Licht, J.D.; Perlman, H.; et al. Mitochondria are required for antigen-specific T cell activation through reactive oxygen species signaling. Immunity 2013, 38, 225–236. [Google Scholar] [CrossRef] [Green Version]
  57. Fransen, M.; Lismont, C.; Walton, P. The Peroxisome-Mitochondria Connection: How and Why? Int. J. Mol. Sci. 2017, 18, 1126. [Google Scholar] [CrossRef]
  58. Lazarow, P.B. Peroxisome biogenesis: Advances and conundrums. Curr. Opin. Cell Biol. 2003, 15, 489–497. [Google Scholar] [CrossRef]
  59. Tabak, F.; Braakman, I.; van der Zand, A. Peroxisome formation and maintenance are dependent on the endoplasmic reticulum. Annu. Rev. Biochem. 2013, 82, 723–744. [Google Scholar] [CrossRef]
  60. Sakai, Y.; Oku, M.; van der Klei, I.J.; Kiel, J.A. Pexophagy: Autophagic degradation of peroxisomes. Biochim. Biophys. Acta 2006, 1763, 1767–1775. [Google Scholar] [CrossRef] [Green Version]
  61. Robinson, A.; Waddington, K.E.; Pineda-Torra, I.; Jury, E.C. Transcriptional Regulation of T-Cell Lipid Metabolism: Implications for Plasma Membrane Lipid Rafts and T-Cell Function. Front. Immunol. 2017, 8, 1636. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  62. Mothe-Satney, I.; Murdaca, J.; Sibille, B.; Rousseau, A.S.; Squillance, R.; Le Menn, G.; Rekima, A.; Larbret, F.; Pele, J.; Verhasselt, V.; et al. A role for Peroxisome Proliferator-Activated Receptor Beta in T cell development. Sci. Rep. 2016, 6, 34317. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  63. Lismont, C.; Nordgren, M.; van Veldhoven, P.P.; Fransen, M. Redox interplay between mitochondria and peroxisomes. Front. Cell Dev. Biol. 2015, 3, 35. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Wanders, R.J.; Waterham, H.R.; Ferdinandusse, S. Metabolic Interplay between Peroxisomes and Other Subcellular Organelles Including Mitochondria and the Endoplasmic Reticulum. Front. Cell Dev. Biol. 2015, 3, 83. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Ganguli, G.; Mukherjee, U.; Sonawane, A. Peroxisomes and Oxidative Stress: Their Implications in the Modulation of Cellular Immunity During Mycobacterial Infection. Front. Microbiol. 2019, 10, 1121. [Google Scholar] [CrossRef] [Green Version]
  66. Panieri, E.; Santoro, M.M. ROS homeostasis and metabolism: A dangerous liason in cancer cells. Cell Death Dis. 2016, 7, e2253. [Google Scholar] [CrossRef]
  67. Brenner, D.; Mak, T.W. Mitochondrial cell death effectors. Curr. Opin. Cell Biol. 2009, 21, 871–877. [Google Scholar] [CrossRef]
  68. Huybrechts, S.J.; van Veldhoven, P.P.; Brees, C.; Mannaerts, G.P.; Los, G.V.; Fransen, M. Peroxisome dynamics in cultured mammalian cells. Traffic 2009, 10, 1722–1733. [Google Scholar] [CrossRef]
  69. Schmielau, J.; Finn, O.J. Activated granulocytes and granulocyte-derived hydrogen peroxide are the underlying mechanism of suppression of t-cell function in advanced cancer patients. Cancer Res. 2001, 61, 4756–4760. [Google Scholar]
  70. Schmielau, J.; Nalesnik, M.A.; Finn, O.J. Suppressed T-cell receptor zeta chain expression and cytokine production in pancreatic cancer patients. Clin. Cancer Res. 2001, 7 (Suppl. 3), 933s–939s. [Google Scholar]
  71. Klemke, M.; Samstag, Y. Molecular mechanisms mediating oxidative stress-induced T-cell suppression in cancer. Adv. Enzym. Regul. 2009, 49, 107–112. [Google Scholar] [CrossRef] [PubMed]
  72. Franchina, D.G.; Dostert, C.; Brenner, D. Reactive Oxygen Species: Involvement in T Cell Signaling and Metabolism. Trends Immunol. 2018, 39, 489–502. [Google Scholar] [CrossRef] [PubMed]
  73. Belikov, A.V.; Schraven, B.; Simeoni, L. T cells and reactive oxygen species. J. Biomed. Sci. 2015, 22, 85. [Google Scholar] [CrossRef] [Green Version]
  74. Sena, A.; Chandel, N.S. Physiological roles of mitochondrial reactive oxygen species. Mol. Cell 2012, 48, 158–167. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Brand, K.A.; Hermfisse, U. Aerobic glycolysis by proliferating cells: A protective strategy against reactive oxygen species. FASEB J. 1997, 11, 388–395. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Van der Windt, J.; Pearce, E.L. Metabolic switching and fuel choice during T-cell differentiation and memory development. Immunol. Rev. 2012, 249, 27–42. [Google Scholar] [CrossRef] [Green Version]
  77. Carr, E.L.; Kelman, A.; Wu, G.S.; Gopaul, R.; Senkevitch, E.; Aghvanyan, A.; Turay, A.M.; Frauwirth, K.A. Glutamine uptake and metabolism are coordinately regulated by ERK/MAPK during T lymphocyte activation. J. Immunol. 2010, 185, 1037–1044. [Google Scholar] [CrossRef] [Green Version]
  78. Jacobs, S.R.; Herman, C.E.; Maciver, N.J.; Wofford, J.A.; Wieman, H.L.; Hammen, J.J.; Rathmell, J.C. Glucose uptake is limiting in T cell activation and requires CD28-mediated Akt-dependent and independent pathways. J. Immunol. 2008, 180, 4476–4486. [Google Scholar] [CrossRef] [Green Version]
  79. Chang, C.H.; Curtis, J.D.; Maggi, L.B., Jr.; Faubert, B.; Villarino, A.V.; O’Sullivan, D.; Huang, S.C.; van der Windt, G.J.; Blagih, J.; Qiu, J.; et al. Posttranscriptional control of T cell effector function by aerobic glycolysis. Cell 2013, 153, 1239–1251. [Google Scholar] [CrossRef] [Green Version]
  80. Düvel, K.; Yecies, J.L.; Menon, S.; Raman, P.; Lipovsky, A.I.; Souza, A.L.; Triantafellow, E.; Ma, Q.; Gorski, R.; Cleaver, S.; et al. Activation of a metabolic gene regulatory network downstream of mTOR complex 1. Mol. Cell 2010, 39, 171–183. [Google Scholar] [CrossRef] [Green Version]
  81. Frauwirth, K.A.; Riley, J.L.; Harris, M.H.; Parry, R.V.; Rathmell, J.C.; Elstrom, R.L.; June, C.H.; Thompson, C.B. The CD28 signaling pathway regulates glucose metabolism. Immunity 2002, 16, 769–777. [Google Scholar] [CrossRef] [Green Version]
  82. Jackson, S.H.; Devadas, S.; Kwon, J.; Pinto, L.A.; Williams, M.S. T cells express a phagocyte-type NADPH oxidase that is activated after T cell receptor stimulation. Nat. Immunol. 2004, 5, 818–827. [Google Scholar] [CrossRef]
  83. Ron-Harel, N.; Santos, D.; Ghergurovich, J.M.; Sage, P.T.; Reddy, A.; Lovitch, S.B.; Dephoure, N.; Satterstrom, F.K.; Sheffer, M.; Spinelli, J.B.; et al. Mitochondrial Biogenesis and Proteome Remodeling Promote One-Carbon Metabolism for T Cell Activation. Cell Metab. 2016, 24, 104–117. [Google Scholar] [CrossRef] [Green Version]
  84. Youngblood, B.; Hale, J.S.; Kissick, H.T.; Ahn, E.; Xu, X.; Wieland, A.; Araki, K.; West, E.E.; Ghoneim, H.E.; Fan, Y.; et al. Effector CD8 T cells dedifferentiate into long-lived memory cells. Nature 2017, 552, 404–409. [Google Scholar] [CrossRef]
  85. Gray, S.M.; Kaech, S.M.; Staron, M.M. The interface between transcriptional and epigenetic control of effector and memory CD8⁺ T-cell differentiation. Immunol. Rev. 2014, 261, 157–168. [Google Scholar] [CrossRef] [Green Version]
  86. Sukumar, M.; Liu, J.; Mehta, G.U.; Patel, S.J.; Roychoudhuri, R.; Crompton, J.G.; Klebanoff, C.A.; Ji, Y.; Li, P.; Yu, Z.; et al. Mitochondrial Membrane Potential Identifies Cells with Enhanced Stemness for Cellular Therapy. Cell Metab. 2016, 23, 63–76. [Google Scholar] [CrossRef] [Green Version]
  87. Lochner, M.; Berod, L.; Sparwasser, T. Fatty acid metabolism in the regulation of T cell function. Trends Immunol. 2015, 36, 81–91. [Google Scholar] [CrossRef]
  88. Yang, K.; Shrestha, S.; Zeng, H.; Karmaus, P.W.; Neale, G.; Vogel, P.; Guertin, D.A.; Lamb, R.F.; Chi, H. T cell exit from quiescence and differentiation into Th2 cells depend on Raptor-mTORC1-mediated metabolic reprogramming. Immunity 2013, 39, 1043–1056. [Google Scholar] [CrossRef] [Green Version]
  89. McGarry, D.; Brown, N.F. The mitochondrial carnitine palmitoyltransferase system. From concept to molecular analysis. Eur. J. Biochem. 1997, 244, 1–14. [Google Scholar] [CrossRef]
  90. Longo, N.; Frigeni, M.; Pasquali, M. Carnitine transport and fatty acid oxidation. Biochim. Biophys. Acta 2016, 1863, 2422–2435. [Google Scholar] [CrossRef]
  91. Sayre, N.L.; Lechleiter, J.D. Fatty acid metabolism and thyroid hormones. Curr. Trends Endocinol. 2012, 6, 65–76. [Google Scholar]
  92. Houten, S.M.; Wanders, R.J. A general introduction to the biochemistry of mitochondrial fatty acid β-oxidation. J. Inherit. Metab. Dis. 2010, 33, 469–477. [Google Scholar] [CrossRef] [Green Version]
  93. Cluxton, D.; Petrasca, A.; Moran, B.; Fletcher, J.M. Differential Regulation of Human Treg and Th17 Cells by Fatty Acid Synthesis and Glycolysis. Front. Immunol. 2019, 10, 115. [Google Scholar] [CrossRef] [Green Version]
  94. Pan, Y.; Tian, T.; Park, C.O.; Lofftus, S.Y.; Mei, S.; Liu, X.; Luo, C.; O’Malley, J.T.; Gehad, A.; Teague, J.E.; et al. Survival of tissue-resident memory T cells requires exogenous lipid uptake and metabolism. Nature 2017, 543, 252–256. [Google Scholar] [CrossRef] [Green Version]
  95. Singleton, L.; Roybal, K.T.; Sun, Y.; Fu, G.; Gascoigne, N.R.; van Oers, N.S.; Wulfing, C. Spatiotemporal patterning during T cell activation is highly diverse. Sci. Signal. 2009, 2, ra15. [Google Scholar] [CrossRef] [Green Version]
  96. Zaru, R.; Berrie, C.P.; Iurisci, C.; Corda, D.; Valitutti, S. CD28 co-stimulates TCR/CD3-induced phosphoinositide turnover in human T lymphocytes. Eur. J. Immunol. 2001, 31, 2438–2447. [Google Scholar] [CrossRef]
  97. Sun, Y.; Dandekar, R.D.; Mao, Y.S.; Yin, H.L.; Wulfing, C. Phosphatidylinositol (4,5) bisphosphate controls T cell activation by regulating T cell rigidity and organization. PLoS ONE 2011, 6, e27227. [Google Scholar] [CrossRef] [Green Version]
  98. Zhang, L.; Mao, Y.S.; Janmey, P.A.; Yin, H.L. Phosphatidylinositol 4, 5 bisphosphate and the actin cytoskeleton. Subcell Biochem. 2012, 59, 177–215. [Google Scholar] [CrossRef]
  99. Rocha-Perugini, V.; Gordon-Alonso, M.; Sanchez-Madrid, F. PIP2: Choreographer of actin-adaptor proteins in the HIV-1 dance. Trends Microbiol. 2014, 22, 379–388. [Google Scholar] [CrossRef] [Green Version]
  100. Andreu, Z.; Yáñez-Mó, M. Tetraspanins in extracellular vesicle formation and function. Front. Immunol. 2014, 5, 442. [Google Scholar] [CrossRef] [Green Version]
  101. Quann, E.J.; Merino, E.; Furuta, T.; Huse, M. Localized diacylglycerol drives the polarization of the microtubule-organizing center in T cells. Nat. Immunol. 2009, 10, 627–635. [Google Scholar] [CrossRef]
  102. Nakamura, Y.; Fukami, K. Regulation and physiological functions of mammalian phospholipase C. J. Biochem. 2017, 161, 315–321. [Google Scholar] [CrossRef] [Green Version]
  103. Ladygina, N.; Martin, B.R.; Altman, A. Dynamic palmitoylation and the role of DHHC proteins in T cell activation and anergy. Adv. Immunol. 2011, 109, 1–44. [Google Scholar] [CrossRef] [Green Version]
  104. Almeida, L.; Lochner, M.; Berod, L.; Sparwasser, T. Metabolic pathways in T cell activation and lineage differentiation. Semin. Immunol. 2016, 28, 514–524. [Google Scholar] [CrossRef] [Green Version]
  105. Udenwobele, D.I.; Su, R.C.; Good, S.V.; Ball, T.B.; Shrivastav, S.V.; Shrivastav, A. Myristoylation: An Important Protein Modification in the Immune Response. Front. Immunol. 2017, 8, 751. [Google Scholar] [CrossRef] [Green Version]
  106. Pereira-Leal, J.B.; Hume, A.N.; Seabra, M.C. Prenylation of Rab GTPases: Molecular mechanisms and involvement in genetic disease. FEBS Lett. 2001, 498, 197–200. [Google Scholar] [CrossRef]
  107. Phan, A.T.; Goldrath, A.W.; Glass, C.K. Metabolic and Epigenetic Coordination of T Cell and Macrophage Immunity. Immunity 2017, 46, 714–729. [Google Scholar] [CrossRef] [Green Version]
  108. Valapour, M.; Guo, J.; Schoroeder, J.T.; Keen, J.; Cianferoni, A.; Casolaro, V.; Georas, S.N. Histone deacetylation inhibits IL4 gene expression in T cells. J. Allergy Clin. Immunol. 2002, 109, 238–245. [Google Scholar] [CrossRef]
  109. Warren, J.L.; MacIver, N.J. Regulation of Adaptive Immune Cells by Sirtuins. Front. Endocrinol. 2019, 10, 466. [Google Scholar] [CrossRef]
  110. Harayama, T.; Riezman, H. Understanding the diversity of membrane lipid composition. Nat. Rev. Mol. Cell Biol. 2018, 19, 281–296. [Google Scholar] [CrossRef]
  111. Bartke, N.; Hannun, Y.A. Bioactive sphingolipids: Metabolism and function. J. Lipid Res. 2009, 50, S91–S96. [Google Scholar] [CrossRef] [Green Version]
  112. Avota, E.; de Lira, M.N.; Schneider-Schaulies, S. Sphingomyelin Breakdown in T Cells: Role of Membrane Compartmentalization in T Cell Signaling and Interference by a Pathogen. Front. Cell Dev. Biol. 2019, 7, 152. [Google Scholar] [CrossRef] [Green Version]
  113. Castro, B.M.; Prieto, M.; Silva, L.C. Ceramide: A simple sphingolipid with unique biophysical properties. Prog. Lipid Res. 2014, 54, 53–67. [Google Scholar] [CrossRef]
  114. Börtlein, C.; Draeger, A.; Schoenauer, R.; Kuhlemann, A.; Sauer, M.; Schneider-Schaulies, S.; Avota, E. The Neutral Sphingomyelinase 2 Is Required to Polarize and Sustain T Cell Receptor Signaling. Front. Immunol. 2018, 9, 815. [Google Scholar] [CrossRef]
  115. Trajkovic, K.; Hsu, C.; Chiantia, S.; Rajendran, L.; Wenzel, D.; Wieland, F.; Schwille, P.; Brugger, B.; Simons, M. Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science 2008, 319, 1244–1247. [Google Scholar] [CrossRef]
  116. Lamana, A.; Martin, P.; de la Fuente, H.; Martinez-Muñoz, L.; Cruz-Adalia, A.; Ramirez-Huesca, M.; Escribano, C.; Gollmer, K.; Mellado, M.; Stein, J.V.; et al. CD69 modulates sphingosine-1-phosphate-induced migration of skin dendritic cells. J. Investig. Dermatol. 2011, 131, 1503–1512. [Google Scholar] [CrossRef]
  117. Shiow, L.R.; Rosen, D.B.; Brdickova, N.; Xu, Y.; An, J.; Lanier, L.L.; Cyster, J.G.; Matloubian, M. CD69 acts downstream of interferon-alpha/beta to inhibit S1P1 and lymphocyte egress from lymphoid organs. Nature 2006, 440, 540–544. [Google Scholar] [CrossRef]
  118. Aoki, M.; Aoki, H.; Ramanathan, R.; Hait, N.C.; Takabe, K. Sphingosine-1-Phosphate Signaling in Immune Cells and Inflammation: Roles and Therapeutic Potential. Mediat. Inflamm. 2016, 2016, 8606878. [Google Scholar] [CrossRef] [Green Version]
  119. Chakraborty, P.; Vaena, S.G.; Thyagarajan, K.; Chaetterjee, S.; Al-Khami, A.; Selvam, S.P.; Nguyen, H.; Kang, I.; Wyatt, M.W.; Baliga, U.; et al. Pro-Survival Lipid Sphingosine-1-Phosphate Metabolically Programs T Cells to Limit Anti-tumor Activity. Cell Rep. 2019, 28, 1879–1893.e7. [Google Scholar] [CrossRef] [Green Version]
  120. Chi, H. Sphingosine-1-phosphate and immune regulation: Trafficking and beyond. Trends Pharm. Sci. 2011, 32, 16–24. [Google Scholar] [CrossRef] [Green Version]
  121. Karuppuchamy, T.; Tyler, C.J.; Lundborg, L.R.; Perez-Jeldres, T.; Kimball, A.K.; Clambey, E.T.; Jedlicka, P.; Rivera-Nieves, J. Sphingosine-1-Phosphate Lyase Inhibition Alters the S1P Gradient and Ameliorates Crohn’s-Like Ileitis by Suppressing Thymocyte Maturation. Inflamm. Bowel Dis. 2020, 26, 216–228. [Google Scholar] [CrossRef]
  122. Maceyka, M.; Harikumar, K.B.; Milstien, S.; Spiegel, S. Sphingosine-1-phosphate signaling and its role in disease. Trends Cell Biol. 2012, 22, 50–60. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  123. Colina, C.; Flores, A.; Castillo, C.; Garrido Mdel, R.; Israel, A.; DiPolo, R.; Benaim, G. Ceramide-1-P induces Ca2+ mobilization in Jurkat T-cells by elevation of Ins(1,4,5)-P3 and activation of a store-operated calcium channel. Biochem. Biophys. Res. Commun. 2005, 336, 54–60. [Google Scholar] [CrossRef] [PubMed]
  124. Stahelin, R.V.; Subramanian, P.; Vora, M.; Cho, W.; Chalfant, C.E. Ceramide-1-phosphate binds group IVA cytosolic phospholipase a2 via a novel site in the C2 domain. J. Biol. Chem. 2007, 282, 20467–20474. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Subramanian, P.; Stahelin, R.V.; Szulc, Z.; Bielawska, A.; Cho, W.; Chalfant, C.E. Ceramide 1-phosphate acts as a positive allosteric activator of group IVA cytosolic phospholipase A2 alpha and enhances the interaction of the enzyme with phosphatidylcholine. J. Biol. Chem. 2005, 280, 17601–17607. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  126. Pettus, B.J.; Bielawska, A.; Subramanian, P.; Wijesinghe, D.S.; Maceyka, M.; Leslie, C.C.; Evans, J.H.; Freiberg, J.; Roddy, P.; Hannun, Y.A.; et al. Ceramide 1-phosphate is a direct activator of cytosolic phospholipase A2. J. Biol. Chem. 2004, 279, 11320–11326. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Figure 1. Reactive oxygen species (ROS) and lipids interplay between mitochondria and peroxisomes. Lipid metabolism is tightly regulated during T cell activation. Catabolic routes for energy production as fatty acid oxidation are located inside mitochondria and peroxisomes. The enzymatic production of ROS and different routes regulating their intracellular accumulation also localize at mitochondria and peroxisomes. These processes have an important impact on the transcription of genes, such as cytokines, involved in T cell differentiation and regulatory and effector responses. ROS: Reactive Oxygen Species.
Figure 1. Reactive oxygen species (ROS) and lipids interplay between mitochondria and peroxisomes. Lipid metabolism is tightly regulated during T cell activation. Catabolic routes for energy production as fatty acid oxidation are located inside mitochondria and peroxisomes. The enzymatic production of ROS and different routes regulating their intracellular accumulation also localize at mitochondria and peroxisomes. These processes have an important impact on the transcription of genes, such as cytokines, involved in T cell differentiation and regulatory and effector responses. ROS: Reactive Oxygen Species.
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Figure 2. The cycle of bioactive sphingolipids upon T cell activation. T cell receptor (TCR) stimulation produces endocytic vesicles that regulate signaling. Sphingomyelin (SM) transport to lysosomes by endocytic vesicles allows its conversion to Cer and Sph. Then, the direct or mediated arrival of Sph to the endoplasmic reticulum (ER) leads to its re-conversion into Cer, which goes to the GA through vesicular transport. In the GA, Cer is phosphorylated and ceramide-1-phosphate (C1P) induces Inositol-1, 4, 5-trisphosphate (IP3) production, leading to an opening of plasma membrane and ER Calcium channels that regulates TCR signaling. Besides that, in the ER, Cer is converted to SM, which is then recycled to the plasma membrane, thereby ending the cycle. TCR: T Cell Receptor; Cer: Ceramide; Sph: Sphingosine; ER: Endoplasmic Reticulum; GA: Golgi Apparatus; C1P: Ceramide-1-Phosphate; IP3: Inositol-1, 4, 5-trisphosphate.
Figure 2. The cycle of bioactive sphingolipids upon T cell activation. T cell receptor (TCR) stimulation produces endocytic vesicles that regulate signaling. Sphingomyelin (SM) transport to lysosomes by endocytic vesicles allows its conversion to Cer and Sph. Then, the direct or mediated arrival of Sph to the endoplasmic reticulum (ER) leads to its re-conversion into Cer, which goes to the GA through vesicular transport. In the GA, Cer is phosphorylated and ceramide-1-phosphate (C1P) induces Inositol-1, 4, 5-trisphosphate (IP3) production, leading to an opening of plasma membrane and ER Calcium channels that regulates TCR signaling. Besides that, in the ER, Cer is converted to SM, which is then recycled to the plasma membrane, thereby ending the cycle. TCR: T Cell Receptor; Cer: Ceramide; Sph: Sphingosine; ER: Endoplasmic Reticulum; GA: Golgi Apparatus; C1P: Ceramide-1-Phosphate; IP3: Inositol-1, 4, 5-trisphosphate.
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Dosil, S.G.; Rojas-Gomez, A.; Sánchez-Madrid, F.; Martín-Cófreces, N.B. The Swing of Lipids at Peroxisomes and Endolysosomes in T Cell Activation. Int. J. Mol. Sci. 2020, 21, 2859. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms21082859

AMA Style

Dosil SG, Rojas-Gomez A, Sánchez-Madrid F, Martín-Cófreces NB. The Swing of Lipids at Peroxisomes and Endolysosomes in T Cell Activation. International Journal of Molecular Sciences. 2020; 21(8):2859. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms21082859

Chicago/Turabian Style

Dosil, Sara G., Amelia Rojas-Gomez, Francisco Sánchez-Madrid, and Noa B. Martín-Cófreces. 2020. "The Swing of Lipids at Peroxisomes and Endolysosomes in T Cell Activation" International Journal of Molecular Sciences 21, no. 8: 2859. https://0-doi-org.brum.beds.ac.uk/10.3390/ijms21082859

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