ER as master regulator of membrane trafficking and organelle function

Perinuclear retention of endosomes

The bulk of endosomes and lysosomes exhibit a perinuclear localization clustered around the microtubule-organizing center, together with vesicles of the trans-Golgi network (TGN). This localization enables efficient endosome maturation and cargo trafficking, important for endocytic pathway functions including nutrient uptake, receptor downregulation and cell signaling, host defense against pathogens, and control of cell polarity and cell migration (Alanko et al., 2016; Huotari and Helenius, 2011; Scott et al., 2014). Although the perinuclear clustering of endosomes and Golgi vesicles has been observed for decades, how they are organized and retained was not understood until recently. The ER plays a direct role in the maintenance of this endosomal architecture, orchestrated by two ER-resident ubiquitination enzymes (Cremer et al., 2021; Jongsma et al., 2016; Fig. 1, 1). The E2 ubiquitin conjugation enzyme UBE2J1 interacts with and activates the multimembrane spanning RING domain E3 ubiquitin ligase RNF26. This induces the recruitment and ubiquitination of SQSTM1/p62, a cytosolic ubiquitin adapter best known for its role in selective autophagy. Ubiquitinated SQSTM1/p62 in turn interacts with many ubiquitin-binding organelle-specific adaptor proteins, including T6BP/TAX1BP1 at the TGN (Morriswood et al., 2007) and EPS15B or TOLLIP on endosomes (Katoh et al., 2004; Roxrud et al., 2008). The localization of the E2/E3 pair UBE2J1/RNF26 is confined to the perinuclear ER, which ensures the perinuclear retention of vesicles until released. The ubiquitin-dependent vesicle tethering is released by the deubiquitination enzyme USP15, which is recruited by RNF26 (Jongsma et al., 2016). Although the perinuclear retention of RNF26 depends on its RING domain, it is not known how this mechanism is regulated or how it is coordinated with mechanisms that translocate vesicles to the cell periphery. Dysregulation of ER-UBE2J1/RNF26-mediated vesicle tethering leads to the increased half-life of phosphorylated epidermal growth factor receptors (EGFR) accompanied by prolonged AKT-S473 phosphorylation due to impaired endocytic downregulation (Cremer et al., 2021).

Regulation of endosome translocation to the cell periphery

The nutritional status as well as cellular stress responses influence how endosomes are positioned and utilized for cellular functions (Korolchuk et al., 2011; Raiborg, 2018). During stress and low nutrient conditions, endosomes and lysosomes cluster perinuclearly to facilitate cargo degradation for nutrient supply. When nutrients are available and in the absence of cellular stress, motile endosomes engage in a variety of cellular processes to support cell growth and development. The motile and dispersed late endosomes are less acidic and contain lesser hydrolytic enzymes than the perinuclear late endosomes (Johnson et al., 2016), consistent with their role in functions other than cargo degradation. Indeed, although some endosomes recycle cargo back to the cell surface, others are engaged in plasma membrane repair, protrusion formation, mTORC1 signaling, or secretion of exosomes (Ballabio and Bonifacino, 2020; Pu et al., 2016). Importantly many of these responses are coordinated through the ER. There are two established mechanisms that facilitate the centrifugal transport of late endosomes: the protrudin-dependent pathway, whose function depends on ER-resident proteins, and the BORC-dependent pathway, which is inhibited by ER stress (see below). Thus, the ER is a master regulator of endosome positioning through the control of mechanisms that promote their perinuclear or peripheral localization.

Inhibition of BORC-dependent endosome translocation upon cellular stress

The eight-subunit protein BLOC-one-related complex (BORC) localizes to late endosomes. When nutrient supplies are rich, BORC recruits the small GTPase ARL8B, which through its effector SKIP/PLEKHM2 engages the plus-end-directed microtubule motor Kinesin-1, thus promoting late endosome translocation to the cell periphery (Fig. 1, 2 a). This mechanism is important for cell migration and axonal growth (Farías et al., 2017; Pu et al., 2015). Under cellular stress conditions, however, this pathway is turned off. When cells are deprived of amino acids and growth factors, the BORC complex binds to the endosomal Ragulator complex, making it unable to engage Kinesin-1 (Filipek et al., 2017; Pu et al., 2017). In addition, the ER-resident transmembrane nuclease inositol requiring enzyme 1 (IRE1) plays a role in shutting off BORC-dependent endosome translocation. One branch of the unfolded protein response triggered by ER stress goes through the activation of IRE1. Once activated, IRE1 cleaves and initiates the degradation of certain mRNAs, including the mRNA encoding Blos1, a subunit of the BORC complex. Thus, endosomes cluster perinuclearly, facilitating the lysosomal degradation and clearance of ubiquitinated protein aggregates by microautophagy during ER stress (Bae et al., 2019).

Protrudin-mediated endosome translocation

Protrudin is a transmembrane ER-resident protein that induces ER–endosome MCSs by binding to the late endosomal small GTPase RAB7 in combination with the endosomally enriched lipid phosphatidylinositol 3-phosphate (PtdIns3P; see text box for RAB GTPases and phosphoinositides). In such MCSs, Protrudin hands over Kinesin-1 to the endosomal adapter protein FYCO1, which also interacts with RAB7 and PtdIns3P. This facilitates the translocation of late endosomes along microtubules to the plasma membrane (Raiborg et al., 2015a; Fig. 1, 2 b). The ER-resident pseudoenzyme carnitine palmitoyltransferase 1C (CPT1C) is found in a complex with Protrudin and functions as a nutrient sensor. Under glucose-rich conditions, malonyl-CoA binds CPT1C, and this activates Protrudin-mediated Kinesin-1 handover, which is inhibited upon cellular stress by signaling from the 5' AMP-activated protein kinase (AMPK; Palomo-Guerrero et al., 2019). It is not clear if the seemingly parallel BORC and Protrudin pathways are redundant. As they depend on different small GTPases, ARL8B and RAB7, respectively, they likely translocate different subpopulations of late endosomes (Jongsma et al., 2020). The Protrudin pathway is important for the formation of cellular protrusions like neurites or invadopodia, and this requires that endosomes fuse with the plasma membrane in a Synaptotagmin-VII-dependent manner (Palomo-Guerrero et al., 2019; Pedersen et al., 2020; Raiborg et al., 2015a; Shirane and Nakayama, 2006). In addition, the endosomes contain cargo, such as the metalloprotease MT1-MMP, and the overexpression of Protrudin increases the cell’s invasive behavior by facilitating exocytosis of MT1-MMP in growing invadopodia (Pedersen et al., 2020). Moreover, the Protrudin pathway facilitates mTORC1 signaling from late endosomes (Hong et al., 2017) and stimulates angiogenesis (Arora et al., 2022) and axon regeneration (Petrova et al., 2020).

Synchronization of endosome translocation and lipid transfer

Lipid transfer between closely opposed organelles is mediated by lipid transfer proteins (Reinisch and Prinz, 2021). The ER-resident PDZ domain containing protein 8 (PDZD8) harbors lipid transfer activity and transfers glycerophospholipids and ceramide between membranes in vitro by the use of its synaptotagmin-like mitochondrial-lipid-binding (SMP) domain (Gao et al., 2022; Shirane et al., 2020). In vivo, the depletion of PDZD8 results in a decrease in the abundance of phosphatidylserine (PS) in neuronal endosomes (Shirane et al., 2020) and the accumulation of endosomal PtdIns(4,5)P₂ (Jeyasimman et al., 2021). PDZD8 is important for endosome maturation and their degradative capacity, neuronal integrity, and neurite outgrowth (Gao et al., 2022; Jeyasimman et al., 2021; Shirane et al., 2020).

PDZD8 mediates ER–endosome contact sites by binding to RAB7 and interacts with Protrudin via its transmembrane domain (Elbaz-Alon et al., 2020; Gao et al., 2022; Guillén-Samander et al., 2019; Khan et al., 2021; Shirane et al., 2020; Fig. 1, 2 b). The potential functional relationship between Protrudin and PDZD8 is not completely understood. Since both proteins can form ER–endosome contact sites, why would they need to interact? It is tempting to speculate that these proteins cooperate in the regulation of endosome maturation and translocation, PDZD8, by mediating lipid transfer, and Protrudin by providing a microtubule motor protein. Thus, endosome maturation, function, and translocation can be coordinated efficiently by the ER. This might be especially important in neurons, which depend heavily on endosomal trafficking for their function.

Coordination of microtubule-mediated retrograde and anterograde endosome transport

Endosome positioning entails a constant balance between minus- and plus-end-directed transport along microtubules, mediated by dynein or kinesins, respectively (Bonifacino and Neefjes, 2017; Gennerich and Vale, 2009). With its widespread connection to endosomes, the ER constitutes a unique platform for the organization of the required motor proteins (Friedman et al., 2013). One such possible coordination point centers on the ER-resident protein VAP-A (see text box). Despite being a transmembrane ER protein, Protrudin harbors a VAP-binding FFAT motif, and VAP-A is important for the proper distribution of Protrudin in the ER and for the function of Protrudin in protrusion formation, suggesting that VAP-A facilitates Kinesin-1-dependent endosome translocation (Saita et al., 2009; Fig. 1, 2 b). VAP-A is also implicated in the loss of dynein from endosomal membranes. The dynein binding endosomal protein RILP forms a tripartite complex with RAB7 and the endosomal cholesterol sensor ORP1L, a member of the ORP family (see text box). Under low endosomal cholesterol concentration, the endosomes become tethered to the ER by ORP1L binding to VAP-A, leading to the dissociation of dynein from RILP (Rocha et al., 2009; Fig. 1, 2 c). Thus, although not yet experimentally verified, it is conceivable that VAP-A sites in the ER coordinate the loss of endosomal dynein with the gain of Kinesin-1 through ORP1L-RILP and Protrudin.

Another clue to the role of ER as a coordinator of endosomal motor protein switching comes from the association between Protrudin and the long M1 isoform of the microtubule-severing AAA-ATPase, Spastin. Spastin interacts with Protrudin in the ER and inhibits Protrudin-dependent polarized membrane traffic (Connell et al., 2020). The inhibitory effect of Spastin on endosome translocation requires its ability to interact with the endosomal-sorting complex required for transport (ESCRT)-III proteins, IST1 and CHMP1B, in addition to its microtubule severing-activity. Although not completely understood, this effect might be related to the role of Spastin in the fission of endosomal recycling tubules, which requires the same functional properties as Spastin (Allison et al., 2013). The recruitment of dynein to Spastin-induced microtubule plus ends (Fassier et al., 2013; Lenz et al., 2006; Riano et al., 2009; Zhang et al., 2003) likely counteracts the Protrudin-mediated Kinesin-1-dependent movement of endosomes on microtubule rails toward the cell periphery (Wassmer et al., 2009). The interaction between Spastin and Protrudin in the ER could ensure that the microtubule severing is positioned in close proximity to Protrudin. Thus, the ER coordinates the recruitment of dynein and Kinesin-1 via Spastin and Protrudin, respectively.

Shaping of endosomal tubules

Endocytic cargo that is not destined for lysosomal degradation is sorted into endosomal membrane tubules for recycling back to the plasma membrane or to the Golgi (Huotari and Helenius, 2011; Scott et al., 2014). This process involves membrane budding, tubule extension, and fission to generate cargo-containing vesicles. The ER appears to control both endosomal tubule formation and fission, involving different types of ER–endosome contact sites.

The formation of endosomal recycling tubules requires actin polymerization by the WASH complex, which is coupled to the cargo-sorting retromer machinery by its subunit FAM21 (Derivery et al., 2009; Gomez and Billadeau, 2009; Harbour et al., 2012; Puthenveedu et al., 2010). The transient accumulation of PtdIns4P on endosomes is coupled to a transient burst of WASH-dependent actin nucleation to facilitate retromer function, and the ER is the master regulator of these dynamics (Dong et al., 2016). A type II PI 4-kinase localizes to the WASH complex and produces a local pool of PtdIns4P (Ryder et al., 2013). Endosomal OSBP interacts with PtdIns4P via its PH domain and tethers the endosome to the ER by interacting with VAP-A/B (Fig. 1 3). Here, OSBP transfers PtdIns4P to the ER-resident lipid-phosphatase SAC1, which dephosphorylates PtdIns4P, ensuring the transient PtdIns4P pool on the endosome required for tubule dynamics. In addition, the PtdIns3P binding retromer subunit SNX2 interacts with the ER through VAP-A/B. As actin nucleation by WASH is tightly coupled to retromer-dependent cargo sorting, the ER presumably coordinates their activities through the interaction between VAP-A/B (ER), SNX2 (retromer), and OSBP/PtdIns4P (WASH), all of which localize to the same intracellular hotspots. When this mechanism is perturbed by the depletion of VAP-A/B, SNX2, or OSBP, both PtdIns4P and actin hyper-accumulate on endosomes, and the traffic between endosomes and the Golgi complex is disrupted (Dong et al., 2016). Thus, by regulating endosomal PtdIns4P levels, the ER affects WASH-dependent actin nucleation and retromer function; however, it remains to be seen how PtdIns4P mechanistically interacts with WASH activity.

In addition to regulating endosomal actin dynamics, the ER defines the position and timing of endosome fission (Hoyer et al., 2018; Rowland et al., 2014). Immediately prior to fission, contact sites are formed between ER tubules and endosome buds on sites marked by the WASH component FAM21. The organelles are tethered by the ER membrane protein TMCC1 and endosomal Coronin1C, which is connected to actin on the endosomal buds (Fig. 1 3). Both proteins are required for contact site formation and fission of WASH-containing endosome tubules. Depletion of TMCC1 disrupts recycling of the CI-MPR from endosomes to the Golgi to a similar extent as the depletion of FAM21 (WASH) or VPS35 (retromer), emphasizing the role of ER in this process (Hoyer et al., 2018). Coronin1C confines the localization of actin to bud necks, thereby defining membrane availability for ER–endosome contact sites (Striepen and Voeltz, 2022). How the ER promotes fission is, however, not understood. It will be important to investigate a possible connection with the PtdIns4P-regulated mechanism discussed above. It is tempting to speculate that the final fission step is facilitated by the ESCRT-III-related proteins, IST1 and CHMP1B, which are known to mediate positive membrane bending and constriction (Nguyen et al., 2020) and are connected to the ER by Spastin M1, which is indeed required for the fission of tubules and the recycling of endosomal cargo (Allison et al., 2013).

Coordination of receptor dephosphorylation and formation of multivesicular endosomes

Upon growth factor stimulation, activated growth factor receptors, such as EGFR, are internalized by endocytosis for their final degradation in lysosomes, a process referred to as receptor downregulation (Huotari and Helenius, 2011; Scott et al., 2014). This process ensures that signaling is switched off in a timely manner to prevent hyperproliferation. To attenuate EGFR signaling, the receptors are dephosphorylated and sorted into forming intraluminal vesicles (ILVs) of multivesicular endosomes (MVEs) on their way to the lysosome. It is interesting to note that EGF-stimulation itself induces this process by triggering the dephosphorylation of EGFR and at the same time stimulates ILV formation. Intriguingly, the ER is recruited to promote both tasks.

The phospholipid-binding protein Annexin A1 associates with EGFR-containing MVEs, whereas its ligand S100A11 localizes to the ER (Futter et al., 1993; Gerke and Moss, 2002; Liu et al., 2012; Fig. 1, 4 a). EGFR-induced phosphorylation of Annexin A1 induces the formation of Annexin A1/S100A11-mediated ER–endosome contact sites (Eden et al., 2016). Both Annexin A1 and S100A11 are Ca²⁺ binding proteins, and the contact site formation is aided by the local increase in Ca²⁺, which is induced by the endosomal NAADP-sensitive two-pore Ca²⁺ channel TPC1 (Kilpatrick et al., 2017). These contact sites promote EGF-induced ILV formation (Eden et al., 2016; White et al., 2006; Wong et al., 2018). First, the protein tyrosine phosphatase 1B (PTP1B), which is embedded in the cytoplasmic face of the ER, dephosphorylates EGFRs on the endosomes, depending on Annexin A1/S100A11-mediated ER–endosome contact sites (Eden et al., 2010). At the same time, the EGFRs are sorted into forming ILVs by the ESCRT protein machinery, which interacts with the ubiquitinated EGFRs and mediates membrane deformation and scission to generate ILVs (Migliano et al., 2022). Interestingly, the ESCRT proteins HRS and STAM are dephosphorylated by PTP1B, implying that the Annexin A1/S100A11-mediated ER–endosome contact sites can regulate ESCRT function (Eden et al., 2010; Stuible et al., 2010). This functional relationship, which could facilitate the progression of cargo through downstream ESCRTs and ILV formation, needs further investigation.

In addition to acting on the ESCRT machinery through PTP1B, the Annexin A1/S100A11-mediated ER–endosome contact sites can facilitate ILV formation through a different mechanism. ILVs are rich in cholesterol, and high levels of endosomal cholesterol are required to form ILVs (Möbius et al., 2003). Cholesterol can be supplied by the uptake of low-density lipoprotein (LDL) by receptor-mediated endocytosis (Anderson et al., 1977). To fuel ILV formation in the absence of LDL, cholesterol needs to come from internal sources such as the ER. When endosomal cholesterol levels are low, EGF-stimulated ILV formation depends on Annexin A1/S100A11-mediated ER–endosome contacts (Eden et al., 2016). As Annexin A1/S100A11 does not harbor sterol transfer properties, such delivery has to be coordinated with a lipid transfer protein. The endosomal cholesterol sensor ORP1L localizes to Annexin A1-dependent ER–endosome contact sites in the absence of LDL and is a plausible candidate for this activity. When endosomal cholesterol levels are low, a conformational change exposes the ORP1L FFAT-motif, inducing binding to VAP-A in the ER (Rocha et al., 2009; Fig.1, 4 b). Here, ORP1L facilitates the transfer of cholesterol from the ER to EGFR-containing endosomes and stimulates ILV formation in a manner that requires its interaction with VAP-A (Eden et al., 2016). Whether ORP1L is directly responsible for transfer or regulates another lipid transfer protein is unresolved. It is interesting to note that under cholesterol depletion, the interaction of ORP1L with VAP-A in the ER at the same time leads to the loss of dynein and the HOPS complex from the endosomes (van der Kant et al., 2013). This will inhibit the perinuclear translocation and fusion of MVEs when cholesterol levels are low, and could thus halt the maturation of endosomes to ensure proper sorting of EGFRs into ILVs by use of cholesterol from the ER.

Control of endosome maturation and homeostasis

To maintain lipid homeostasis, the ER influences the transport of lipids from the ER to endosomes and vice versa. In addition to the lipid transporters PDZD8 and ORP1L mentioned above, STARD3 resides in EGFR-negative endosomes and facilitates cholesterol transport from the ER to the endosomes, which are anchored to the ER through VAP-A/B and MOSPD2 (Alpy et al., 2013; Voilquin et al., 2019; Wilhelm et al., 2017). Conversely, NPC1 facilitates the transfer of cholesterol from endosomes to the ER by interaction with ORP5 in the ER (Fig.1, 4 c; Du et al., 2011; Raiborg et al., 2015b). The ER plays a pivotal role in maintaining endosome homeostasis and maturation by regulating endosomal identity (Wu and Voeltz, 2021), and the sorting and trafficking of hydrolytic enzymes and endocytosed proteins and lipids, as exemplified above. Mutations in NPC1 lead to the accumulation of cholesterol in endosomes, causing the neurodegenerative disease Nieman-Pick (Mukherjee and Maxfield, 2004). Accumulation of endocytosed or cellular material caused by dysfunctional ER–endosome MCS proteins can thus lead to severe metabolic and developmental defects, as manifested by genetic diseases, collectively termed lysosomal storage disorders (Platt et al., 2012).

Biogenesis of the phagophore membrane

Although several cellular membranes have been proposed as the origin of phagophore membranes, there is little doubt that the ER is a major source (Lamb et al., 2013; Melia et al., 2020). The fact that autophagosome membranes, in contrast to other cellular membranes, are almost devoid of transmembrane proteins (Fengsrud et al., 2000) suggests that much of the phagophore could originate from de novo membrane synthesis rather than budding from existing membranes. In support of this, a large cytosolic protein required for autophagosome biogenesis, ATG2, is an elongated lipid transporter that contains a hydrophobic groove through which lipids can slide in an efficient way (Ghanbarpour et al., 2021; Maeda et al., 2019). ATG2 could thus function in MCSs that bridge the lipid-synthesizing ER and the forming phagophore.

ATG2 interacts with two lipid scramblases in the ER membrane, TMEM41B and VMP1, and with a lipid scramblase on Golgi-derived vesicles, ATG9 (Ghanbarpour et al., 2021; Judith et al., 2019; Noda, 2021). Lipid scramblases transfer lipids from one membrane leaflet to the other, and it has been proposed that even a single ATG9-containing vesicle might act as a seed for phagophore biogenesis (Ghanbarpour et al., 2021). In this model, ATG2 mediates lipid transport from the ER membrane to the seeding vesicle, whereas TMEM41B and VMP1 re-equilibrate the leaflets of the ER during lipid extraction. In the seed vesicle, ATG9 scrambles ER-derived lipids upon their delivery to allow phagophore expansion (Fig. 2). Even though this is an attractive model that explains the requirement for lipid transporters and scramblases in autophagosome biogenesis, it still needs to be verified experimentally. Hybrid organelles consisting of membranes from endosomes and the cis-Golgi have recently been put forward as precursors of phagophores (Kumar et al., 2021), and it remains plausible that autophagosomes can originate from membranes other than the ER, at least under some conditions (Melia et al., 2020).

Class III PI 3-kinase (PI3K-III), which phosphorylates PtdIns into PtdIns3P, is required for phagophore biogenesis, and it is conceivable that PtdIns3P contributes to defining the sites of phagophore initiation. Indeed, ATG14, a subunit of the autophagy-specific version of PI3K-III, localizes to ER sites, and this localization is required for autophagy (Matsunaga et al., 2010). The PtdIns3P-binding protein DFCP1 is recruited to PtdIns3P-containing ER subdomains in response to amino acid starvation, a classical way to induce autophagy, and is a likely PtdIns3P effector in autophagosome biogenesis. Due to their omega shape in light microscopy, DFCP1-containing ER subdomains are referred to as omegasomes (Axe et al., 2008). The exact spatial and functional relationships of omegasomes with phagophores are not known, but current evidence suggests that omegasomes could represent ER subdomains that are involved in the elongation and sculpting of the phagophore.

PtdIns3P is not only found on ER subdomains but also on phagophore membranes (Cheng et al., 2014), suggesting the involvement of additional PtdIns3P-binding proteins in phagophore biogenesis. PtdIns3P-binding proteins of the WIPI family are good candidates as they interact with ATG2 and localize to the growing phagophore. WIPI4, which shows the highest affinity to ATG2, binds to one of the tips of ATG2, consistent with the idea that ATG2 could be recruited by WIPI4 to form a lipid-transporting bridge between the ER and the tip of the phagophore (Chowdhury et al., 2018). However, the spatiotemporal relationships between ATG2, ATG9, PtdIns3P, DFCP1, and WIPI4 during phagophore biogenesis remain to be defined.

Control of autophagosome fusion

Autophagy culminates in the fusion of autophagosomes with lysosomes. The membranes of late endosomes and lysosomes contain the small GTPase RAB7, and among the RAB7 effectors they recruit are ORP1L, RILP, and PLEKHM1. As described above, ORP1L is a cholesterol sensor that forms tripartite contacts with RAB7 and the dynein adaptor RILP, thereby promoting dynein-mediated transport of late endosomes and lysosomes toward the microtubule organizing center. The endolysosomal protein PLEKHM1, in concert with RILP, recruits the HOPS complex, which promotes fusion between lysosomes and autophagosomes (McEwan et al., 2015; Wijdeven et al., 2016). If the lysosomes have low cholesterol content, the FFAT motif of ORP1L is exposed and engages in interaction with VAP-A in the ER membrane. The cholesterol-free conformation of ORP1L not only prevents the interaction of RILP with dynein and HOPS, but also dissociates PLEKHM1, and this inhibits both lysosome motility toward the microtubule organizing center and fusion between lysosomes and autophagosomes (Wijdeven et al., 2016). Thus, autophagic flux is positively regulated by cholesterol and negatively controlled by the ER–lysosome MCSs. Since autolysosomes, like lysosomes, contain RAB7 and ORP1L, their motility is regulated in the same manner.

Ca²⁺ transfer between ER and mitochondria

The ER contains a highly concentrated pool of intraluminal Ca²⁺, which is involved in the regulation of processes ranging from ATP production to the onset of apoptosis. Upon activation of the IP₃ gated Ca²⁺ channel (IP3R), the ER can release Ca²⁺ ions to the surrounding milieu. The close proximity of the ER–mitochondria MCSs allows for a directional flow of Ca²⁺ to enter the mitochondria through the voltage-dependent anion channel 1 (VDAC1) in the outer membrane (Gincel et al., 2001; Rapizzi et al., 2002) and the mitochondrial calcium uniporter (MCU1) in the inner membrane (Kirichok et al., 2004). The glucose-regulated protein 75 (GRP75) bridges the two organelles to form a stable “synapse” for the Ca²⁺ transfer by binding both VDAC1 and IP3R (Szabadkai et al., 2006; Fig. 3 a). This synapse is necessary for mitochondrial function, homeostasis, energy production, and viability. The concentration of Ca²⁺ inside the inner mitochondrial membrane has consequences for ATP production through the regulation of Ca²⁺-dependent enzymes in the Krebs cycle (Rossi et al., 2019). However, excessive levels of Ca²⁺ ions can induce apoptosis (Rasola and Bernardi, 2011; see below).

Several protein–protein interactions are involved in the flow of Ca²⁺ between these membranes, likely due to its important functions in the regulation of both cell growth and cell death. One such example is VAP-B on ER, which binds PTPIP51 on the outer mitochondrial membrane to ensure proper Ca²⁺ release from ER lumen to the mitochondria (De Vos et al., 2012). The interaction is mediated via the FFAT-like motif on PTPIP51 and the MSP motif on VAP-B. A mutated version of VAP-B, VAPB^P56S, and the dysregulation of Ca²⁺ flow between ER and mitochondria are both associated with amyotrophic lateral sclerosis, highlighting the physiological importance of this connection (Langou et al., 2010; Nishimura et al., 2004). Mitofusin (MFN) is another example of a protein bridge that supports Ca²⁺ transport from the ER to the mitochondria. The ER membrane carries MFN2, which can bind heterotypically or homotypically to MFN1 or MFN2, respectively, on the mitochondrial membrane. This interaction aids in forming a stable bridge between the organelles during mitochondria Ca²⁺ uptake (de Brito and Scorrano, 2008).

The ER-resident lipid transfer protein PDZD8 (described under “Synchronization of endosome translocation and lipid transfer”) is a mammalian paralog of the yeast protein Mmm1, a member of the ER–mitochondrial encounter structure (ERMES) complex, implicated in the formation of ER–mitochondria contact sites (Hirabayashi et al., 2017; Wideman et al., 2018). In neurons, PDZD8 is necessary for the contact between ER and mitochondria during Ca²⁺ transport. This MCS is proposed to be utilized by neurons to regulate dendritic excitability and plasticity during signal transduction. PDZD8 establishes directionality of Ca²⁺ flux from IP3R and ryanodine receptors toward the mitochondrion. In the absence of PDZD8, the cytosolic Ca²⁺ concentration increases. The protein partner of PDZD8 on the mitochondria membrane has yet to be elucidated (Hirabayashi et al., 2017).

Regulation of apoptosis

In the context of cell death, the ER-resident protein BAP31 is crucial for ER–mitochondrial tethering (Ng et al., 1997), and several BAP31 complexes are associated with cell death, including FIS1 (Iwasawa et al., 2011) and CDIP1 (Namba et al., 2013; Fig. 3 a). These protein–protein contacts regulate cell death by establishing signaling platforms, translating apoptotic cues, and engaging in the onset of apoptosis (Iwasawa et al., 2011; Mattson and Chan, 2003; Namba et al., 2013). The initiation of cell death will activate either the intrinsic or extrinsic apoptotic pathway. However, a few common events occur independently of the mode of action. These include the activation of caspases by cleavage and the release of cytochrome c into the cytosol from the mitochondria. These actions are downstream of an increased flow of Ca²⁺ from ER into the mitochondria. When the concentration of Ca²⁺ reaches a certain threshold, the permeability transition pore (PTP) opens and allows for water molecules and protons to travel freely over the inner mitochondria membrane. This disrupts the proton gradient and induces swelling and rupture of the outer membrane (Halestrap, 2009). As a result, cytochrome c leaks into the cytosol where it binds the apoptotic protease-activating factor 1 (APAF1) machinery (Hardingham and Bading, 2003; Mattson and Chan, 2003), causing the apoptosome to assemble and execute apoptosis (Rasola and Bernardi, 2011).

Cell death can also be mediated through ceramide-induced apoptosis (Obeid et al., 1993). Ceramides synthesized by ER are highly regulated and are normally transported to the Golgi for further processing. A rise in ceramide levels followed by the recruitment of ceramide binding proteins on the outer mitochondria membrane can trigger apoptosis. The exact molecular mechanisms leading to this event remain elusive, but VDAC2 in the outer mitochondrial membrane has recently been shown to bind ceramides and acts as an effector of cell death signals (Dadsena et al., 2019).

Interestingly, BAP31 in the ER–mitochondria MCSs is not restricted to cell death signaling. The mitochondrial membrane protein TOM40 can bind BAP31 to recruit NDUSF4. This protein complex is involved in stress sensing and cellular homeostasis. The lack of BAP31 activates autophagy and glycolysis while reducing mitochondrial oxygen consumption (Namba, 2019). Thus, BAP31 has a role in ER MCS during both self-preservation and self-destruction.

Mitochondrial fission

Homeostasis of mitochondria is maintained by fission and fusion of the organelle. ER–mitochondrial contacts spatially define where the mitochondrion will divide (Abrisch et al., 2020; Friedman et al., 2011), as ER tubules wrap around the mitochondria to indicate the position for fission (Abrisch et al., 2020; Chakrabarti et al., 2018; Korobova et al., 2013; Fig. 3 b). The ER-associated formin, inverted formin-2 (INF2), polymerizes actin filaments to establish a close contact between the two organelles. The INF2-mediated actin polymerization stimulates mitochondria Ca²⁺ uptake. Spire1c is an actin nucleator and resides on the outer mitochondrial membrane during fission. Here it binds INF2 directly to connect the two organelles, as well as initiating nucleation of the actin filaments (Manor et al., 2015). ER tubules in the MCS release Ca²⁺, which enters the mitochondria through the VDAC1 channel. INF2 has been implied in the constriction of the inner mitochondria membrane indirectly by an increase in ER–mitochondria contact sites following an influx of Ca²⁺, which triggers the inner mitochondrial membrane to divide first (Chakrabarti et al., 2018). It is unknown exactly how the inner membrane divides, but the electron transport chain is required for the execution. Constriction of the outer membrane depends on the cytosolic GTPase dynamin related-protein 1 (DRP1) to self-assemble into a ring around the mitochondrion at the fission site. For the Drp1-spiral to form, the outer membrane receptors FIS1 and MFF guide the assembly (Elgass et al., 2013; Koch et al., 2005). Drp1 binds actin filaments and modulates actin bundles in vitro (Ji et al., 2015). It is therefore possible that the actin filaments on the mitochondria surface are anchored directly to the DRP1 spiral. The spiral constricts with the aid of actin–myosin filaments (Smirnova et al., 2001). Thus, the inner membrane scission is followed by constriction of the outer membrane as the Drp1 spiral tightens and ultimately results in the formation of two daughter organelles (Chakrabarti et al., 2018; Korobova et al., 2013).

A recent contribution to the field of mitochondrial fission is the observation that PtdIns4P-containing vesicles derived from lysosomes or the TGN interact with ER–mitochondria MCSs, forming three-way contacts (Boutry and Kim, 2021; Nagashima et al., 2020). Within these contact sites, ORP1L was suggested to transfer PtdIns4P from lysosomes to the mitochondrion, promoting mitochondrial fission (Boutry and Kim, 2021). Likewise, inhibition of the formation of PtdIns4P microdomains on TGN vesicles results in branched and hyperfused mitochondria (Nagashima et al., 2020), thus indicating an important role of ER–mitochondria triple contact sites to finalize fission.

When mitochondria divide, the daughter mitochondria must bear a copy of the mitochondrial DNA (mtDNA) found in the parental mitochondrion. Subpopulations of ER–mitochondria contact sites have been shown to be specifically reserved for and required for the synthesis of mtDNA (Lewis et al., 2016). Following duplication of the nucleoid, DRP1 regulates the mtDNA synthesis and the distribution to the daughter organelles during fission by altering the ER sheets that contact the mitochondria (Ilamathi et al., 2022; Lewis et al., 2016).

Mitochondrial fusion

It has been proposed that mitochondrial fission and fusion events are coordinated to quickly respond to metabolic cues. Hence, as for mitochondrial fission, the ER marks the sites of mitochondrial fusion (Guo et al., 2018), and molecules involved in fusion and fission colocalize in ER–mitochondria contact sites (Abrisch et al., 2020). It has been proposed that ER tethering guides the position and timing of mitochondria fusion, but the exact role of ER–mitochondria MCS is still not clear (Gao and Hu, 2021). Fusion of the outer mitochondria membranes is executed by the mitofusins, MFN1 and MFN2 (Cao et al., 2017; Chen et al., 2003), while Opa1 regulates the fusion of the inner mitochondria membrane (Song et al., 2009; Fig. 3 b). Even though both hetero- and homotypic protein interactions can occur between MFN1 and MFN2, it is the homotypic interactions that are required for fusion (Chen et al., 2003). Interestingly, only a certain protein conformation of mitofusins allows for mitochondrial fusion (Franco et al., 2016). Both MFN1 and DRP1 puncta localize to the ER–mitochondria contact sites during the synchronized fusion and fission events, respectively (Abrisch et al., 2020). To maintain the ER–mitochondria contact during fusion, the Ca²⁺ sensitive molecule Miro will decrease mitochondria motility (Kornmann et al., 2011). Miro is also a motorprotein-adaptor involved in both actin filament and microtubule transport. It is unknown exactly how Miro regulates the translocation of mitochondria during fission and fusion; however, acetylation of Miro at specific Lysine residues has been implicated in the context of mitochondria transportation in neurons (Kalinski et al., 2019). More research is needed to further explore the role of Miro in mitochondria fission and fusion.

Lipid transfer between ER and mitochondria

Mitochondria rely on lipid transport proteins to maintain membrane homeostasis. The evolutionarily conserved ER protein LTC1 is found at MCSs between mitochondria and ER, depending on the mitochondrial import receptors Tom70/71. It has been suggested that the function of LTC1 is to transport and/or sense sterols to maintain correct lipid homeostasis and organelle function (Murley et al., 2015).

Miro is not only involved in mitochondria mobility as discussed above but it also promotes lipid transfer. In this role, Miro has been shown to locate to the outer mitochondrial membrane to recruit the lipid channel transporter VPS13D, which is anchored to the ER by VAP-B. Analogously, Miro regulates lipid transfer in ER–peroxisome contact sites (Guillen-Samander et al., 2021; Kornmann et al., 2011).

The interaction between the ER-resident protein VAP-B and the mitochondrial protein PTPIP51 is also important for lipid transfer (De Vos et al., 2012). The interaction between the proteins is provided by a FFAT-like motif on PTPIP51 and an MSP domain on VAP-B. The contact site can provide the transport of phosphatidic acid from ER to mitochondria via a TRP motif in PTPIP51 (Yeo et al., 2021). Interestingly, the same protein–protein interaction and MCS are also involved in regulating autophagy in mammalian cells. Loss of PTPIP51 or VAP-B results in increased autophagy, while the overexpression of either protein reduces the number of autophagic structures, likely in a Ca²⁺-dependent manner (Gomez-Suaga et al., 2017).

PtdIns4P fuels lipid transfer between ER and Golgi

PtdIns4P is generated by the Golgi-localized phosphatidylinositol kinase 4β (PI4KIIIβ) through the phosphorylation of PtdIns (Balla and Balla, 2006). Oxysterol-binding protein (OSBP) and ORPs (see text box), such as ORP4L, ORP9, and ORP10, are lipid-exchange transporters, which depend on high levels of PtdIns4P in the Golgi, both to form ER–Golgi contact sites and to function in lipid transport. These cytosolic transport proteins contact the Golgi with their PH domain, which binds PtdIns4P and ARF1-GTP, and they tether to the ER via their VAP-binding FFAT motifs, which allows the exchange of PtdIns4P against sterols or PS (Mesmin et al., 2013; Ngo and Ridgway, 2009). The lipid transfer is mediated by the ORD domain and requires PtdIns4P in exchange. The high PtdIns4P levels in the Golgi fuel a counter-transfer of cholesterol (OSBP, ORP4L, and ORP9) or PS (ORP10) to the Golgi (Maeda et al., 2013; Pietrangelo and Ridgway, 2018; Venditti et al., 2019b; Fig. 4 b). The PtdIns4P gradient is maintained by PI4KIIIβ-mediated synthesis of PtdIns4P in the TGN and dephosphorylation of PtdIns4P in the ER by the phosphatase SAC1 (Mesmin et al., 2013; Mesmin et al., 2017).

SAC1 regulates PtdIns4P levels

SAC1 is an ER-resident phosphatase that dephosphorylates PtdIns4P in cis in the ER (Mesmin et al., 2013). However, close contact at ER–Golgi–MCS may allow SAC1 to also act in trans to consume PtdIns4P in the Golgi (Manford et al., 2010), and this activity is promoted by the presence of phosphatidyl-four-phosphate-adaptor-protein-1 (FAPP1) within these contacts (Venditti et al., 2019a; Fig. 4 b). Another regulator of SAC1 phosphatase activity is the neuronally-expressed CPT1C (Sierra et al., 2008), which cooperates with Protrudin as described above. CPT1C senses metabolic changes through binding to malonyl-CoA, an intermediate in de novo long-chain fatty acid synthesis, whose levels correlate with the nutritional state. Under basal conditions, CPT1C inhibits SAC1 activity to maintain normal levels of PtdIns4P in the Golgi, allowing AMPA receptor trafficking to the plasma membrane. Under glucose deprivation, CPT1C releases SAC1 inhibition allowing SAC1 to dephosphorylate PtdIns4P at ER–Golgi–MCS in trans, which results in AMPA receptor retention at the TGN (Casas et al., 2020). In this way, the ER can affect neuronal function and cognition depending on energy status.

Sphingolipid transfer between ER and Golgi

PtdIns4P also plays an important role for sphingolipid transporters: Ceramide transfer protein (CERT) uses its START domain to transfer ceramide from the ER to the TGN for further processing into sphingomyelin (Hanada et al., 2003; Fig. 4 b). CERT forms a contact site by binding to ER-resident VAP proteins via its FFAT motif and its PH domain recognizes PtdIns4P in the Golgi (Hanada et al., 2003; Peretti et al., 2008). The activity of CERT is regulated by phosphorylation and by a negative feedback loop recognizing elevated DAG levels, resulting from sphingomyelin synthesis (Fugmann et al., 2007; Kumagai et al., 2014; Saito et al., 2008). In addition, the START domain can compete with the PH domain for PtdIns4P binding. When the levels of ceramide are high, the START domain is occupied by ceramide, and the shuttling of ceramide from ER to Golgi will occur. When the levels of ceramide are low, the START domain will bind to the PH domain, removing CERT from the Golgi (Prashek et al., 2017).

Phosphatidylinositol-four-phosphate adapter protein 2 (FAPP2; also known as PLEKHA8) possesses a PtdIns4P- and ARF1-GTP-binding PH domain at the N-terminus and a glycolipid transfer protein homology domain at the C-terminus, responsible for glucosylceramide (GlcCer) transport (Godi et al., 2004). Depletion of FAPP2 disrupts GlcCer transport from cis- to trans-Golgi, resulting in a disturbed synthesis of complex glycosphingolipids (D'Angelo et al., 2007; D'Angelo et al., 2013; Halter et al., 2007). How FAPP2 aids in the transport of GlcCer from the cis to the trans-Golgi is not yet fully understood, but may involve ER–Golgi contact sites: FAPP2 has a putative FFAT motif (Backman et al., 2018) and has been suggested to transfer GlcCer retrogradely to the ER, where GlcCer translocates into the lumen. From the ER lumen, GlcCer could be anterogradely transported to the trans-Golgi for further glycosylation into complex glycosphingolipids (Halter et al., 2007). Alternatively or additionally, FAPP2 may mediate the direct transfer of GlcCer from cis- to trans-Golgi (D'Angelo et al., 2007).

Control of Golgi homeostasis

The phosphatidylinositol-transfer protein NIR2 (PYK2 N-terminal domain- interacting receptor 2) contacts the ER through a classical FFAT motif and it has a PtdIns-transfer domain (PITD) that mediates Golgi localization (Amarilio et al., 2005; Kim et al., 2013). Through its PITD domain, NIR2 transfers PtdIns from the ER to the Golgi, and thereby replenishes the substrate for the PtdIns4-kinase. This closes the PtdIns4P cycle of phosphorylation, transfer, and dephosphorylation, which is necessary to fuel sterol and PS transport (Fig. 4 b). In addition, NIR2 has been shown to regulate DAG levels at the Golgi apparatus (Litvak et al., 2005) and it can affect OSBP and CERT localization and activity (Peretti et al., 2008). The action of the lipid-transfer proteins OSBP, CERT, and NIR2 is thus intricately connected and coordinated at ER–Golgi contact sites.

Maintaining lipid homeostasis through lipid-transfer proteins is important for the physiologic function of the secretory pathway. Disturbances in lipid exchange and the resulting imbalances in PtdIns4P or cargo lipids disrupt Golgi morphology, lipid modifications, and anterograde cargo transport processes (Cruz-Garcia et al., 2013; Godi et al., 2004; Litvak et al., 2005; Peretti et al., 2008; Szentpetery et al., 2010; Wakana et al., 2021; Wakana et al., 2015). The ER maintains this balance not only through its direct function at lipid-transfer contact sites but also by anchoring TGN vesicles in the perinuclear area through T6BP/TAX1BP1–SQSTM1/p62–RNF26 as described above (Fig. 1, 1).