Schwann Cell Development and Myelination

Abstract

Glial cells in the peripheral nervous system (PNS), which arise from the neural crest, include axon-associated Schwann cells (SCs) in nerves, synapse-associated SCs at the neuromuscular junction, enteric glia, perikaryon-associated satellite cells in ganglia, and boundary cap cells at the border between the central nervous system (CNS) and the PNS. Here, we focus on axon-associated SCs. These SCs progress through a series of formative stages, which culminate in the generation of myelinating SCs that wrap large-caliber axons and of nonmyelinating (Remak) SCs that enclose multiple, small-caliber axons. In this work, we describe SC development, extrinsic signals from the axon and extracellular matrix (ECM) and the intracellular signaling pathways they activate that regulate SC development, and the morphogenesis and organization of myelinating SCs and the myelin sheath. We review the impact of SCs on the biology and integrity of axons and their emerging role in regulating peripheral nerve architecture. Finally, we explain how transcription and epigenetic factors control and fine-tune SC development and myelination.

STAGES OF SCHWANN CELL DEVELOPMENT

Schwann cells (SCs) derive from neural crest cells, a multipotent group of cells that arises after closure of the neural tube at around embryonic day (E)9.5–E11.5 in the mouse.⁶ Neural crest cells originate at the cranial, trunk, cardiac, and sacral levels of the neural tube, delaminate and migrate throughout the body following three main streams. Depending on the location and migrating stream, neural crest cells give rise to various peripheral nervous system (PNS) neurons, glia, endoneurial fibroblasts (EFs), and other cells including melanocytes and cranial chondrocytes (Le Douarin and Teillet 1974; Joseph et al. 2004).

The formative stages of SC differentiation from neural crest to precursors and then to mature cells (Fig. 1) have been extensively characterized (see Jessen and Mirsky 2019; Muppirala et al. 2021 for recent reviews). Axon-associated SCs originate from trunk neural crest cells that migrate ventrally and rapidly differentiate into SC precursors (Weston 1963; Jessen et al. 1994), which associate with extending axons and surround their growth cones (Wanner et al. 2006). SC precursors retain significant pluripotency and can also give rise to melanocytes, neurons, and chondrocytes (Kastriti et al. 2022; Taveggia and Feltri 2022). In developing nerve trunks, SC precursors interact with axon bundles and generate immature SCs, whose survival depends on the Neuregulin 1 (NRG1) growth factor expressed on axons (Jessen et al. 1994; Meyer and Birchmeier 1995). At this stage, axons of varying caliber are tightly packed into bundles with SCs present typically on the outside. Interaction with axons and NRG1 stimulates the proliferation of SC precursors, generating a sufficient number of cells to surround all axon bundles and deposit a basal lamina.

Figure 1.

Schematic of the formative stages in the Schwann cell (SC) lineage. At the right are electron micrographs showing cross-sections through a myelinating SC and a portion of a Remak SC; axon profiles are labeled A. Key extrinsic signals and their SC receptors (in blue text) are listed below. Axonal signals (red) include type III NRG1, which activates ErbB2/3; Adam 22, which interacts with secreted Lgi4; and prion protein (PrP), which signals via Gpr126. Extracellular matrix components including laminins and collagens in the basal lamina, which signal via the abaxonal SC receptors (black) β integrins, dystroglycan, and Gpr126. Mechanical forces signal through Piezo1 and 2. The basal lamina is shown surrounding SCs at the immature and all later stages. (This figure is modified from schematics in Jessen and Mirsky 2005 and Fledrich et al. 2019.)

SCs that surround an axon bundle and are contained within the same basal lamina are described as an SC “family” (Peters et al. 1991). Basal lamina deposition (see below) contributes to SC precursor proliferation and survival and coincides with their differentiation into immature SCs. With proliferation, a daughter SC detaches from the “family,” bringing along its associated axon (Webster et al. 1973). When such SCs establish a 1:1 relationship with an axon, they are referred to as “promyelinating” SCs (Fig. 1). These SCs typically ensheath axons with a diameter ≥1 µm that will be myelinated. The recognition and isolation of large-caliber axons destined to be myelinated, away from small-caliber axons that will remain unmyelinated, is referred to as radial sorting (Peters and Muir 1959). As promyelinating SCs transition to myelinating SCs, they turn on Egr2 (also known as Krox20), the master transcription factor of SC myelination that drives expression of myelin protein and lipid biosynthetic genes. The small axons that remain in bundles after radial sorting will be sorted into separate pockets of a single SC, which will differentiate into a mature Remak (nonmyelinating) SC. Remak bundles differ from the axon bundles in developing nerves as all axons are small in caliber and are properly ensheathed by SC processes (for review, see Feltri et al. 2016).

These two alternative modes of axon ensheathment by SCs (i.e., myelination vs. Remak ensheathment) (Peters et al. 1991), are responsible for distinct modes of action potential propagation: saltatory conduction versus continuous propagation, respectively. Accordingly, their associated fibers have distinct physiological functions. Axons that are myelinated include large-caliber (13–20 µm), rapidly conducting type A skeletomotor and proprioceptor fibers and mid-caliber (6–12 µm), fast-conducting type B mechanoreceptors, and preganglionic autonomic efferent fibers. Axons enclosed by Remak SCs are small (typically <1 µm diameter), slowly conducting (0.5–4 m/sec) type C fibers. The latter include efferent, postganglionic autonomic nerve fibers and afferent nerve fibers from the skin that carry pain and temperature information.

Myelinating and Remak SCs are transcriptionally quite distinct and accordingly express different repertoires of proteins. For example, the transcription factor Egr1 is present in Remak SCs, whereas, as noted, myelinating SCs express Egr2 (discussed further below). Myelinating and Remak SCs also differ in the cell-adhesion molecules and receptors that mediate their interactions with the axon and the basal lamina (Harty and Monk 2017; Bosch-Queralt et al. 2023). The most striking difference is the selective and robust expression of myelin proteins by myelinating SCs. In contrast, the P75NTR receptor, L1CAM, and GAP43 are characteristic of Remak SCs. For detailed information on markers present in SCs at different stages of differentiation, including Remak SCs, see Jessen et al. (2015).

Recent single-cell transcriptomic data further delineate markers that are differentially expressed during SC development and between myelin-forming versus Remak SCs (Gerber et al. 2021; Tasdemir-Yilmaz et al. 2021). These studies also highlight heterogeneity within mature SC populations, notably of subgroups of myelin-forming SCs (mSCs) with unique patterns of gene expression (Yim et al. 2022). One such group of mSCs expresses PMP2; these mSCs preferentially ensheath motor axons and are selectively reduced in amyotrophic lateral sclerosis nerve samples (Yim et al. 2022). These results suggest myelin-forming SC subtypes exhibit different characteristics, either due to variations in local, extrinsic signals, and/or possibly distinct developmental origins, that may be relevant to nerve pathology. Future studies, that manipulate or genetically fate-map these SC subtypes based on RNA-seq data will be useful to further elucidate their functional significance.

EXTRINSIC SIGNALS THAT REGULATE SCHWANN CELL DEVELOPMENT

The major extrinsic signals that drive SC differentiation are provided by the axon and the extracellular matrix (ECM)/basal lamina (Salzer 2015; Fledrich et al. 2019; Muppirala et al. 2021; Wilson et al. 2021), as discussed below.

Axonal Signals

NRG1 on the axon controls virtually every aspect of SC development including proliferation, survival, and differentiation of immature SCs, radial sorting, and their binary choice to become Remak versus myelinating SCs (Newbern and Birchmeier 2010). In particular, levels of NRG1 type III, an abundant membrane-tethered NRG isoform, correlate with the ensheathment fate of axons. Axons destined to be myelinated present much higher levels of NRG1 type III than do axons in Remak fibers (Taveggia et al. 2005). Haploinsufficiency or full genetic inactivation of NRG1 type III result in significant PNS hypomyelination (Michailov et al. 2004; Taveggia et al. 2005; Brinkmann et al. 2008), a reduced percentage of myelinated axons, and aberrant axon sorting including in Remak fibers (Taveggia et al. 2005; Raphael et al. 2011). Conversely, overexpression of NRG1 type III results in significant hypermyelination evident as increased myelin thickness (Michailov et al. 2004). Thus, threshold levels of NRG1 type III trigger SC myelination, and above that threshold, the amount of compact myelin that forms is graded to the levels of NRG1 (Nave and Salzer 2006). As NRG1 type III also controls SC proliferation and survival, the elevated levels of NRG1 type III of larger axons drive the generation of the additional SCs required for myelination—thereby coordinating SC numbers to their myelinating fate (Taveggia et al. 2005). Once myelination is complete, ongoing signaling from axonal NRG1 or its SC receptors is not required to maintain the myelin sheath (Atanasoski et al. 2006; Fricker et al. 2011).

NRG1 type III is a juxtacrine signal that activates the ErbB2/3 coreceptors on SCs. ErbB receptors are classical tyrosine kinases whose downstream effectors include the mitogen-activated protein kinase (MAPK) ERK, phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3K), and phospholipase Cγ (PLCγ) pathways, each of which has been reported to be essential for proper myelination (Newbern and Birchmeier 2010; Fledrich et al. 2019). Expression of constitutively active ERK1/2 is able to rescue myelination in NRG1-deficient mice, in part via an increase in protein translation (Sheean et al. 2014).

An additional key axonal signal required for myelination is Adam22 (a disintegrin and metalloproteinase domain-containing protein 22) (Özkaynak et al. 2010; Kegel et al. 2014). Adam22 interacts with leucine-rich glioma-inactivated 4 (Lgi4), a SC-secreted protein that is mutated in the claw paw hypomyelinated mutant mouse (Bermingham et al. 2006). Pan-knockouts of Adam22 (Sagane et al. 2005), neuron-specific deletion of Adam22, and SC-specific deletion of Lgi4 (Özkaynak et al. 2010) all result in major defects in myelination, arresting SCs at the promyelinating stage (Kegel et al. 2013). It is unclear presently how binding of Lgi4 to Adam22 on the axon signals back to SCs to regulate ensheathment and myelination. Interestingly, Lgi4–Adam22 interactions are also required for PNS gliogenesis during early development, including for proliferation of glial-restricted progenitors (Nishino et al. 2010).

Recent studies suggest that axon size, independent of its content of signaling molecules, also regulates SC development and myelination (Park et al. 2022). PIEZO1 and PIEZO2 are among the most abundant mechanosensitive ion channels present in SCs and may regulate myelination via activation of Yes-associated protein 1 (YAP1) and WW domain–containing transcription regulator protein 1 (WWTR1 or TAZ) (Acheta et al. 2022), as discussed further below.

Basal Lamina Signals

Axons direct SC assembly of a basal lamina, which in turn has an essential, autocrine role in SC ensheathment and myelination of axons (Madrid et al. 1975; Bunge et al. 1986; Feltri and Wrabetz 2005; Chernousov et al. 2008). The SC basal lamina is a thin, dense mixture of ECM components, including laminins and collagens, that is closely opposed to the abaxonal SC membrane (Chernousov et al. 2008). The basal lamina is synthesized by SCs with contributions from EFs (Obremski et al. 1993). Conditional knockout of laminin (LAMC1) in SCs results in aberrant basal lamina formation, major defects of axon sorting, and hypomyelination in vivo, underscoring laminin's key role in these events (Chen and Strickland 2003). Both the scaffold-forming and cell-adhesion activities of laminin are required to promote SC ensheathment (McKee et al. 2012). Other components of the matrix, notably the collagens, also regulate SC myelination (Chernousov et al. 2008).

SC laminin receptors include dystroglycan and the α6β1 and α6β4 integrins. These receptors activate downstream signaling pathways (e.g., ILK, FAK, Rac1, cdc42, and PKA) to further induce the formation of the lamellipodia-like processes that interdigitate, contact, and separate axons (Benninger et al. 2007; Nodari et al. 2007; Pereira et al. 2009; Pellegatta et al. 2013; Grove and Brophy 2014). In addition, both collagen and laminin bind to and activate Gpr126/Adgrg6 on SCs, which is required for ensheathment and myelination (Monk et al. 2009). Gpr126 mediates mechano-signals from the ECM (collagen IV/laminin 211) (Paavola et al. 2014; Petersen et al. 2015; Mitgau et al. 2022) to activate adenylate cyclase via Gαs (Mogha et al. 2013), elevating cAMP levels that activate PKA and promote myelination (Howe and McCarthy 2000). Interestingly, the prion protein (PrP) expressed on axons has been implicated as an agonistic ligand for Gpr126 to sustain cAMP levels and myelination in the adult (Küffer et al. 2016). Thus, Gpr126 appears to function as a signal on both the abaxonal (outer) and adaxonal (inner) SC membranes.

MECHANISMS OF SCHWANN CELL ENSHEATHMENT AND RADIAL SORTING

The combined activity of these extrinsic signals drives expansion of the SC plasma membrane and promotes radial sorting and myelination of axons. All of these morphogenetic changes in SCs are driven by remodeling of the actin cytoskeleton. Thus, inhibitors of actin assembly (Fernandez-Valle et al. 1997) and of myosin II contractility (Wang et al. 2008) block axon sorting and myelination in cocultures. Among the master regulators of actin assembly and its organization in cells are Rho, Rac1, and cdc42, foundational members of the Rho GTPase family of proteins (Hall 2012). Each of these have been implicated in regulating radial sorting and SC myelination. Thus, conditional inactivation of Rac1 or Cdc42 during SC development results in impaired axon sorting, delayed myelination, and significant hypomyelination (Benninger et al. 2007; Nodari et al. 2007; Guo et al. 2013). Rho and its key effector Rho kinase (ROCK) promote radial sorting (Pereira et al. 2009) and the coordinated progression of the inner turn of the myelin sheath around the axon at the onset of myelination (Melendez-Vasquez et al. 2004).

Extrinsic signals from the basal lamina and axon, in turn, are known to activate these Rho family GTPases. Rac1 is activated downstream of laminin/β1-integrin signaling (Benninger et al. 2007; Nodari et al. 2007) via lymphoid cell kinase (Lck) (Ness et al. 2013). Indeed, overexpression of Rac1 partially rescues the phenotype of SC conditional β1-integrin nulls (Nodari et al. 2007). In addition, increased activity of the PI3K/Akt pathway, which is downstream of NRG1 signaling (Maurel and Salzer 2000), drives exuberant SC wrapping of axons in Remak and myelinated fibers, and even of collagen fibers, via mTOR signaling and Rac1 activity (Goebbels et al. 2010; Doménech-Estevez et al. 2016). RhoA activation is downstream of Gpr56, which is required for radial sorting (Ackerman et al. 2018) and of laminin signaling via ILK (Montani et al. 2014). Cdc42 is activated in SCs by soluble NRG1 in vitro (Benninger et al. 2007); however, the importance of NRG1 as an activator of Cdc42 in vivo is not yet established.

Effectors of these GTPases include N-WASP (the neuronal Wiskott–Aldrich syndrome protein), which is activated downstream of Rac1 and cdc42 and binds to the Arp2/3 complex to nucleate new branched arrays of actin filaments (Rohatgi et al. 1999; Burianek and Soderling 2013). N-Wasp localizes to the leading edge of SC processes. Pharmacological inhibition of N-WASP impairs myelination in cocultures (Bacon et al. 2007) and conditional inactivation markedly delays sorting and profoundly impairs myelin wrapping, resulting in shorter internodes and thinner sheaths (Jin et al. 2011; Novak et al. 2011). These latter results indicate that elongation and circumferential membrane wrapping share similar actin regulators. Finally, the Rho/ROCK pathway promotes lamellipodia by MLCK (myosin light chain kinase) and activation of Myosin II (Melendez-Vasquez et al. 2004) and, separately, via the actin binding protein Profilin1 (PFN1) (Montani et al. 2014). In the latter case, SC-specific gene ablation of Pfn1 in mice results in profound radial sorting and myelination defects (Montani et al. 2014).

In addition to proteins that promote actin assembly and branching, depolymerization of the actin cytoskeleton is also actively regulated, including by the actin-severing protein Cofilin. NRG1 activates cofilin via the cofilin-phosphatase Slingshot-1, which is activated by translocation to and association with F-actin at the leading edge of lamellipodia (Nagata-Ohashi et al. 2004). In agreement, NRG1 promotes translocation of nonphosphorylated Cofilin to the leading edge of cultured SCs (Sparrow et al. 2012). Further, knockdown of Cofilin1 in cultured SCs blocks their engagement or alignment along axons, their assembly of the basal lamina, and their ability to myelinate (Sparrow et al. 2012).

Taken together, these findings suggest that actin is dynamically and obligately regulated during axon segregation and myelination. Based on ligand activation and localization studies, NRG1/ErbB signaling activates Cofilin and Cdc42 at the leading edge of the adaxonal membrane, leading to a cycle of actin depolymerization and N-WASP-dependent assembly, respectively. These results are consistent with actin treadmilling known to underlie lamellipodial extension in most cells (Pollard and Borisy 2003). In addition, Rac1 and Rho/ROCK act in the abaxonal compartment downstream of laminin/β1-integrin to drive actinomyosin activity required for radial sorting.

MORPHOGENESIS OF THE SCHWANN CELL MYELIN SHEATH

Early electron microscopic studies established that the myelin sheath forms as the result of spiral wrapping of the SC plasma membrane around the axon (Geren 1954; Robertson 1955). The myelin sheath is initially loosely spiraled for the first few turns, then compacts with upregulation of myelin proteins, notably P0 (Kidd et al. 2013). In a classic study, Bunge et al. (1989) provided compelling evidence that spiral wrapping results from circumnavigation of the inner myelin turn around the axon rather than by the countervailing movement of the outer SC compartment that contains the SC nucleus. This model was also suggested by earlier, live imaging studies of developing nerves in tadpole tails (Speidel 1964). Growth of the inner membrane requires it to intercalate between the existing inner membrane and the axon (Fig. 1), disrupting existing interactions between the axon and the SC. Whether movement of the inner turn is primarily guided by interactions with the axon (heterotypic interactions), by interactions with the glial membrane (homotypic interactions), or by both is not yet known.

In addition to wrapping of the inner turn, the myelin sheath also expands at its lateral margins. The innermost turns are initially narrower than the full length of the internode, as suggested by older electron microscopy (EM) studies (Webster 1971) and expands laterally as the sheath matures. This interpretation is strongly supported by changes in the pattern of Caspr staining, which delineates the leading and lateral edges of the SC and transitions from a loose spiral around the axon to its subsequent localization at the paranodes (Pedraza et al. 2001). Thus, morphogenesis of the SC myelin sheath mirrors that of oligodendrocyte myelin (Snaidero et al. 2014; Simons et al. 2023).

New components of the membrane are added in the abaxonal and perinuclear region (Gould 1977), and potentially within the paranodes (Gould and Mattingly 1990). Inserted components—in particular membrane lipids—then likely diffuse throughout the forming myelin sheath. Even after compact myelin has formed, the sheath rapidly expands and its internal circumference continues to increase to accommodate radial expansion of the axon (Webster 1971). Together, these results suggest the myelin sheath conforms to the classical model of a fluid mosaic membrane (Singer and Nicolson 1972; Nicolson and Ferreira de Mattos 2023), with diffusion potentially enhanced by its high lipid content.

REGULATION OF MYELIN SHEATH THICKNESS AND INTERNODE LENGTH

Myelination typically commences around axons that are >1 µm in diameter (Peters et al. 1991), in agreement with theoretical models that suggest myelination enhances conduction velocity in PNS axons of this diameter (Rushton 1951). Together with axon diameter, key determinants of nerve conduction velocity include the thickness of the myelin sheath, its length (i.e., internode length), and the width of the node of Ranvier (Moore et al. 1978; Wu et al. 2012; Arancibia-Cárcamo et al. 2017). These parameters are optimized for their role in saltatory conduction (Waxman 1980).

Accordingly, myelin sheath thickness is tightly regulated by and correlates to the diameter of the axon it surrounds. The relationship between sheath thickness and axon diameter is conventionally expressed as the g-ratio (i.e., axon diameter/total fiber diameter) and is typically ∼0.67 in the PNS. The tight correlation of myelin sheath thickness to axon diameter results from a balance of promyelinating signals with other signals that inhibit/terminate myelination (Salzer 2015; Bolino 2021). As discussed above, the major (promyelinating) determinant of myelin sheath thickness is the level of NRG1 on the axon, which is strongly correlated to axon diameter (Michailov et al. 2004; Taveggia et al. 2005). Recent studies also implicate mechanical signals that are sensitive to axonal diameter and act through ACTL6a, a component of the SWI/SNF chromatin remodeling complex, to determine myelin thickness (Park et al. 2022).

Mechanisms that limit sheath thickness include down-regulation of promyelinating signals (e.g., the NRG1/ErbB signaling pathway). NRG1 activity is itself subject to negative regulation, for example via TACE cleavage (La Marca et al. 2011). In addition, SC expression of ErbB2 (Cohen et al. 1992; Jin et al. 1993) and Akt (Heller et al. 2014; Sheean et al. 2014) is markedly down-regulated with myelination. Furthermore, negative regulators of signaling, for example of the PI3K/Akt/mTOR pathway, also control myelin thickness (Cotter et al. 2010; Noseda et al. 2013; Norrmén et al. 2014). Paradoxically, these inhibitors include components of the basal lamina, which are required for initial SC differentiation, radial sorting, and myelination, but also constrain the amount of myelin that forms, preventing hypermyelination. Laminin interactions in the abaxonal compartment activate Sgk1 (Heller et al. 2014) and inhibit PKA, which is activated by NRG1 (Heller et al. 2014; Ghidinelli et al. 2017), to limit myelination of small fibers. Collagen VI in the basal lamina also limits hypermyelination by regulating FAK, Akt, and Erk signaling (Michailov et al. 2004; Chen et al. 2014).

Internode length is regulated independently of sheath thickness (Tricaud 2018) and NRG1 levels (Michailov et al. 2004). Much of the expansion of the internode during development results from axon elongation dictated by growth of the developing limbs (Hildebrand et al. 1994). Thus, experimental manipulations that elongate limbs increase internode length (Abe et al. 2004; Simpson et al. 2013), whereas those that constrain limb growth during development reduce internode length (Hildebrand et al. 1989; Jacobs and Myers 1993). This linkage may reflect adhesive interactions that bind SCs to axons and mechanotransduction signals that are activated during limb growth (Tricaud 2018). Internode length improves conduction speed until a plateau is reached (Simpson et al. 2013). Conversely, the short internodes characteristic of remyelinated segments (Scherer and Salzer 2001) reduce nerve conduction velocities. Such short, remyelinated segments may result from myelin formation around adult axons that are no longer actively elongating.

Formation of the myelin sheath represents a dramatic expansion of the SC plasma membrane. By some estimates, myelinating SCs may generate up to 20 mm² of membrane around the largest axons—some 2000 times that of a typical epithelial cell (Kidd et al. 2013). The rapid membrane production that occurs during myelination requires coordination of gene transcription, protein translation, directed trafficking and insertion of myelin components to sites of membrane addition, and mechanisms to establish the proper stoichiometry of these myelin components. The mechanisms that coordinate these various cell biological processes during myelination are poorly understood. Production of this huge expanse of membrane imposes significant stresses upon the SCs, including in the ER, rendering them especially vulnerable to mutations that affect the stoichiometry, the folding and/or the trafficking of myelin proteins (Scherer and Svaren 2023).

ORGANIZATION OF THE MYELINATING SCHWANN CELL AND ITS MYELIN SHEATH

Myelinating SCs are radially and longitudinally polarized cells (Salzer 2003; Özçelik et al. 2010; Pereira et al. 2012; Tricaud 2018). With myelination, SCs organize into distinct membrane domains, each with a unique array of proteins, and a communicating set of cytoplasmic compartments (Fig. 2). Longitudinal (axial) polarity is evident by the overall organization of the myelinating SCs and axons into nodal, paranodal, juxtaparanodal, and internodal domains (Rasband and Peles 2023).

Figure 2.

Organization of myelinating Schwann cells (SCs). Schematic organization of myelinating SCs (blue) surrounding an axon (gray); the left cell is shown in longitudinal cross-section and the right cell is shown unwrapped. Myelinating SCs are surrounded by a basal lamina (shown only on the left), which is in direct contact with the abaxonal membrane. The abaxonal compartment contains the SC nucleus (SN) and is divided into Cajal bands and periodic appositions that form between the abaxonal membrane and outer turn of compact myelin. The SC adaxonal membrane is separated from the axonal membrane by the periaxonal space (shown in yellow). Compact myelin is interrupted by Schmidt Lanterman incisures (SLIs), which retain cytoplasm and are enriched in gap and other junctions; a similar autotypic junctional complex of adherens, tight and gap junctions form between the apposed membranes of the paranodal loops. Also shown are the paranodal loops and junctions (red) and the SC microvilli contacting the axon at the node. The axon diameter is reduced in the region of the node and paranodes. (This figure legend is reprinted from Salzer 2015 © Cold Spring Harbor Perspectives in Biology; the left side of the image is adapted, with permission, from Salzer 2003; the right side is adapted, with permission, from Nave 2010.)

Radial polarity is characterized by the distinct compositions of the adaxonal and abaxonal membranes, which lie on either side of the SC—with the compacted membranes of the myelin sheath in between. The adaxonal membrane, which is separated from the axolemma in the internode by a gap of ∼15 nm (the periaxonal space), is enriched in adhesion molecules—the Myelin-associated glycoprotein (MAG) and members of the CADM (Necl) family of proteins (Maurel et al. 2007; Spiegel et al. 2007); the latter are tethered to a 4.1G-based cytoskeleton in the SC. These cell adhesion molecules (CAMs) in the inner SC membrane likely maintain spacing and interactions with the axon during wrapping of promyelinating and myelinating SCs. CAMs that mediate interactions with other domains are discussed in detail elsewhere (Salzer et al. 2008; Rasband and Peles 2023). The adaxonal SC membrane also contains solute carriers that maintain the periaxonal space (Marshall-Phelps et al. 2020) and monocarboxylate transporters that contribute to axo-glial metabolic exchange (Doménech-Estevez et al. 2015). SC receptors for axonal signals (e.g., NRG1 and PrP) are also likely to be expressed in the adaxonal membrane, although this remains to be convincingly demonstrated.

The outer, abaxonal membrane lies adjacent to and interacts with laminin and other components of the basal lamina via integrins (initially via α6β1 and later via α6β4), dystroglycan, and Gpr126. Unlike the inner cytoplasmic compartment, which is uniformly spaced, the outer cytoplasmic compartment is interrupted by periodic appositions between the abaxonal membrane and the outer wrap of the compact myelin sheath (Fig. 2). These appositions are enriched in a dystroglycan complex linked via dystrophin-related protein 2 (Drp2) to periaxin (Sherman et al. 2012); they delineate a network of anastomosing, cytoplasmic channels termed Cajal bands (Court et al. 2004) that depend on expression of periaxin and dystroglycan and signaling from Gpr56 (Court et al. 2004; Ackerman et al. 2018). These cytoplasmic channels provide a conduit for centrifugal transport of RNA and proteins originating in the cell soma en route to the paranodal collar (Gould and Mattingly 1990; Court et al. 2004). Consistent with their proposed role in transport, Cajal bands are enriched in large arrays of cytoskeletal proteins, including microtubules, intermediate filaments, utrophin family members (Court et al. 2009, 2011), and an actin/spectrin complex (Susuki et al. 2011; Kidd et al. 2013; Walko et al. 2013). They also represent sites of localized protein synthesis (Gould and Mattingly 1990), intracellular signaling from laminin receptors (Heller et al. 2014), and based on their enrichment in caveolae (Mugnaini et al. 1977; Köling 1985; Mikol et al. 1999), may function in the uptake of metabolites from the extracellular space.

In between these two membranes is the multilamellar myelin sheath—the result of compaction of the circumferentially wrapped, SC plasma membrane. Myelin provides a high resistance, low capacitance sheath essential for impulse propagation by saltatory conduction. The myelin sheath itself is comprised of 40 or more lamellae (Peters et al. 1991). Electron micrographs of compact myelin reveal interperiod lines, which represent the appositions of the extracellular leaflets, alternating with major dense lines, representing the tight apposition of the cytoplasmic leaflets (Fig. 3).

Figure 3.

Schematic organization of the compact myelin sheath. Myelin forms, initially, as loose wraps of uncompacted membrane. With the onset of myelin transcription, myelin proteins are up-regulated including P0, myelin basic protein (MBP), and PMP22, which are shown diagrammatically. Compaction is mediated by extracellular interactions of P0 tetramers on one membrane interacting with P0 tetramers on the opposing membrane, shown here only as dimer/dimer interactions for simplicity. Compaction of the cytoplasmic leaflets is mediated by electrostatic interactions of MBP with the phospholipid bilayer supplemented by the interactions with the cytoplasmic tail of P0. Compact myelin appears in electron microscopy (EM) as a major dense line (MDL) (representing tight apposition of the cytoplasmic leaflets) alternating with the intraperiod lines (representing the two apposed extracellular leaflets) as shown. (This figure and figure legend is reprinted from Salzer 2015 © Cold Spring Harbor Perspectives in Biology.)

Myelin has a unique composition among plasma membrane equivalents. It has an unusually high lipid content (∼70%), which includes galactosphingolipids, certain phospholipids (correspondingly named sphingomyelin), saturated long-chain fatty acids, and, in particular, cholesterol—which is required for myelin sheath assembly (Saher and Simons 2010). In addition, myelin is highly enriched in relatively few proteins (Fig. 3). Like central nervous system (CNS) myelin proteins (Toyama et al. 2013), these are likely to be quite stable in the mature sheath. The most abundant SC myelin protein by far is P0, which is encoded by the Myelin protein zero gene MPZ. P0 is a transmembrane adhesion molecule of the immunoglobulin superfamily and constitutes nearly half of the protein mass of PNS myelin (Siems et al. 2020). P0 promotes apposition of the extracellular leaflets via homophilic adhesion (Fig. 3; Filbin et al. 1990; Shapiro et al. 1996). Another key set of proteins are isoforms of myelin basic protein (MBP), which constitute ∼20% of myelin protein mass. MBP is a peripheral membrane protein that neutralizes the charges of membrane phospholipids, in particular of phosphatidylserines (Kidd et al. 2013). MBP and the cytoplasmic segment of P0 together promote apposition of the intracellular leaflets to form the major dense line (Martini et al. 1995). Other compact myelin proteins include several tetraspanins, notably PMP22, and periaxin, which are frequently mutated in inherited neuropathies (Scherer and Svaren 2023). Mass spectroscopy of myelin-enriched fractions has identified many additional proteins that are present at much lower levels (Siems et al. 2020). Characterization of these proteins is, as yet, incomplete, including whether they are localized to compact myelin or to other sites in the SC. A number of these have been implicated in the etiology of inherited neuropathies (Siems et al. 2020).

Compact myelin is interrupted by Schmidt–Lanterman incisures (SLIs) (Fig. 2). These are interspersed along the internode and are most abundant in heavily myelinated, large-diameter fibers. SLIs (or clefts) provide a conduit for communication between the inner and outer collars of cytoplasm via the presence of gap junctions, which are arrayed between adjacent membranes (Balice-Gordon et al. 1998). Adjacent membranes in the SLIs also harbor autotypic adherens junctions (Fannon et al. 1995), tight junctions, and a series of PDZ-domain-containing proteins, including the MAGUKs, that are linked to the 4.1G cytoskeleton (Terada et al. 2019) and F-actin (Trapp et al. 1989). SLIs also show enrichment of the tyrosine kinase Src (Terada et al. 2013) and are likely to be a site of autocrine signaling (Heller et al. 2014), consistent with their rich array of cell junctions. Many of the adhesion molecules present in the adaxonal membrane (i.e., MAG and the Necls [Cadms]) are also enriched in the SLIs (Maurel et al. 2007; Spiegel et al. 2007) where they likely contribute to the integrity of these structures. As these clefts develop after the myelin sheath has substantially formed (Small et al. 1987), they are not required for myelin formation, but rather may assist in its maintenance (Gillespie et al. 2000).

In keeping with its polarized organization, components of classical polarity complexes are enriched at distinct sites of myelinating SCs (Masaki 2012; Pereira et al. 2012; Tricaud 2018). The mechanisms that drive the targeting of these complexes to their respective intracellular sites, the regulation of their activity, and their precise roles during myelination in vivo are incompletely understood.

SCHWANN CELLS REGULATE AXON SURVIVAL, DIAMETER, AND TRANSPORT

The interactions between axons and SCs are bidirectional, and SCs have a profound impact on the biology, function, and integrity of axons. As noted, SCs support distinct modes of axon ensheathment and axon conduction. The separation of axons by Remak SCs serves to minimize epiphatic conduction. Myelinating SCs enable saltatory conduction via their elaboration of the myelin sheath and also by their dramatic reorganization of the content and distribution of axonal surface proteins, including the accumulation of ion channels at nodes of Ranvier (Rasband and Peles 2023). SCs also coordinate the injury response by transdifferentiating into repair SCs that promote axonal regeneration in the PNS (Stassart et al. 2023). Here, we briefly consider the role of SCs in supporting axon integrity and regulating axonal diameter and transport.

Both myelin-forming and Remak SCs have an essential role in supporting axon integrity and neuronal survival during early development, postnatally, and in the adult (Harty and Monk 2017; Bosch-Queralt et al. 2023). A fundamental role of SCs is to ensheath axons—a highly conserved function of glia throughout evolution (Nave and Werner 2021). Among the key roles of glial ensheathment is the metabolic support of axons (Simons et al. 2023). Because axons extend a significant distance from their cell bodies—up to 1 meter in humans—they depend on axonal transport, an energy-dependent process, to survive. This renders axons vulnerable, so that axon degeneration is an early event in many neurological diseases. SCs are evenly distributed along the entire length of the axon and are therefore ideally poised to provide local support to axons, both structurally and metabolically. This support is independent from myelination and is believed to rely on the intimate apposition of the SC and axon membranes at various sites. Indeed, similar to oligodendrocytes in the CNS that transfer lactate and pyruvate to axons to support their metabolic needs (Nave and Werner 2021), SCs utilize monocarboxylate transporters to transfer lactate and pyruvate to PNS axons (Doménech-Estevez et al. 2015; Bouçanova and Chrast 2020; Bouçanova et al. 2021; Jia et al. 2021; Deck et al. 2022). Presumably, SCs in close contact with axons sense and respond to their metabolic status. Although the molecular mechanisms that mediate this sensing and energy delivery are still largely unknown, they are the subject of intense investigation.

A striking example of the dependence of axons on SCs is the massive loss of axons and neurons during development that results from depletion of immature SCs due to disruption of the NRG1/ErbB signaling axis (Meyer and Birchmeier 1995; Riethmacher et al. 1997). In addition to loss of metabolic support, these effects may result from loss of trophic signals from immature SCs, which are a rich source of growth factors including neurotrophins (Davies 1998; Scherer and Salzer 2001). Loss of SCs may also impair the targeting and peripheral innervation of axons thereby depriving them of access to their normal, target-derived neurotrophic factors (Birchmeier 2009; Bosch-Queralt et al. 2023).

In addition to prospective metabolic support of axons, myelinating SCs have long been known to promote radial expansion of the underlying axon (Aguayo et al. 1979; Windebank et al. 1985; de Waegh et al. 1992), which in turn increases nerve conduction velocity (Gasser and Grundfest 1939). Radial expansion is locally regulated and is most evident along the internode of large PNS fibers. The local nature of this expansion is evident in the reduced diameter of the axon—underneath regions of noncompact myelin (e.g., SLIs [Price et al. 1993] and in the node and paranodes) where the diameter may be as little as 20% of that of the internode of large axons (Berthold 1996). Reduced diameter of the axon at the node may enhance conduction velocity by reducing the surface area and thereby the capacitance of the nodal axolemma (Halter and Clark 1993; Johnson et al. 2015).

Radial expansion is driven in part by increasing the spacing of the neurofilament (NF) cytoskeleton via increased numbers and phosphorylation of NF subunits (Yuan et al. 2017; see also Dale and Garcia 2012 for an alternate view). In agreement, NF numbers and phosphorylation are increased along the myelinated internode (de Waegh et al. 1992; Cole et al. 1994) and are reduced, less phosphorylated, and more tightly packed at the node (Price et al. 1993; Hsieh et al. 1994). Axon diameter and NF phosphorylation are significantly reduced in dysmyelinating mouse mutants (de Waegh and Brady 1990, 1991; de Waegh et al. 1992; Kirkpatrick and Brady 1994; Brady et al. 1999; Martini 2001). It is not yet known whether myelinating SCs also impact other mechanisms that regulate axon diameter for example, via the submembranous actin cytoskeleton (Leite et al. 2016) and myosin II contractility (Fan et al. 2017; Costa et al. 2018).

Signals on SCs that regulate axon diameter include MAG and CMTM6 (chemokine-like factor-like MARVEL-transmembrane domain-containing family member-6); it is likely that other signals remain to be identified (Nave and Werner 2021). MAG expression modestly increases NF phosphorylation and axon caliber by activating axonal Cdk5 (Yin et al. 1998; Eichel et al. 2020). Another adaxonal SC protein, CMTM6 is up-regulated with myelination and interacts with MAG on the adaxonal membrane. However, CMTM6 constrains the expansion of axons during myelination and is required to prevent overexpansion of axons in adult nerves (Eichel et al. 2020). Surprisingly, NF packing density and phosphorylation are unaltered in the axons of CMTM6 knockouts, suggesting alternate mechanisms drive their expanded axon diameters. Enlarged axons in the Remak fibers of the Cmtm6 knockouts indicate that nonmyelinating SCs also regulate axon diameter.

In addition, SC myelination regulates axonal transport. The rates of slow and fast axonal transport are reduced in the nodal and paranodal regions (de Waegh et al. 1992; Zimmermann 1996; Salzer 2003). Indeed, more than 90% of the membranous organelles of the entire axon accumulate in the nodes and paranodes (Berthold et al. 1993). Reduced transport rates in the nodal region were evident by live imaging organelle transport in myelinated axons of Xenopus laevis (Cooper and Smith 1974) and labeled (fluorescent) NFs in myelinating cocultures (Monsma et al. 2014). It is also consistent with the accumulation of metabolically labeled, axonal glycoproteins at this site (Armstrong et al. 1987). The mechanism(s) responsible for the reduction of transport in the nodal/paranodal region are not known but may result from the constricted axon diameter at this site (Zimmermann 1996) and potentially from alterations of microtubule–motor interactions and organization. Axon–SC interactions at the paranodal junctions are also known to regulate local axonal transport (Zhang et al. 2020; Ishibashi and Baba 2022), although the mechanisms involved remain to be elucidated.

SCHWANN CELLS REGULATE THE ARCHITECTURE OF PERIPHERAL NERVES

SCs are the most abundant cells of the PNS, corresponding to ∼70% of all cells (Stierli et al. 2018). SCs interact with many of the cellular components of peripheral nerves to regulate their development and pathology (see Taveggia and Feltri 2022; Bosch-Queralt et al. 2023 for recent reviews). This includes regulating their organization into various compartments (e.g., the endoneurium, the perineurium, and the epineurium) (Fig. 4). These compartments, with their distinct cellular and ECM compositions, provide critical mechanical and metabolic support for nerves. The cellular components of the endoneurial compartment include bundles of axon–SC units with interspersed EFs and a collagenous ECM (Richard et al. 2012). Remak bundles and myelinated axons form fascicles in the endoneurium that are enclosed by the perineurium, which contributes to the formation of the blood–nerve barrier (BNB) (Rechthand and Rapoport 1987; Iwanaga et al. 2022).

Figure 4.

Architecture of peripheral nerves. Schematic of a peripheral nerve highlighting the three cellular compartments: the epineurium, perineurium, and endoneurium. The perineurium is comprised of layers of perineurial glia (PNG) that surround the endoneurium, which, as shown on the right, contain large axons (orange) that are myelinated (myelin sheath is dark blue), smaller axons (orange) that are ensheathed by Remak Schwann cells (SCs), and scattered endoneurial fibroblasts (EFs) (green). These fascicles are joined together by the epineurium. The vasculature, including small arterioles of the vasa nervorum, is also shown (red). (This figure is adapted from Zotter et al. 2022 and reprinted with permission from the author(s) © 2022.)

SC secretion of the morphogen Desert hedgehog (Dhh) is required during development to form the perineurium and its BNB, the epineurium, and for accumulation of endoneurial collagen (Parmantier et al. 1999; Jessen and Mirsky 2019). Dhh also suppresses the postnatal formation of mini-fascicles that are otherwise generated by EFs (Parmantier et al. 1999; Zotter et al. 2022). The hedgehog-activated transcription factor Gli1 is present in epineurial and perineurial cells, EFs, and pericytes, and regulates endoneurial development (Zotter et al. 2022). Surprisingly, myelinating SCs regulate Gli1 expression independent of Dhh, suggesting other SC signals cooperate with Dhh to control postnatal architecture. Finally, myelinating SCs regulate the formation of the peripheral nerve vasculature via both positive (Mukouyama et al. 2005) and negative (Taïb et al. 2022) signals.

TRANSCRIPTIONAL AND EPIGENETIC REGULATION OF SCHWANN CELL DEVELOPMENT AND MYELINATION

Each step of SC development, from the specification of neural crest cells into SC precursors to their progression through the SC lineage culminating in myelination, is controlled by transcription factors and epigenetic regulators (Fig. 5; Jacob et al. 2011a; Jacob 2015, 2017; Ma and Svaren 2018; Sock and Wegner 2019; Duman et al. 2020a). Epigenetic regulators comprise nucleosome-remodeling complexes, DNA methylation enzymes, histone modifiers, and noncoding RNAs.

Figure 5.

Transcriptional and epigenetic regulation of Schwann cell (SC) development. Schematic representation of the transcriptional (orange hexagons) and epigenetic (pink ovals) regulation of the different stages of SC development from neural crest cells to myelinating SCs is shown. Positive (green arrows) and negative/inhibitory (red arrows) regulatory effects are indicated. Additional regulatory factors, including corepressors (yellow) are also shown. The differentiation into nonmyelinating (Remak) SCs and the maintenance of the myelinating SC transcriptional state in adults are not included in this figure for reasons of simplicity.

Transcriptional Control of SC Specification and Progression to the Immature SC Stage

Many transcription factors are sequentially expressed at different stages of the SC lineage to control the progression of SC development. However, the transcription factor Sox10 is required for each developmental step in the SC lineage (Kuhlbrodt et al. 1998; Britsch et al. 2001; Paratore et al. 2001; Schreiner et al. 2007; Finzsch et al. 2010; Fröb et al. 2012; Weider and Wegner 2017). Therefore, Sox10 can be considered as one of the main drivers of SC identity and differentiation. Although Sox10 is essential for the specification of the SC lineage (Britsch et al. 2001; Paratore et al. 2001), it is present in all neural crest cells, thus additional factors are required. Indeed, the two highly homologous class I histone deacetylases HDAC1 and HDAC2 (HDAC1/2) interact with Sox10 to promote the activation of the Pax3 promoter (Jacob et al. 2014). Pax3 is a transcription factor that acts in synergy with Sox10 to activate the expression of several genes including Sox10 itself (Werner et al. 2007; Wahlbuhl et al. 2012) and the early markers of the SC lineage Fabp7 and P0. In the absence of HDAC1/2 or of Pax3, Fabp7 and P0 are not expressed, Sox10 expression is strongly reduced, and SC precursors are not formed (Jacob et al. 2014). HDAC1/2 and Pax3 are thus essential for the specification of neural crest cells into the SC lineage. In addition to Pax3, Fabp7 and P0, Sox10 also activates the expression of the NRG1 receptor ErbB3 in neural crest cells, which promotes the specification into SC precursors (Britsch et al. 2001; Adameyko et al. 2009; Prasad et al. 2011).

As noted, SC precursors are directly generated from multipotent neural crest cells. They can also originate from an intermediate progenitor, which is already PNS-committed, such as boundary cap cells. In this case, specification into SC precursors occurs in cells where the expression of the transcription factor FoxD3 is maintained (Nitzan et al. 2013). The manipulation of other factors such as the down-regulation of the homeodomain transcription factor Hmx1 or the overexpression of constitutively active Notch intracellular domain (NICD) in neural crest cells leads to reduced neurogenesis in favor of SC precursors (Morrison et al. 2000; Adameyko et al. 2009), suggesting a potential involvement of these factors in regulating the generation of SC precursors. While the transcription factors Sox10, Pax3, FoxD3, and AP-2α continue to be expressed by neural crest cells as they specify into SC precursors, other key neural crest transcription factors such as Sox9, Snail, and Slug are down-regulated (Del Barrio and Nieto 2002; Spokony et al. 2002). Before delaminating from the dorsal neural tube and migrating to their final destination to give rise to different cell types, neural crest cells first need to undergo epithelial-to-mesenchymal transition (EMT) (Theveneau and Mayor 2012). Sox9, Snail, and/or Slug are necessary or synergize with other factors to induce EMT (Locascio et al. 2002; Cheung et al. 2005) but need to be down-regulated in migrating neural crest cells giving rise to most derived lineages including SC precursors (Del Barrio and Nieto 2002; Spokony et al. 2002). Delamination from the dorsal neural tube is also controlled by transcription factors: Slug, Sox9, Sox10, and FoxD3 cooperate to induce a transient cadherin switch from N-cadherin and cadherin 6B to cadherins 7 and 11, which is necessary to trigger delamination (Theveneau and Mayor 2012).

SC precursors retain a high level of multipotency; they can further differentiate into immature SCs, but also into several other lineages (for details see Kastriti et al. 2022). Woodhoo et al. (2009) found high levels of NICD in rat peripheral nerves from the embryonic SC precursor stage until the immature SC stage at birth. In this study, the authors show that canonical Notch signaling is not required for the specification into SC precursors but is critical for differentiation of SC precursors into immature SCs; however, Notch effectors prevent further differentiation of the lineage (Woodhoo et al. 2009). In contrast, AP-2α, which persists as neural crest cells progress to SC precursors, is down-regulated in immature SCs and has been shown to repress the progression of the lineage into immature SCs (Stewart et al. 2001).

Gene Regulatory Networks Control the Transition to Promyelinating and Myelinating Stages

The last steps of the differentiation process, namely, differentiation into promyelinating SCs and then into myelinating and nonmyelinating Remak SCs are controlled by several transcription factors, among which Sox10, Egr2, Pou3f1, and Brn-2 hold key functions (Topilko et al. 1994; Bermingham et al. 1996; Jaegle et al. 1996, 2003). These transcription factors are interconnected: Sox10 activates the transcription of Pou3f1 and Brn-2 (Jalagur et al. 2011), which in turn synergize with Sox10 to activate the transcription of Egr2, the master transcription factor of myelination (Murphy et al. 1996; Ghislain and Charnay 2006; Reiprich et al. 2010). Egr2 then synergizes with Sox10 to activate the transcription of myelin protein genes, lipid biosynthesis genes, and genes involved in the formation of the node of Ranvier (Topilko et al. 1994; Leblanc et al. 2005; Saur et al. 2021). Together, these feedforward transcriptional mechanisms promote radial sorting and are essential for SC myelination. Other factors are also prominently involved in the regulation of these processes. Indeed, upon mechanical stimuli, the Hippo pathway effectors—the transcription factor Tead and its coactivators Yap and Taz—induce expression of laminin receptors, which are necessary for radial sorting and myelination (Poitelon et al. 2016; Deng et al. 2017). Simultaneously, the nuclear actin-related protein ACTL6A, which is part of several chromatin-remodeling complexes such as the SWI/SNF complex, acts as a sensor for large-caliber axons and ensures their radial sorting and myelination by inducing the derepression of ECM-related genes to produce SC basal lamina and of promyelinating factors such as Pou3f1 (Park et al. 2022). In addition, the transcription factors Yy1, Nfatc3, and Nfatc4 and the effectors Yap and Taz contribute to the induction of Egr2 expression (Kao et al. 2009; He et al. 2010; Grove et al. 2017; Weintraub et al. 2017).

As noted, a striking feature of the transcriptional phenotype of SCs is that they can adopt alternate fates as a myelinating versus Remak SC depending on extrinsic signals. This binary distinction is reinforced via transcriptional cross-repression in which Sox2 and c-Jun, two major inhibitors of myelination, inhibit Egr2 expression and vice versa (Le et al. 2005; Parkinson et al. 2008). Maintenance of the myelinating phenotype requires ongoing expression of both Egr2 and Sox10; conditional inactivation of either results in dedifferentiation, which is particularly rapid in the case of Egr2 and is associated with Sox2 up-regulation (Decker et al. 2006; Bremer et al. 2011). These results indicate that Sox10 and Egr2 are subject to ongoing turnover in mature myelinating SCs and provide the framework for their differentiation into repair SCs following injury (Stassart et al. 2023).

Regulation of the Transcriptional Program of Myelination by Epigenetic and Posttranslational Modifications: HDAC-Dependent Mechanisms

Epigenetic regulation of transcription plays a crucial role in PNS myelination (see Fig. 5 for summary; Pereira et al. 2012). Among the best-characterized modifiers are the HDACs. HDAC1 and HDAC2 are highly homologous and can compensate for the loss of each other; however, they have primary functions when both are expressed. HDAC2 interacts with Sox10 to protect it from degradation and maintain it on its target genes (Duman et al. 2020b) and recruits KDM3A and JMJD2C, two histone H3K9 demethylases, to remove repressive H3K9 methylation marks on Sox10-target genes (Brügger et al. 2017). HDAC1 prevents the activation of β-catenin in early postnatal SCs to protect SCs from apoptosis. Once SCs have entered the myelinating stage, HDAC1 is down-regulated and levels of active β-catenin (ABC) increase, up-regulating Sox10 and Egr2. In turn, Sox10 increases the levels of ABC and Egr2. This feedforward loop enhances the myelination process (Jacob et al. 2011b) and is likely necessary for timely myelination (Lewallen et al. 2011; Tawk et al. 2011). Sox10 also recruits the nucleosome remodeling complex BAF/BRG1 and interacts with HDAC1/2 to activate the transcription of Pou3f1 and Egr2 and of myelin proteins such as P0 (Jacob et al. 2011b; Weider et al. 2012). In addition, Sox10 interacts with HDAC1/2 to activate its own expression (Jacob et al. 2011b). NF-κB has also been shown to promote Pou3f1 expression (Nickols et al. 2003) and to recruit the BAF/BRG1 complex and HDAC1/2 on the Sox10 promoter to activate Sox10 expression (Chen et al. 2011; Limpert et al. 2013); however, ablation of NF-κB in SCs leads to a minimal phenotype (Morton et al. 2013). Therefore, this NF-κB-dependent mechanism is not essential for myelination. In adult SCs, HDAC1/2, presumably through their interaction with Sox10, are necessary to maintain high levels of P0, the integrity of paranodes and nodes of Ranvier, and optimal myelination (Brügger et al. 2015). Two other HDACs, HDAC3 (class I) and HDAC4 (class II), are also involved: HDAC4 has been shown to recruit the repressor complex NcoR1 together with HDAC3 upon cAMP signaling to silence the Jun promoter and thereby favor myelination (Gomis-Coloma et al. 2018). However, two other studies on the functions of HDAC3 in SCs show a hypermyelination phenotype in the absence of HDAC3, which can appear somewhat contradictory to a function of HDAC3 in silencing the Jun promoter. One of these two studies proposes that HDAC3 replaces HDAC2 at the transition between developmental myelination and myelination maintenance to slow down the activation of the P0 promoter and the production of myelin while still maintaining myelin homeostasis (Rosenberg et al. 2018). The replacement of HDAC2 by HDAC3 may, however, only be partial since HDAC1/2 are necessary in adult SCs for the maintenance of appropriate levels of P0 (Brügger et al. 2015).

Mechanisms Related to Other Epigenetic Factors

Similar to Sox10, Egr2 recruits histone-modifying enzymes to promote activation of its target genes. Indeed, Egr2-mediated recruitment of the Rnf40-containing E3 ligase leads to histone H2B monoubiquitination of Egr2 target genes. This results in high expression levels of myelin and lipid biosynthesis genes and repression of inhibitors of myelination, which appears essential for the myelination process (Wüst et al. 2020). In addition, the CCCTC-binding factor (CTCF) that mediates chromatin looping contributes to the induction of Egr2 expression and thereby to the transition from the promyelinating to the myelinating stage (Wang et al. 2020).

The NuRD remodeling complex, which comprises the ATPases CHD3 or CHD4 and recruits HDAC1/2, is also required for radial sorting and myelination. The transcription factor Zeb2 recruits the NuRD complex at the immature SC stage to repress the transcription of the inhibitors of differentiation Sox2, Hey2, and Ednrb and thereby favors differentiation into the promyelinating and then to the myelinating stages (Quintes et al. 2016; Wu et al. 2016). In addition, at late developmental myelination stages, Egr2 interacts with the transcriptional repressors NAB1/2, which recruit the NuRD complex to silence the expression of other inhibitors of myelination including Id2, Id4, and Jun (Parkinson et al. 2004; Srinivasan et al. 2006; Mager et al. 2008; Hung et al. 2012). Pou3f1 and Sox2 are also known to be repressed by Egr2 (Zorick et al. 1999; Le et al. 2005; Parkinson et al. 2008), a mechanism likely occurring through the interaction of Egr2 with NAB1/2/NuRD (Srinivasan et al. 2006; Mager et al. 2008). Indeed, Pou3f1 is an intermediate activator of myelination; it is first required to induce the up-regulation of Egr2 but needs to be down-regulated afterward for myelination to proceed (Ryu et al. 2007). In addition to the Egr2-mediated down-regulation of Pou3f1 and Sox2, the chromatin-remodeling complex Tip60/Ep400, which replaces the histones H2A.Z and H3.1 by H2A and H3.3, respectively, is necessary for the timely down-regulation of Pou3f1, Sox2, and other immature SC markers such as AP-2α, Pax3, Sox1, and Sox3. Tip60/Ep400 is also required for terminal differentiation of myelinating and nonmyelinating SCs (Fröb et al. 2019).

To achieve optimal myelination, the regulation of DNA methylation in specific genes is required. During the myelination process, DNA methyltransferases and the methyl donor SAMe are down-regulated, whereas Gadd45a, Gadd45b, and Apobec1, which promote DNA demethylation, and glycine N-methyltransferase, which transforms SAMe into S-adenosylhomocysteine and N-methylglycine, are up-regulated (Varela-Rey et al. 2014). These regulations are necessary for appropriate expression of myelin-related genes and myelin thickness (Varela-Rey et al. 2014).

Finally, microRNAs (miRNAs) have been implicated in regulating SC biology. miRNAs are single-stranded, small noncoding RNAs that can regulate gene expression by inducing the degradation of their target mRNAs. They are processed from hairpin-containing primary transcripts into precursor miRNAs by the microprocessor proteins Drosha and DGCR8, and then into miRNA duplexes by Dicer (Suster and Feng 2021). miRNAs have a critical function in the regulation of radial sorting and myelination and in the maintenance of myelin integrity (Bremer et al. 2010; Pereira et al. 2010; Verrier et al. 2010; Yun et al. 2010; Gokey et al. 2012; Gökbuget et al. 2015, 2018; Lin et al. 2015). Indeed, these studies show that miRNAs down-regulate immature SC factors and prevent inappropriate expression of injury-induced genes in developing and mature SCs.

In addition to promyelinating transcription mechanisms, there are also inhibitory mechanisms that ensure optimal myelin sheath thickness and prevent pathological hypermyelination. Repressive H3K27 trimethylation marks (H3K27me3) have been implicated in these mechanisms. For instance, a nuclear variant of ErbB3 binds to the DNA to induce enrichment of H3K27me3 at the promoter of myelin-related genes, probably through activation of the histone methyltransferase EZH2 (Ness et al. 2016). In addition, the PRC2 complex, which contains EZH2, represses the promoter of Igfbp2 and thereby prevents Igfbp2-induced activation of Akt, which is an activator of the myelination process (Ma et al. 2015).

Wrapping Up and Looking Forward

There has been substantial progress in the identification of axonal, basal lamina, and mechanotransduction signals that converge on SCs to regulate their development and differentiation. The axon–SC unit functions within the context of the much larger ecosystem of the PNS. There is increasing appreciation that SCs help to coordinate and direct peripheral nerve development. Factors from axons drive SC differentiation, which in turn signal to other cells to direct the development of nerve compartments and vasculature. The best studied of such axon-regulated SC signals is Dhh, but myelinating SCs likely produce other signals that drive morphological changes during differentiation, including Gli1 activation in EFs (Zotter et al. 2022). SCs also contribute to the development and function of other structures and organs, such as the skin, synapses, and the vascular, enteric, and immune systems. Thus, in addition to the myriad of primary peripheral neuropathies, SC pathology is also relevant to cancer, pain, enteric, and dermatological diseases (Taveggia and Feltri 2022).

The extent of reciprocal signaling from other peripheral nerve cell types and compartments back to the axon–SC unit is less well understood. Recent transcriptomic data indicate that a significant number of candidate ligands and their cognate receptor mRNAs are reciprocally expressed by EFs and SCs under homeostatic conditions (Peng et al. 2020; Toma et al. 2020). These include well-characterized growth factors such as nerve growth factor, hepatocyte growth factor, and NRG1 in fibroblasts versus PDGFA and EGF in SCs—in addition to many novel ligands with unknown roles in the PNS. Two candidate factors secreted by fibroblasts that may impinge directly on SCs include the vasodilator peptide adrenomedullin, which was previously shown to up-regulate cAMP expression in SCs (Dumont et al. 2002), and BMP7, previously shown to inhibit myelin gene expression in SCs (Liu et al. 2016). These data highlight the prospective richness of intercellular signaling that may coordinate peripheral nerve development.

The trophic and metabolic support of axons by immature and mature SCs is well known. However, the precise mechanisms involved remain to be established. Given that this trophic support of axons may be relevant to the pathology in primary axonal diseases and to neuropathies secondary to demyelination, elucidating the mechanisms involved has significant clinical implications.

Recent transcriptomic data has identified unexpected heterogeneity within SCs that may underlie the different sensitivities of SCs and the resultant clinical presentations of peripheral nerve disease. Understanding the functional significance of SC diversity revealed by recent scRNA-seq studies is an emerging area with important implications for PNS functions in health and disease.

Finally, more work is needed to elucidate how transcription and epigenetic factors are regulated and to better understand the mechanisms these factors control. The integration of these mechanisms into dynamic regulatory networks promises to greatly diversify the possibilities of therapeutic interventions for developmental disorders, for the maintenance of peripheral nerve integrity in adults during aging, and to promote regeneration after injury and pathology.

IN MEMORIUM: LAURA FELTRI

As this article was going to press, Laura Feltri, our co-author and beloved colleague, passed away after a long illness. Trained as a neurologist, Laura focused her career on and made seminal contributions to elucidating the biology and pathobiology of Schwann cells and peripheral nerves. Her many transformative scientific contributions will of necessity be described elsewhere. Together with Larry Wrabetz, her husband and frequent collaborator, she established major centers of PNS research in Milan and Buffalo where they trained many current and next-generation leaders in the field. Despite her illness, Laura courageously assumed key leadership positions, authored a series of major reviews, and completed some of the most important research of her career. We are impoverished by her loss. We dedicate this article to her memory.