The “Sweet” Side of Ion Channels

Rev Physiol Biochem Pharmacol (2014) 167: 67–114 DOI: 10.1007/112_2014_20 © Springer-Verlag Berlin Heidelberg 2014 Published online: 20 September 2014

Joanna Lazniewska and Norbert Weiss

Abstract Ion channels play a crucial role in cell functioning, contributing to transmembrane potential and participating in cell signalling and homeostasis. To fulﬁl highly specialised functions, cells have developed various mechanisms to regulate channel expression and activity at particular subcellular loci, and alteration of ion channel regulation can lead to serious disorders. Glycosylation, one of the most common forms of co- and post-translational protein modiﬁcation, is rapidly emerging as a fundamental mechanism not only controlling the proper folding of nascent channels but also their subcellular localisation, gating and function. More- over, studies on various channel subtypes have revealed that glycosylation represents an important determinant by which other signalling pathways modulate channel activity. The discovery of detailed mechanisms of regulation of ion channels by glycosylation provides new insights in the physiology of ion channels and may allow developing new pharmaceutics for the treatment of ion channel-related disorders.

Keywords Ion channel • N-linked glycosylation • O-linked glycosylation • Glycan • Protein glycosylation

Contents 1 Introduction ...... 69 2 Protein Glycosylation in a Nutshell ...... 70 3 Means to Investigate Glycosylation of Ion Channels ...... 72 3.1 Identiﬁcation of N-Linked Glycosylation ...... 72 3.2 Identiﬁcation of O-Linked Glycosylation ...... 73 4 Glycosylation-Dependent Regulation of Ion Channels ...... 73

J. Lazniewska and N. Weiss (*) Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic, Prague, Czech Republic e-mail: [email protected]

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4.1 Voltage-Gated Potassium Channels (Kv)...... 73 4.2 Voltage-Gated Sodium Channels (Nav)...... 80 4.3 Voltage-Gated Calcium Channels (Cav)...... 82 4.4 Hyperpolarisation-Activated Cyclic Nucleotide-Gated Channels (HCN) ...... 83 4.5 Transient Receptor Potential Channels (TRPs) ...... 84 4.6 Acid-Sensing Ion Channels (ASICs) ...... 88 4.7 Two-Pore Domain K+ Channels (TASK) ...... 89 4.8 Two-Pore Channels (TPCs) ...... 90 4.9 Pannexin Channels (Panx) ...... 91 4.10 CFTR Channel ...... 92 4.11 CLC Channels ...... 93 5 Crosstalk Between Glycosylation and Other Signalling Pathways ...... 93 5.1 Regulation by Auxiliary Subunits ...... 93 5.2 Klotho-Dependent Regulation of TRPV5 ...... 97 5.3 SP1 Induces CLC-2 Gene Expression ...... 97 6 Modulation of Ion Channel Glycosylation by External Factors ...... 97 6.1 Modulation by External Sugar ...... 97 6.2 Modulation by Temperature ...... 98 7 Glycosylation-Mediated Channelopathy ...... 99 7.1 Long QT Syndrome ...... 99 7.2 Diabetic Neuropathic Pain ...... 100 7.3 Cystic Fibrosis ...... 100 7.4 Varitint–Waddler Phenotype ...... 101 Conclusion and Perspectives ...... 101 References ...... 103

Abbreviations

ASIC Acid-sensing ion channel B35 Neuroblastoma cell CAD Cath.a-differentiated cell Cav Voltage-gated calcium channel CDG Congenital disorders of glycosylation CFTR Cystic ﬁbrosis transmembrane conductance regulator CHO Chinese hamster ovary cell ER Endoplasmic reticulum ERGIC ER–Golgi intermediate compartment GalNAc N-acetylgalactosamine GlcNAc-1-P N-acetylglucosamine-1-phosphate HCN Hyperpolarisation-activated cyclic nucleotide-gated channel K2P Two-pore domain potassium channel Kv Voltage-gated potassium channel LacNAc N-acetyllactosamine LQT Long QT syndrome M1 Murine cortical collecting duct Nav Voltage-gated sodium channel O-GalNAc Mucin-type O-glycan

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Panx Pannexin channel PNGase F Peptide-N-glycosidase F Sf9 Spodoptera frugiperda cell SKBR3 Human breast cancer cell ST3Gal4 Beta-galactoside alpha-2,3-sialyltransferase 4 ST8sia2 Alpha-2,8-sialyltransferase 2 TPC Two-pore channel TRIP8b Tetratricopeptide repeat-containing Rab8b-interacting protein TRP Transient receptor potential channel TRPC Transient receptor potential canonical channel TRPM Transient receptor potential melastatin channel TRPP Transient receptor potential polycystin channel TRPV Transient receptor potential vanilloid channel α-benzyl- 1-Benzyl-2-acetamido-2-deoxy-α-D-galactopyranoside GalNAc

1 Introduction

Ion channels are pore-forming integral membrane proteins whose main function is controlling the flow of ions across cell membranes. They are multimeric protein complexes expressed both in the plasma membrane and intracellular organelles and are key contributors to the cell “well-being” (Hille 2001). It is well established that ion channels undergo chemical and post-translational modifications including acet- ylation (Shipston 2014), phosphorylation (Davis et al. 2001) (Ismailov and Benos 1995) (Levitan 1994), methylation (Baek et al. 2014) (Palmieri et al. 2012), ubiquitination (Altier et al. 2011) (Abriel and Staub 2005) (Kantamneni et al. 2011) (Marangoudakis et al. 2012) (Waithe et al. 2011), SUMOylation (Rougier et al. 2010) (Rajan et al. 2005) (Dai et al. 2009) or palmitoylation (Shipston 2011), a fundamental process controlling their expression, activity and cellular function. Recently, protein glycosylation, a process commonly known to contribute to the endoplasmic reticulum (ER) quality control of nascent polypeptide chains and critical for the proper folding, sorting and trafficking of proteins that enter the secretory pathway (Weerapana and Imperiali 2006) (Roth et al. 2010) (Moremen et al. 2012), has been also shown to play a fundamental role in modulating ion channel function (Ednie and Bennett 2012). Using site-directed mutagenesis of glycosylation loci, pharmacological inhibition of protein glycosylation and enzymatic removal of glycans, it was shown that preventing protein glycosylation drastically alters life cycle of ion channels (Gong et al. 2002) (Mant et al. 2013) (Weiss et al. 2013) and most importantly their intrinsic activity (Jing et al. 2012) (Pertusa et al. 2012). Moreover, glycosylation at specific channel loci appears to produce variable effects on a given channel, contributing to fine-tune channel activity and fulfil highly specialised cellular functions. Interestingly enough, glycosylation of ion channels also represents a fundamental regulatory

[email protected] 70 J. Lazniewska and N. Weiss platform by which various signalling pathways modulate channel activity (Leunissen et al. 2013) (Ding et al. 2014) (Laedermann et al. 2013). The network of interactions underlying glycosylation-related regulation of ion channels is grad- ually being discovered. This is of extremely high importance considering that dysregulation of ion channels gives rise to numerous human disorders, either congenital or acquired, the so-called channelopathies (Weiss and Koschak 2014) (Kim 2014), and that the list of inborn defects resulting from defective protein glycosylation (congenital disorders of glycosylation, CDG) is still growing (Scott et al. 2014). The focus of this review is to provide a comprehensive understanding of our current knowledge of the role of glycosylation in ion channel function and physiology.

2 Protein Glycosylation in a Nutshell

The biochemical and enzymatic basis of protein glycosylation has recently been extensively reviewed elsewhere (Moremen et al. 2012), and thus we only provide here a succinct overview. Glycans are built from nine monosaccharides – fucose, galactose, N-acetylgalactosamine (GalNAc), glucose, N-acetylglucosamine (GlcNAc), glucuronic acid, mannose, sialic acid and xylose (Fig. 1a). Glycosylation is the enzymatic process mediated by various glycosidases and glycosyl- transferases, as opposed to glycation. Proteins can be O- and N-linked glycosylated. In the case of O-glycosylation, the oligosaccharide is attached to the protein via an oxygen atom on a serine (S) or threonine (T). In contrast, N-glycosylation involves addition of saccharides to an asparagine (N) located within a canonical sequence N-X-S/T (X, any amino acid except a proline) (Schwarz and Aebi 2011). N-glycosylation as well as several types of O-glycosylation (O-mannose, O-fucose, O-glucose and O-galactose) (Van den Steen et al. 1998) begins in the ER and is continued throughout the Golgi apparatus, while attachment of mucin-type O-glycan (O-GalNAc) occurs only in the Golgi apparatus (Bennett et al. 2012) (Ohtsubo and Marth 2006). Oligosaccharides that are destined for attachment to asparagine of nascent polypeptide chain are first anchored to the lipid molecule dolichol phosphate facing the cytoplasmic side of the ER. First, a saccharide, which consists of two GlcNAc and five mannose residues, is enzymatically added to the dolichol phosphate. Next, this structure “flips” to the lumen side of the ER where various enzymes add more monosaccharides to the already existing glycan tree. Such precursor is then trans- ferred onto the asparagine residue of the polypeptide chain. Before leaving the ER, three glucose residues are removed from the oligosaccharide, and this is part of the quality control “checkpoint” of the ER. After entering the Golgi apparatus, the glycan tree is further modified to form more complex oligosaccharides and is usually terminated with sialic acid moieties. All N-glycans, however, posses a common sugar core comprising two GlcNAc and three mannose residues. The

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Fig. 1 N-linked oligosaccharides. (a) Chemical structure of the nine monosaccharides used by mammalian cells to build N-glycans. (b) Schematic representation of the three main types of N-linked glycan structures. Glycosidases commonly used to strip off the glycan at various levels are also shown. PNGase F cleaves between the innermost GlcNAc and the asparagine residue of complex, hybrid and high-mannose oligosaccharides. In contrast, endoglycosidase H (Endo H) cleaves within the chitobiose core of high-mannose and some hybrid oligosaccharides, leaving a GlcNAc residue bound to the asparagine. Neuraminidase catalyses the hydrolysis of α2-3-, α2-6- and α2-8-linked N-acetyl-neuraminic acid residues and is classically used in mammalian cells to remove the sialic acid moiety main types of N-glycan include high-mannose, hybrid and complex oligosaccharide trees (Fig. 1b) (Weerapana and Imperiali 2006). The most common type of O-linked glycosylation is the so-called mucin type, which involves addition of GalNAc to the hydroxyl group of a serine or threonine and takes place in the Golgi apparatus (Bennett et al. 2012). The process is mediated by UDP-N-acetylgalactosamine and polypeptide N-acetylgalactosaminyltransferase

[email protected] 72 J. Lazniewska and N. Weiss enzymes (Tian and Ten Hagen 2009). It is noteworthy that O-glycosylation can overlap with protein phosphorylation at the same serine or threonine residues (Hart et al. 2011; Wang et al. 2012).

3 Means to Investigate Glycosylation of Ion Channels

3.1 Identiﬁcation of N-Linked Glycosylation

Various enzymatic and pharmacological tools can be used to prevent the formation of N-glycans or, in contrast, to remove N-glycans a posteriori, allowing investiga- tion of N-glycan formation and function at different stages of the process (Fig. 1b). For instance, tunicamycin, a nucleoside antibiotic, blocks the very first step of N-glycosylation by preventing the transfer of N-acetylglucosamine-1-phosphate (GlcNAc-1-P) from UDP-GlcNAc to dichol-P (Heifetz et al. 1979), generating non-glycosylated proteins. Peptide-N-glycosidase F (PNGase F), an amidase that cleaves between the innermost GlcNAc and the asparagine residue of high- mannose, hybrid and complex oligosaccharides from N-glycoproteins, is usually used to strip off the entire N-glycan a posteriori, either in vitro or in vivo (Tarentino et al. 1985). In contrast, endoglycosidase H cleaves within the chitobiose core of high-mannose and some hybrid oligosaccharides, leaving a GlcNAc residue bound to the asparagine (Freeze and Kranz 2010). Finally, acetyl-neuraminyl hydrolase (sialidase), commonly known as neuraminidase, catalyses the hydrolysis of α2-3-, α2-6- and α2-8-linked N-acetyl-neuraminic acid residues and is classically used in mammalian cells to remove the sialic acid moiety (and corresponding negative charge) from glycoproteins and oligosaccharides (Schauer 1982). A broader spec- trum of chemicals and enzymes used to manipulate protein N-glycosylation is also presented elsewhere (Hagglund et al. 2007; Esko and Bertozzi 2009). Despite the fact that pharmacological and enzymatic disruption of N-glycan is a valuable approach, other methods can be used to exclude nonspecific modifications of cellular N-glycans per se rather than from specific deglycosylation of the studied channel. Hence, site-directed mutagenesis is commonly used to disrupt canonical glycosylation sites (N-X-S/T) by replacing asparagine (N) residue either with glutamine (Q), glycine (G), alanine (A) or threonine (T), although glutamine is most commonly used because of its structural similarity with asparagine residues, differing only by one methyl group in the amino acid side chains, which is consequently expected to preserve the local charge distribution within the protein and the secondary structure of the channel. Finally, a broad panel of mammalian cell lines presenting alterations in the glycosylation of proteins and lipids have been isolated and can be used to charac- terise glycosylation of ion channels that elucidate specific functional roles of glycan structures on channel activity reintroduced in those cells (Patnaik and Stanley 2006). Molecular characterisation of the glycan structure can also be further

[email protected] The “Sweet” Side of Ion Channels 73 analysed by liquid chromatography and mass spectrometry (Wuhrer et al. 2005a, b; Morelle and Michalski 2007).

3.2 Identiﬁcation of O-Linked Glycosylation

Although the analysis of protein O-linked glycosylation may be more challenging, different approaches have been described (Calvete and Sanz 2008; Zauner et al. 2012). A relatively simple approach to interfere with the enzymatic attachment of O-glycan on the polypeptide chain is the application of 1-benzyl-2- acetamido-2-deoxy-α-D-galactopyranoside (α-benzyl-GalNAc), a competitive inhibitor of O-GlcNAc transferase (Zanetta et al. 2000). A more sophisticated way to reveal O-linked glycosylation of ion channels includes the application of azide monosaccharide analogues, which can be detected by a reaction called Staudinger ligation (Hang et al. 2003), or the use of specific antibodies binding to the glycan moiety (Comer et al. 2001). In contrast to N-glycosylation, no canonical consensus site for protein O-glycosylation has been identified. However, predictive tools using experimentally verified O-glycosylation site entries and comparing the surrounding amino acid sequence similarity have been used successfully to char- acterise potential new O-glycosylation sites (Thanka Christlet and Veluraja 2001; Aoki-Kinoshita 2013). Indeed, a profound analysis of O-glycosylation sites validated experimentally revealed an increase in proline residues in the proximity of O- glycosylated serine and threonine. Moreover, in the case of mucin-type glycosylation, acidic amino acids were shown to be preferred around serine and threonine residues. In contrast, aromatic amino acids like cysteine, or amino acids with large side chains, have been shown to disfavour O-glycosylation. Hence, several amino acid motifs seem to especially undergo O-glycosylation and include T-A-P-P, T-V- X-P, S/T-P-X-P and T-S-A-P (Thanka Christlet and Veluraja 2001).

4 Glycosylation-Dependent Regulation of Ion Channels

4.1 Voltage-Gated Potassium Channels (Kv)

Sequence analysis of the human cardiac pore-forming subunit of voltage-gated inwardly rectifying potassium channel Kv11.1 (hERG), often associated with cardiac arrhythmias, identified two potential N-glycosylation sites (N598 and N629) (Fig. 2, Table 1). Although some discrepancies exist between experimental studies, likely due to intrinsic specificities in glycosylation pathways inherent to cellular expression systems, functional examination of glycosylation-deficient channels revealed that glycosylation at asparagine N598 contributes to the steady-state expression of the channel at the plasma membrane, possibly by stabilising and/or

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Fig. 2 Schematic representation of the location of validated N-glycosylation sites in the scheme of voltage-gated potassium channel (Kv) structure. The corresponding residues in the human channels are indicated in brackets promoting channel trafficking (Petrecca et al. 1999; Gong et al. 2002). In addition, preventing glycosylation at asparagine N629 drastically disrupts channel trafficking to the plasma membrane. However, it is not fully clear if this effect relies on the absence of glycosylation per se or results from a nonspecific alteration of the neighbouring channel structure. Experimental electrophysiological recordings of glycosylation-deficient Kv11.1 reexpressed in CHO cells indicate that proper glycosylation is also critical for channel gating (Norring et al. 2013) (Du et al. 2014). Based on in silico modelling, reduced channel glycosylation was shown to shorten the repolarisation phase of cardiac action potentials, thereby influencing cardiac

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Table 1 Ion channels identiﬁed in glycosylation studies Glyco. site/ Channel Cell type (eq. human) Location Functional role References

Voltage-gated potassium channels (Kv)

Kv1.1 (rat) CHO N207 (N207) S1–S2 Channel gating; Watanabe linker no inﬂuence on et al. (2003), channel surface Watanabe trafﬁcking or et al. (2004) stability

Kv1.2 (rat) CHO N207 (N207) S1–S2 Channel gating Watanabe linker and surface traf- et al. (2007) ﬁcking; no inﬂuence on channel stability

Kv1.3 (rat) CHO, CAD N229 (N279) S1–S2 Channel gating Zhu linker and surface et al. (2012) trafﬁcking

Kv1.4 (rat) CHO N354 (N352) S1–S2 Channel surface Watanabe linker trafﬁcking and et al. (2004), stability; no Schwetz inﬂuence on et al. (2010) channel gating

Kv1.5 (rat) CHO N290 (N299) S1–S2 Channel gating Schwetz linker et al. (2010)

Kv2.1 (rat) CHO, CHO Glycosidase ND Channel gating Schwetz Pro-5/Lec2 et al. (2011)

Kv3.1b (rat) CHO, Sf9, N220 (N220) S1–S2 Channel gating Brooks B35 N229 (N229) linker and cell–cell et al. (2006), S1–S2 border targeting Hall linker et al. (2011), Hall et al. (2013)

Kv4.2 (rat) CHO, CHO Glycosidase ND Channel gating Schwetz Pro-5/Lec2 et al. (2011)

Kv4.3 (rat) CHO, CHO Glycosidase ND Channel gating Schwetz Pro-5/Lec2 et al. (2011)

Kv11.1 HEK293, N598 S5–S6 Channel gating, Petrecca (hERG) CHO, in silico N629 linker surface trafﬁck- et al. (1999), (human) modelling S5–S6 ing and stability Gong linker et al. (2002), Du et al. (2014)

Voltage-gated sodium channels (Nav)

Nav (electric Lipid bilayer Glycosidase ND Channel gating Recio-Pinto eel) et al. (1990), Cronin et al. (2005)

Nav1.4 (rat) CHO, Glycosidase ND Channel gating Bennett HEK293 et al. (1997), Zhang et al. (1999) (continued)

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Table 1 (continued) Glyco. site/ Channel Cell type (eq. human) Location Functional role References

Nav1.5 CHO, Glycosidase ND Channel gating Bennett (human, HEK293, et al. (1997), mouse) cardiomyocyte Zhang et al. (1999), Ufret- Vincenty et al. (2001a, 2001b), Montpetit et al. (2009), Ednie et al. (2013)

Nav1.9 (rat) DRG Glycosidase ND Channel gating Tyrrell et al. (2001)

Voltage-gated calcium channels (Cav)

Cav3.2 HEK293 N192 D1 S3–S4 Channel surface Weiss (human) N271 linker trafﬁcking, sta- et al. (2013), N1466 D1 S5–S6 bility and gating Orestes N1710 linker et al. (2013) D3 S5–S6 linker D4 S3–S4 linker

Cav3.2 (rat) DRG Neuraminidase ND Channel gating Orestes et al. (2013) Hyperpolarisation-activated cyclic nucleotide-gated channels (HCN) HCN1 CHO N327 (N338) S5–S6 No influence on Hegle (mouse) linker channel expres- et al. (2010), sion and gating; Much important for et al. (2003) subunit heteromerisation HCN2 CHO N380 (N407) S5–S6 Channel surface Hegle (mouse) linker trafficking; no et al. (2010), influence on Much channel gating; et al. (2003) important for subunit heteromerisation Transient receptor potential channels (TRPs) TRPC3 COS-7 N418 (N404 S1–S2 No influence on Vannier (human iso- canonical linker channel expres- et al. (1998) form 3) isoform) sion and gating TRPC6 HEK293 N473 S1–S2 Channel gating Dietrich (human) N561 linker et al. (2003) S3–S4 linker (continued)

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Table 1 (continued) Glyco. site/ Channel Cell type (eq. human) Location Functional role References TRPM4b COS-7, N988 (N992) S5–S6 Channel surface Woo (mouse, HEK293 linker activity, possi- et al. (2013), human) bly by Syam stabilising the et al. (2014) protein or controlling gating TRPM5 HEK293 N932 S5–S6 Channel surface (Syam (human) linker activity, possi- et al. 2014) bly by stabilising the protein or controlling gating TRPM8 HEK293, N934 (N934) S5–S6 Channel gating Pertusa (mouse) DRG linker (temperature- et al. (2012), dependent acti- Morenilla- vation) and Palao membrane et al. (2009) localisation into raft TRPV1 (rat) HEK293, F11 N604 (N604) S5–S6 Channel gating Veldhuis linker et al. (2012) TRPV4 HEK293, N651 (N651) S5–S6 Channel surface Xu (mouse) COS-7 linker trafficking and et al. (2006), possibly gating Arniges et al. (2006) TRPV5 HEK293 N358 S5–S6 Channel surface Chang (human) linker trafficking et al. (2005) TRPP2 HeLa, N299 S1–S2 Channel surface Hofherr (human) HEK293, N305 linker trafficking and et al. (2014) LLCPK1, N328 S1–S2 stability IMCD3 N362 linker N375 S1–S2 linker S1–S2 linker S1–S2 linker TRPML1 HSF Glycosidase ND Likely channel Kiselyov (human) trafficking et al. (2005) through the Golgi TRPML3 HEK293 Glycosidase ND Likely channel Kim (human) gating et al. (2007) Acid-sensing ion channels (ASICs) ASIC1a (rat, CHO, hippo- N366 (N368) S1–S2 Channel surface Kadurin mouse) campal N393 (N395) linker trafficking and et al. (2008), dendritic (continued)

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Table 1 (continued) Glyco. site/ Channel Cell type (eq. human) Location Functional role References neurons, S1–S2 targeting; likely Jing Xenopus linker channel gating et al. (2012) oocyte ASIC1b Xenopus N192 (N193) S1–S2 Channel surface Kadurin (rat) oocyte N216 (N217) linker trafﬁcking et al. (2008) N399 (N402) S1–S2 N426 (N429) linker S1–S2 linker S1–S2 linker Two-pore domain K+ channels (TASK)

K2P3.1 (rat) COS-7, N53 (N53) S1–S2 Channel surface Mant HEK293 linker trafﬁcking and et al. (2013) gating

K2P9.1 (rat) COS-7, N53 (N53) S1–S2 Weak influence Mant HEK293 linker on channel sur- et al. (2013) face trafficking and gating Two-pore channels (TPCs) TPC1 HEK293, N599 D2 S5–S6 Channel gating Hooper (human) SKBR3 N611 linker et al. (2011) N616 D2 S5–S6 linker D2 S5–S6 linker TPC2 HEK293, N594 (N611) D2 S5–S6 ND Zong (mouse) COS-7 N601 (N618) linker et al. (2009) D2 S5–S6 linker Pannexin channels (Panx) Panx1 HEK293, N254 S3–S4 Channel surface Boassa (human) NRK linker trafficking; Panx et al. (2007), Panx2 HEK293, N86 S1–S2 heteromerisation Boassa (human) NRK linker et al. (2008), Panx3 HEK293, N71 S1–S2 Penuela (human) NRK linker et al. (2007) Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) CFTR BHK-21, N894 D2 S1–S2 ERAD degrada- Chang (human) CHO-K1, N900 linker tion (N894); et al. (2008) human epithe- D2 S1–S2 channel surface lial tissue linker trafficking (N900) CLC Channels CLC-1 Xenopus N430 L–M linker Channel surface Schmidt-Rose (human) oocyte trafficking and Jentsch (1997) (continued)

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Table 1 (continued) Glyco. site/ Channel Cell type (eq. human) Location Functional role References CLC-3 Xenopus Glycosidase ND ND Schmieder (mouse, oocyte, mouse et al. (2001) Xenopus brain, kidney laevis) and intestine CLC-5 Xenopus N169 (/) B–C linker Channel surface Schmieder (Xenopus oocyte N470 (N408) L–M linker trafﬁcking and et al. (2007) laevis) stability HEK-293 human embryonic kidney 293, CHO Chinese hamster ovary, COS ﬁbroblast-like cell from monkey kidney tissue, CAD Cath.a differentiated, Sf9 Spodoptera frugiperda, B35 neuroblastoma cell, M1 murine cortical collecting duct, SKBR3 human breast cancer cell. Amino acid residues indicated into brackets correspond to the human equivalent.

electrical activity (Norring et al. 2013; Du et al. 2014). A possible role of Kv11.1 glycosylation for the pathogenesis of long QT syndrome is discussed in Sect. 7. N-glycosylation was also shown to play a significant role in the trafficking, stability and surface expression of Kv1.4 channel (Watanabe et al. 2004) (Fig. 2, Table 1). However, while glycosylation-deficient Kv1.4 mutants were characterised by decreased stability and cell surface expression, preventing glycosylation of the closely related Kv1.1 isoform had minor effects in these respects (Watanabe et al. 2004). In contrast, N-glycosylation status, especially terminal sialylation, was shown to affect gating properties of Kv1.1, slowing activation and inactivation kinetics of the channel as well as recovery from C-type inactivation, while Kv1.4 gating was not affected (Watanabe et al. 2003; Schwetz et al. 2010). Similarly, terminal sialic acid residues have major impact on the gating of Kv1.5, and glycosylation-deficient channels showed a large depolarising shift in the voltage dependence of activation as well as a slowing of the activation kinetics (Schwetz et al. 2010). Channel gating alterations are in accordance with the surface potential theory, which assumes that external negative charges on the channel, provided by terminal sialic acid moieties, contribute to the surface electrical potential by establishing the electric field, affecting the channel voltage sensor (Schwetz et al. 2010). It is however not the case for Kv1.4 where desialylation has no impact on the channel gating, in contrast to Kv1.1 and Kv1.5 (Watanabe et al. 2003). Functional studies on Kv1.2 and Kv1.3 also revealed that preventing glycosylation drastically alters surface trafficking (increased ER retention) and gating of the channels (Watanabe et al. 2007; Zhu et al. 2012). In contrast to Kv1 channels, preventing N-glycosylation of Kv3.1b by mutating the two canonical glycosylation sites present in the protein (N220 and N229) has no effect on the trafficking and surface expression of the channel (Brooks et al. 2006; Hall et al. 2011). However, functional analysis revealed a significant alteration in the gating properties, including slower activation and deactivation kinetics, faster inactivation kinetics and a depolarising shift of the voltage dependence of activation. In addition, a profound study on Kv3.1 using various mutated CHO cell lines producing different types of oligosaccharides revealed that the nature of the

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N-glycan plays a critical role in the targeting of the channel into specific subcellular membrane loci, which becomes especially apparent during cell–cell interactions. For instance, Kv3.1 channels carrying complex N-glycans produced by CHO Pro-5 cells were found expressed mostly at the cell–cell border, while channels carrying mannose-rich glycans produced by CHO Lec1 were significantly less present. Furthermore, the molecular structure of N-glycan significantly influences cell migration assessed by wound healing assay. Indeed, cells producing glycosylated channel variants migrate faster than cells producing non-glycosylated channels. Moreover, faster migration was also observed for cells producing channels with complex N-glycan (CHO Pro-5) compared to cells expressing mannose-rich channels (CHO Lec1) or carrying bisecting-type N-glycans (CHO Lec10B) (Hall et al. 2013). This observation is of critical importance since expression of Kv3.1 with complex N-glycans has been documented during cancer and could possibly explain the increased migratory rate of cancerous cells during malignant transfor- mation. Moreover, it reveals the functional importance of N-glycosylation and highlights how N-glycan structures of glycoproteins contribute to store information in respect to the channel localisation and function. The importance of N-glycosylation in cellular distribution and clustering of Kv3.1 channels was further demon- strated in a recent study using neuroblastoma cells (B35) where monoglycosylated and non-glycosylated Kv3.1 channels accumulated predominantly in the soma as opposed to the wild-type channel, which was distributed almost evenly in the cell body and outgrowths (Hall et al. 2014). Furthermore, cells expressing non-glycosylated channels lost cell adhesion capacity between neighbouring cells. Altogether, these data document the essential role of N-glycans in carrying information about spatial cellular arrangement of the channel, which translates into specific cellular properties, function and cell–cell interaction. Finally, N-glycosylation is not the only form of post-translational glycosylation affecting voltage-gated potassium channels. For instance, Kv2.1, Kv4.2 and Kv4.3 were shown to not be N-glycosylated but rather O-glycosylated (Schwetz et al. 2011). However, similarly to what was observed with N-glycans, alteration of the terminal sialic acid moiety from O-glycans has also a significant impact on channel gating (Ufret-Vincenty et al. 2001b; Schwetz et al. 2011). This discloses that O-glycosylation can also be responsible for modulating activity of ion channels and suggests that this post-translational modification should be also taken into account as an important contributor to channel function.

4.2 Voltage-Gated Sodium Channels (Nav)

Voltage-gated Na+ channels are multimers that consist of the pore-forming Nav-subunit and auxiliary β-subunits (Catterall 2000). Early biochemical analyses have revealed that Nav channels are heavily glycosylated (Barchi et al. 1980; Miller et al. 1983; Messner and Catterall 1985; Schmidt and Catterall 1986, 1987), which is critical for proper folding and assembly of the channel (Schmidt and Catterall

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1987). In addition, glycosylation (sialylation) has been shown to modulate gating properties of purified eel Na+ channels reconstituted in artificial lipid bilayer (Recio-Pinto et al. 1990; Cronin et al. 2005). Similarly, desialylation of skeletal Nav1.4 and cardiac Nav1.5 channels expressed in mammalian cell lines induces a depolarised shift of the voltage dependence of activation of both channel types, while a depolarising shift of the voltage dependence of inactivation was only observed for Nav1.4, suggesting that glycosylation differentially regulates skeletal and cardiac Nav channels (Bennett et al. 1997; Zhang et al. 1999). However, channel desialylation had no effect on single-channel properties (single conductance, mean open time and open probability). In addition, Nav1.4 was expressed in CHO cells impaired in their ability to produce either sialylated or polysialylated N-glycans. While the loss of sialylation produced similar channel gating alteration as pharmacological desialylation (i.e. depolarising shift of activation and inactivation), the loss of polysialylated N-glycans produced opposite gating effect (i.e. hyperpolarising shift of activation and inactivation) (Ahrens et al. 2011), indicating that sialylation and polysialylation differentially modulate skeletal Nav1.4 channel. The functional importance of Nav1.5 glycosylation on cardiac electrical activity was further investigated in primary cardiomyocytes. Interest- ingly, the analysis of glycosylation-associated genes (glycogenes) revealed that the polysialyltransferase ST8sia2 is specifically expressed in neonatal atrium, and / electrophysiological recordings on ST8sia2À À atrial myocytes showed significant alterations of Nav1.5 gating (consistent with what was observed in heterologous expression systems) and action potential waveforms (Montpetit et al. 2009). In contrast, the absence of ST8sia2 had no influence on ventricular electrical activity, / which is however potently altered in ST3Gal4-deficient myocytes (ST3Gal4À À) (Ednie et al. 2013). These results offer insights into the importance of channel glycosylation in cardiac physiology and reveal a fundamental mechanism by which cell-type-specific expression profile of glycogenes contributes to specialised cellular functions. The potential role of Nav1.5 glycosylation in the genesis of cardiac arrhythmias is further discussed in Sect. 7 of this review. Finally, an interesting report documented a role for glycosylation in the control of Nav1.9/NaN channels. Nav1.9 is preferentially expressed in small dorsal root ganglion and trigeminal ganglion neurons (Dib-Hajj et al. 1998; Dib-Hajj et al. 2002; Fjell et al. 2000; Coste et al. 2007) where it contributes to pain signalling pathway (Bird et al. 2013; Leipold et al. 2013; Huang et al. 2014). Biochemical analysis indicated that endogenous Nav1.9 expressed in dorsal root ganglion presents two glycosylated states at early neonatal ages. In contrast, only the lightly glycosylated form was detected in adults (Tyrrell et al. 2001). Interest- ingly, patch-clamp recordings on dissociated DRG neurons revealed that the steady-state inactivation of the channel in neonatal neurons is shifted by 7 mV toward hyperpolarised potentials compared with adult neurons. Consistent with a role of glycosylation in differential channel gating, the application of neuraminidase on neonatal neurons produced a depolarised shift of the steady-state inactivation, making the channel indistinguishable from that of adult neurons (Tyrrell et al. 2001). It is likely that differences in glycosylation pattern of Nav1.9 arise

[email protected] 82 J. Lazniewska and N. Weiss from developmentally regulated glycogene expression that in turn may cause neonatal and adult sensory neurons to respond differently to similar stimuli.

4.3 Voltage-Gated Calcium Channels (Cav)

Voltage-gated Ca2+ channels (VGCCs) have evolved as one of the most important source of Ca2+ entry into the cells, orchestrating a plethora of cellular functions (Catterall 2011). Recently, glycosylation of low-voltage-activated Ca2+ channels, the so-called T-type channels, was described. Perhaps one of the most remarkable features of T-type channels is their low threshold of activation that makes these channels important candidates for Ca2+ entry near the resting membrane potential of the cells (Perez-Reyes 2003; Nilius and Carbone 2014). Hence, they mediate low-threshold burst discharges (Huguenard and Prince 1992; Huguenard 1996) that occur during different forms of neuronal rhythmogenesis (Crunelli et al. 2006; Bal and McCormick 1997; Beurrier et al. 1999; Sotty et al. 2003) but also play important roles in sensory transmission (Francois et al. 2014; Krahe and Gabbiani 2004; Jacus et al. 2012; Todorovic and Jevtovic-Todorovic 2013; Kim et al. 2003; Bourinet et al. 2005), as well as in hormone and neurotransmitter release (Weiss and Zamponi 2013; Weiss et al. 2012b; Weiss et al. 2012a; Carbone et al. 2014). Additionally, they have been associated in an increasing number of pathological situations (Nilius and Carbone 2014; Dziegielewska et al. 2014; Yang et al. 2014; Mesirca et al. 2014; Proft and Weiss 2014) including painful diabetic neuropathy (Todorovic and Jevtovic-Todorovic 2014) and some forms of epilepsy (Cain et al. 2014; Powell et al. 2014; Chen et al. 2014; Cheong and Shin 2014; Zamponi et al. 2010; Khosravani and Zamponi 2006). Using site-directed mutagenesis of canonical N-glycosylation sites present in human Cav3.2 T-type channels, Weiss et al. (2013) identiﬁed two N-glycosylation sites, N192 and N1466, responsible for surface expression and activity of the channel, respectively (Fig. 3, Table 1). In contrast, preventing glycosylation at asparagines N271 and N1710 resulted in low expression level and nonfunctional channel, respectively. At the same time, Orestes

Fig. 3 Schematic representation of the location of validated N-glycosylation sites in the scheme of voltage-gated calcium channel (Cav) structure

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at al. (2013) also showed that Cav3.2 channel mutated at asparagines N192Q and N1466Q displayed impaired surface expression and altered current kinetics. In this study, however, N192 was identified as a regulator of channel activity, while N1466 was identified as a determinant of channel surface expression. It is possible that different experimental conditions, such as glucose level or specific enzymatic deglycosylation, may have produced specific N-glycans contributing to the different observations. However, regardless of observed differences, it is unambiguous that Cav3.2 channels undergo N-glycosylation, which has profound impact on channel expression and activity and possibly in pathophysiology (this aspect is further detailed in Sect. 7).

4.4 Hyperpolarisation-Activated Cyclic Nucleotide-Gated Channels (HCN)

HCN channels are considered as a subfamily of voltage-gated potassium channels. They are characterised by dual-gating mode (i.e. are activated by both hyperpolarisation and cyclic nucleotides). They conduct mix currents and are permeable for both Na+ and K+. HCN channels are expressed mainly in the nervous system and the heart (and in a much lesser extent in other tissues) where they contribute to rhythmic activities. They present the same basic structure as voltage- gated cation channels (Biel et al. 2009). Two isoforms, HCN1 and HCN2, were shown to be N-glycosylated (Fig. 4, Table 1). Surprisingly, preventing N-glycosylation of HCN1 and HCN2 does not signiﬁcantly impact the functioning of the channels (Hegle et al. 2010). Interestingly, although HCN1 and HCN2 channel isoforms are closely related, proper N-glycosylation of HCN2 is critical for efﬁcient

Fig. 4 Schematic representation of the location of validated N-glycosylation sites in the scheme of hyperpolarisation-activated cyclic nucleotide-gated channel (HCN) structure. The corresponding residues in the human channels are indicated in brackets

[email protected] 84 J. Lazniewska and N. Weiss expression of the channel at the plasma membrane, while HCN1 surface expression is independent of N-glycan attachment. From an evolutionary point of view, it is possible that glycosylation was not a prerequisite for proper expression of ancestor channels, and as a consequence of gene duplication and evolution of complex organisms, glycosylation appeared as a means to ﬁne-tune channel activity and fulﬁl highly specialised cellular functions. However, a recent study documented an increased surface expression of HCN1 by the auxiliary subunit TRIP8b (tetratricopeptide repeat-containing Rab8b-interacting protein), possibly by enhancing glycosylation of the channel (Wilkars et al. 2014). In addition, proper glycosylation of HCN channels is critical for subunit heteromerisation (Much et al. 2003) and possibly regulated by neuronal electrical activity, which may have important consequences during epileptic seizures (Zha et al. 2008). This aspect is further discussed in Sect. 7.

4.5 Transient Receptor Potential Channels (TRPs)

The transient receptor potential (TRP) channels encompass several subfamilies including TRPC (the “C” stands for canonical), TRPV (“V” for vanilloid), TRPM (“M” for melastin), TRPP (“P” for polycystic), TRPML (“ML” for mucolipin), TRPA (“A” for ankyrin) and TRPN (“N” for no mechanoreceptor potential) (Nilius and Owsianik 2011; Nilius and Owsianik 2011). TRP channels differ from other channel families by their lack of common gating mode and ion selectivity. They are broadly expressed in many tissues and share the same general structure with voltage-gated cation channels and HCN channels, although the 4th transmembrane segment (S4) is usually devoid of a complete set of positively charged residues responsible for efficient voltage sensing (Huang 2004), even though putative voltage sensors have been documented for some of the TRP channels which appeared to be weakly voltage gated (Voets et al. 2007; Nilius et al. 2005; Nilius et al. 2007). TRP channels undergo numerous post-translational modifications (Voolstra and Huber 2014), and a subset of channels undergoes N-glycosylation (Cohen 2006). TRPC3 and TRPC6 pose a group of channels, activated by diacylglycerols (DAGs) and permeable for Ca2+. It was shown that the difference in their getting is governed by specific N-glycosylation patterns (Dietrich et al. 2003). For instance, TRPC3 is monoglycosylated (Vannier et al. 1998), while TRPC6 is dually glycosylated (N473, N561) (Fig. 5, Table 1). Monoglycosylated TRPC3 channel exhibits constitutive activity, while the activity of dually glycosylated TRPC6 is tightly receptor and phospholipase C dependent. Interestingly, the removal of one N-glycosylation site (N561) in TRPC6 is sufficient to convert the channel to a constitutive active mode. On the other hand, the introduction of a second N- glycosylation site into TRPC3 gives rise to a substantial loss of constitutive activity of the channel. These results indicate that N-glycans, to a large degree, determine the mode of gating of TRPC channels. However, although the glycosylation status

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Fig. 5 Schematic representation of the location of validated N-glycosylation sites in the scheme of transient receptor potential canonical channel (TRPC) structure

Fig. 6 Schematic representation of the location of validated N-glycosylation sites in the scheme of transient receptor potential melastatin-related channel (TRPM) structure. The corresponding residues in the human channels are indicated in brackets influences channel activity, trafficking and surface expression were undisturbed in glycosylation-deficient mutants. TRPM4 and TRPM5 were shown to be N-glycosylated at asparagines N992 and N932, respectively, in the pore-forming region between the 5th and 6th transmembrane segments (Fig. 6, Table 1). TRPM4a and TRPM4b are products of the alternative splicing of the same transcript. In contrast to the short variant TRPM4a, TRPM4b is activated by Ca2+ but is impermeable to Ca2+ (Petersen 2002; Nilius and Vennekens 2006; Launay et al. 2002). Interestingly, while surface expression of TRPM4b and TRPM5 does not rely on proper glycosylation of the channel, the functional expression, in contrast, depends on the attachment of complex N-glycans (Woo et al. 2013; Syam et al. 2014). However, the exact role of N-glycan in the control of TRPM4b and TRPM5 activity is not fully understood at this stage since opposite results were observed depending if the glycosylation status of the channel was modified by genetic manipulation or pharmacological approach (Syam et al. 2014).

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An interesting representative of TRPM subfamily is TRPM8. Expressed primar- ily in nociceptive neurons, it is involved in the sensation of cold and is activated by both physical (temperature) and chemical (e.g. methanol) stimuli (McKemy et al. 2002). N-glycosylation at asparagine N934 located in the proximity of the extracellular end of the transmembrane domain S6 was shown to target TRPM8 into lipid raft (cholesterol-rich) membrane domains, critical for proper channel activity (Morenilla-Palao et al. 2009). Indeed, disruption of TRPM8 glycosylation drastically decreases channel sensitivity to cold and methanol by shifting the voltage dependence of channel activation toward more positive potentials. Consistent with this observation, enzymatic deglycosylation of native TRPM8 channels expressed in trigeminal sensory neurons markedly altered thermal sensitivity, causing an important shift of the temperature threshold of cold-sensitive thermoreceptors, indicating the fundamental role of N-glycan in TRPM8-dependent cold perception (Pertusa et al. 2012). Considering that the close homologues TRPM4 and TRPM5 are, in contrast to TRPM8, activated by heat (Talavera et al. 2005) and potentially expressed in primary nociceptive neurons (Vandewauw et al. 2013), it is possible that glycosylation of these channels may also contribute to proper thermal perception. Several TRPV channels are also regulated by N-glycosylation (Fig. 7, Table 1). For instance, glycosylation of TRPV1, a key sensor in pain-sensing nociceptors, has been reported when expressed in HEK293 cells and in the more native context of the dorsal ganglion-derived F-11 cell line (Jahnel et al. 2001). Further functional analysis revealed that glycosylation at asparagine N604 is critical for channel Ca2+ permeation and desensitisation triggered by capsaicin (Wirkner et al. 2005; Veldhuis et al. 2012). Indeed, while capsaicin induces sustained increase in intracellular Ca2+ level in cells expressing wild-type TRPV1, a rapid desensitisation was observed in cells expressing TRPV1N604T mutant channels. Although it is unambiguous that TRPV1 undergoes functional glycosylation when transiently expressed in mammalian cell lines, an earlier report on endogenous TRPV1 natively expressed in dorsal root ganglion did not detect glycosylation of the channel (Kedei et al. 2001). As for TRPV1, glycosylation of the osmoresponsive

Fig. 7 Schematic representation of the location of validated N-glycosylation sites in the scheme of transient receptor potential vanilloid channel (TRPV) structure. The corresponding residues in the human channels are indicated in brackets

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Fig. 8 Schematic representation of the location of validated N- glycosylation sites in the scheme of transient receptor potential polycystin channel (TRPP) structure

TRPV4 channel was initially suspected based on gel migration of TRPV4 transiently transfected in mammalian cells or natively expressed in kidney tissue (Xu et al. 2003). In a subsequent study, it was shown that preventing N-glycosylation at asparagine N651 produces an unexpected increase in channel activity and Ca2+ influx upon hypotonic stimulation, consistent with an increased trafficking and surface expression of the channel (Xu et al. 2006; Arniges et al. 2006). In contrast, while glycosylation of TRPV5 at asparagine N358 was also shown to favour channel trafficking and expression at the plasma membrane, it did not substantially influence Ca2+ currents (Chang et al. 2005). An interesting mode of regulation of TRPV5 by klotho via glycosylation of the channel is discussed in Sect. 5.2. TRPP channels pose the next subfamily of TRP channels that undergoes glycosylation. One of the distinctive features of TRPP channels is the presence of a large extracellular loop between the transmembrane segments S1 and S2. Earlier biochemical studies revealed that TRPP2 is particularly highly glycosylated (Cai et al. 1999; Newby et al. 2002). Indeed, sequence analysis revealed not less than five potential glycosylation sites (N299, N305, N328, N362 and N375), all of them located in this first extracellular loop (Fig. 8, Table 1). Recently, sequential disruption of those canonical sites by site-directed mutagenesis revealed that glycosylation of TRPP2 is critical for proper trafficking and stability of the channel at the membrane (Hofherr et al. 2014). In addition, glucosidase II (GII) was shown to mediate glycan trimming of TRPP2. Interestingly, genetic knockout in mouse of the non-catalytic β-subunit of GII (GIIβ) encoded by the gene Prkcsh, which has been associated with autosomal dominant polycystic liver disease (ADPLD), impaired TRPP2 glycosylation and expression (Hofherr et al. 2014). The impaired GIIβ-dependent glucose trimming of TRPP2 glycosylation in ADPLD may explain

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/ the decreased TRPP2 protein expression in PrkcshÀ À mice and the genetic interaction observed between TRPP2 and PRKCSH in ADPLD. Finally, TRPML channels are essentially expressed in the endosomal pathway where they may function as Ca2+/cation release channels in endosomes, lysosomes and lysosome-related organelles. They may also contribute to the endolysosomal transport and fusion processes. However, the exact physiological roles of TRPML channels remain quite elusive, and whether they are strict endolysosomal channels or whether they may also be functionally expressed at the plasma membrane has to be determined (Zeevi et al. 2007; Cheng et al. 2010; Grimm et al. 2012; Weiss 2012). Although glycosylation of TRPML channels per se has not been really investigated, it is likely that TRPML1 undergoes sialylation in the Golgi prior to the transport of the channel to lysosomes (Kiselyov et al. 2005). Glycosylation of TRPML3 has also been documented (Kim et al. 2007) and is detailed in section 7 in the context of the varitint–waddler phenotype.

4.6 Acid-Sensing Ion Channels (ASICs)

ASIC channels belong to the amiloride-sensitive cationic channel superfamily. Widely expressed throughout the central and the peripheral nervous system, they have been implicated in various diseased states including pain and neurological and psychiatric diseases (Wemmie et al. 2013; Chu et al. 2014). They are sensitive to pH (activated by H+) and selective for Na+, except for ASIC1, which is also permeable for Ca2+. The characteristic feature of these channels arises from their sensitivity to amiloride, a nonspecific inhibitor that also inhibits other Na+ channels and exchangers. ASIC channels are trimeric protein complexes made up of different combinations of subunits consisting of two transmembrane segments and a large extracellular cysteine-rich domain (Saugstad et al. 2004; Wemmie et al. 2006). The importance of glycosylation in the control of ASIC channels has been documented for ASIC1 (Jing et al. 2011). While ASIC1a and ASIC1b variants (both encoded by a single gene ASIC1) present many structural similarities, they differ in their functionality such as affinity for H+ (higher for ASIC1a) or permeability for Ca2+ (supported only by ASIC1a), and recent studies indicate that differences between these closely related channels arise, at least partly, from different glycosylation status. Both ASIC1a and ASIC1b present two canonical N-glycosylation sites located in the distal part of their ectodomain (N366 and N393 for ASIC1a; N399 and N426 for ASIC1b) (Fig. 9, Table 1). However, ASIC1b also contains two additional N-glycosylation sites (N192 and N216) in the proximal part of its ectodomain. Functional analysis of glycosylation-deficient channels revealed that glycosylation at distal sites significantly increases surface expression of both ASIC1a and ASIC1b but is not an absolute prerequisite for channel activity (Kadurin et al. 2008). In contrast, glycosylation of at least one of the two proximal sites in ASIC1b is an absolute requirement for proper surface expression of the channel. In addition, functional analysis of native ASIC1a indicates that N-

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Fig. 9 Schematic representation of the location of validated N-glycosylation sites in the scheme of acid-sensing ion channel (ASIC) structure. The corresponding residues in the human channels are indicated in brackets glycosylation is critical for dendritic targeting of the channel and acidosis-induced spine remodelling (Jing et al. 2012). This observation nicely illustrates how glycosylation of a given channel, by controlling not only its surface expression and activity but also its appropriate targeting into speciﬁc subcellular loci, contributes to the proper physiology of the neuron.

4.7 Two-Pore Domain K+ Channels (TASK)

+ TASK channels pose a subfamily of two-pore domain (K2P)K channels, which are characterised by the existence of two-pore-forming loops between the transmembrane segments S1–S2 and S3–S4. Two repeats of four segments are assembled as dimers to form a functional channel (Lotshaw 2007). These channels are expressed in many different tissues throughout the body where they contribute to a plethora of physiological functions and pathological conditions (Bittner et al. 2010). K2P channels are not voltage dependent and thus are active at resting electrical membrane potentials where they support background K+ currents (Enyedi and Czirjak 2010). K2P3.1 and K2P9.1 belong to a subgroup of acid-sensitive K2P channels and are both N-glycosylated at asparagine N53 (Fig. 10, Table 1). While glycosylation of K2P3.1 significantly affects channel activity, it influences K2P9.1 to a much lesser extent (Mant et al. 2013). Functional analysis of glycosylation-deficient K2P3.1 showed reduced membrane K+ currents and channel surface expression. Co-localisation studies with ER–Golgi intermediate compartment (ERGIC) revealed that non-glycosylated channels efficiently traffic from the ER to the

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Fig. 10 Schematic representation of the location of validated N- glycosylation sites in the scheme of two-pore domain potassium channel (K2P) structure. The corresponding residues in the human channels are indicated in brackets

ERGIC and Golgi apparatus, although they were barely detected at the cell surface. In addition, turnover assay revealed that a fraction of glycosylation-deﬁcient channels eventually reaches the cell surface but was immediately directed to endosomal degradation pathway. Hence, N-glycosylation is responsible for proper surface expression, stability and activity of K2P3.1, while activity of K2P9.1 is much less dependent on the glycosylation state of the channel, highlighting once again important differences in the functional impact of glycosylation among different channels even closely related.

4.8 Two-Pore Channels (TPCs)

TPCs (two-pore channels) are composed of two repeats of six transmembrane segments. Expressed exclusively in acidic organelles (endolysosomes), TPC channels are considered as non-selective channels, likely permeable to Ca2+, Na+ and K+ (Galione et al. 2009; Patel and Brailoiu 2012; Morgan and Galione 2014). They were found to be sensitive to nicotinic acid adenine dinucleotide phosphate (NAADP), which can induce Ca2+ release from lysosome-related stores (Calcraft et al. 2009). Glycosylation of human TPC1 was shown at asparagines N599, N611 and N616 (Hooper et al. 2011), whereas mouse TPC2 is glycosylated at asparagines N594 and N601 (Zong et al. 2009) (Fig. 11, Table 1). While preventing glycosylation of TPC1 does not alter intracellular distribution of the channel, it produces increased

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Fig. 11 Schematic representation of the location of validated N-glycosylation sites in the scheme of two-pore channel (TPC) structure. The corresponding residues in the human channels are indicated in brackets

NAADP-induced Ca2+ release, indicating that proper glycosylation of the channel is critical to prevent aberrant intracellular Ca2+ elevations under NAADP stimulation (Hooper et al. 2011).

4.9 Pannexin Channels (Panx)

Pannexin channels comprise three isoforms, Panx1, Panx2 and Panx3. These proteins are composed of four transmembrane segments, connected by two extracellular and intracellular loops (Baranova et al. 2004). Pannexins are predicted to form hexamers (Panx1 and Panx3) or heptamers (Panx2) in the plasma membrane (Boassa et al. 2007; Ambrosi et al. 2010), and although they bear signiﬁcant sequence homology with the invertebrate gap junction proteins innexins, there is

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Fig. 12 Schematic representation of the location of validated N-glycosylation sites in the scheme of pannexin channel (Panx) structure currently no evidence in the literature that pannexins form gap junctions (Sosinsky et al. 2011). These non-selective channels are expressed in virtually every mammalian organ, and altered expression of pannexin channels has been associated with numerous human diseases (Penuela et al. 2014a). N-glycosylation of Panx1, Panx2 and Panx3 was documented at asparagines N254, N86 and N71, respectively, and was shown to be important for surface targeting of the channels (Fig. 12, Table 1) (Boassa et al. 2008; Boassa et al. 2007; Penuela et al. 2009). However, although pannexin channels are functionally N-glycosylated, they do not undergo sialylation (Penuela et al. 2014b). Moreover, a tissue-dependent glycosylation pattern of Panx1 was documented, suggesting that Panx1 activity may be dependent of the cellular environment (Penuela et al. 2007). In addition, proper glycosylation of pannexins was shown to be necessary for protein– protein interaction among pannexin family members (Penuela et al. 2009),

4.10 CFTR Channel

The cystic fibrosis transmembrane conductance regulator (CFTR) is a member of the ABC transporter family that forms a chloride channel in the apical membrane of epithelia where it mediates transepithelial salt and liquid movement, and mutations in the gene encoding the channel cause congenital cystic fibrosis (Sheppard and Welsh 1999). Numerous earlier works have documented the biochemical glycosylation of the CFTR (Scanlin and Glick 1999;O’Riordan et al. 2000; Scanlin and Glick 2001), and defective N-glycosylation was shown to reduce cell surface expression of the CFTR by impairing both early and endocytic trafficking of the channel (Glozman et al. 2009). Sequence analysis of CFTR channel indicates two canonical N-glycosylation sites, both located in the extracellular S1–S2 loop of the domain two (Fig. 13, Table 1). Using glycosylation-deficient CFTR mutants, it was proposed that glycosylation at asparagine N894 accelerates ERAD degradation of the channel, whereas the glycan at asparagine N900 promotes maturation and progression of the protein through the Golgi (Chang et al. 2008). Hence, individual glycan loci direct the ER processing of CFTR in different ways, suggesting that

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Fig. 13 Schematic representation of the location of validated N-glycosylation sites in the scheme of cystic ﬁbrosis transmembrane conductance regulator channel (CFTR) structure balanced N-glycosylation between those two loci represents a powerful mechanism to control channel fate and surface expression level. However, the cellular signals controlling differential glycosylation pattern remain to be explored.

4.11 CLC Channels

CLC channels are integral membrane proteins whose main function is to translocate chloride ion (ClÀ) across the cell membranes (Jentsch 2008; Accardi and Picollo 2010; Vandewalle 2002). They have been implicated in many human diseased states including renal hypercalciuria (Scheinman 1998; Lippiat and Smith 2012), stroke (Zhang et al. 2013) and various cardiovascular diseases (Duan 2011). Glycosylation of Xenopus laevis CLC-5 has been reported at asparagines N169 and N470 as critical for proper surface expression and stability of the channel (Figure 14, Table 1) (Schmieder et al. 2007). In addition, biochemical glycosylation of CLC-1 and CLC-3 has been reported but was not investigated at the functional level (Schmidt-Rose and Jentsch 1997) (Schmieder et al. 2001).

5 Crosstalk Between Glycosylation and Other Signalling Pathways

5.1 Regulation by Auxiliary Subunits

Glycosylation-mediated modulation of ion channels not only relies on the glycosylation status of the pore-forming subunit but also on the proper glycosylation of the channel auxiliary subunits (Table 2). For instance, DPP10, a Kv4 channel

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Fig. 14 Schematic representation of the location of validated N-glycosylation sites in the scheme of chloride CLC channel structure. The corresponding residues in the human channels are indicated in brackets. The schematic topology of the mammalian CLC channels is based on the crystallographic structure of the prokaryotic channel (Dutzler et al. 2002) auxiliary subunit, which belongs to the dipeptidyl aminopeptidase-like (DPPL) protein family, was shown to modulate Kv4.3 in a glycosylation-dependent manner. Indeed, preventing N-glycosylation of DPP10 significantly alters Kv4.3 channel gating (Cotella et al. 2010), indicating that glycosylation of proteins closely associated with ion channels also plays a fundamental role in the proper activity of the channel. Along this line, N-glycosylation of the Cavα2δ1-subunit of high-voltage- activated Ca2+ channels (Davies et al. 2007) at asparagines N136 and N184 located in the α2 domain of the protein is critical for Cav1.3 channel potentiation induced by Cavα2δ1 (Andrade et al. 2009). Similarly, proper O-glycosylation of the KCNE1 auxiliary subunit of Kv7.1 channel is necessary for efficient surface trafficking and expression of the channel (Chandrasekhar et al. 2011). Also, proper glycosylation of SUR1, a member of the ATP-binding cassette family of proteins, which associates with the inward rectifier Kir6.x to form ATP-sensitive potassium channels

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Table 2 Glycosylation-dependent factors modulating ion channel function Modulation Channel Cell type factor Functional role References Modulation by channel auxiliary subunits

Kv4.3 CHO DPP10 DPP10 N-glycosylation required Cotella for proper channel gating et al. (2010)

Kv7.1 CHO KCNE1 KCNE1 O-glycosylation required Chandrasekhar for surface trafﬁcking and expres- et al. (2011) sion of the channel Kir6.2 COS-1 SUR1 SUR1 N-glycosylation required for Conti (KATP) proper trafﬁcking and surface et al. (2002) expression of KATP channel

Cav1.3 HEK293 Cavα2δ1 Cavα2δ1 N-glycosylation required Andrade for functional surface expression of et al. (2009) the channel

Nav1.7 HEK293 Navβ1/Navβ3 Navβ1 and Navβ3 inﬂuence N-gly- Laedermann cosylation of Nav1.7 that in turn et al. (2013) modulates channel gating Klotho-dependent regulation TRPV5 HEK293 Klotho Klotho-dependent hydrolysis of Cha TRPV5 sialic acids allows binding et al. (2008), of gelactin-1/3 that in turn Leunissen stabilises channel membrane et al. (2013) expression and activity Glycosylation-dependent channel gene expression CLC-2 L2 SP1 Glycosylation of the transcription Vij and Zeitlin factor SP1 enhances transcriptional (2006) activity and CLC-2 gene expression Modulation by external sugar

Kv1.3 CHO L-fucose/ Increased channel expression, pos- Zhu Neu5Ac sibly caused by increased et al. (2012) GlcNAc fucosylation and sialylation of N- glycans Increased channel expression likely caused by decreased size of N-glycan branches but increased level of branching

K2P3.1 COS-7, Glucose Increased external glucose concen- Mant HEK293 tration induces glycosylation- et al. (2013) dependent channel potentiation

Cav3.2 HEK293 Glucose Increased external glucose concen- Weiss tration induces glycosylation- et al. (2013), dependent channel potentiation Orestes et al. (2013) Modulation by temperature

Kv1.5 CHO Temperature Lowering temperature from 37C Ding to 28C favours glycosylation and et al. (2014) functional expression of the channel (continued)

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Table 2 (continued) Modulation Channel Cell type factor Functional role References ASIC1a CHO Temperature Lowering temperature favours gly- Jing cosylation and functional expres- et al. (2011) sion of the channel CFTR Mammalian Temperature Lowering temperature rescues Denning cell lines expression of the cystic ﬁbrosis et al. (1992) CFTR mutant ΔF508 by enhancing glycosylation and processing of the channel HEK-293 human embryonic kidney 293, CHO Chinese hamster ovary, COS ﬁbroblast-like cell from monkey kidney tissue, L2 rat lung cell

(KATP), is essential for the proper trafficking and surface expression of KATP channels (Conti et al. 2002). In contrast to Kv4.3, Kv7.1, Cav1.3 and KATP channels, where glycosylation of the auxiliary subunit itself modulates channel activity, the β-subunit of voltage- gated Na+ channels directly influences glycosylation status of the pore-forming subunit of the channel complex. Indeed, while Nav1.7 expressed alone is present in two glycosylated forms, a core-glycosylated (<250 kDa) and a fully glycosylated form (<280 kDa), co-expression of the β1-subunit mediates a third and intermediate form of the channel (<260 kDa) with specific gating properties. In contrast, co-expression of the β3-subunit increases expression of the core-glycosylated channel (Laedermann et al. 2013). One possible explanation is that in the presence of a β-subunit, the Nav-pore-forming subunit adopts different conformations possibly affecting the availability of the glycosylation sites. However, it is also conceivable that besides regulating channel activity, β-subunits affect cellular glycosylation pathways resulting in altered glycosylation patterns of the proteins. Further investigations will certainly reveal the exact role of Navβ-subunit in the control of channel glycosylation. Noteworthy, hyperglycosylation of Navβ4-subunit has been documented in aging human brains, as well as in mouse neonatal Parkinson’s disease (PD) transgenic brains. Interestingly, preventing glycosylation of Navβ4 increases neurite outgrowth and number of filopodia-like protrusions (Zhou et al. 2012). Considering that aberrant protein glycosylation has been documented in neurodegenerative diseases including PD, the possibility that altered ion channel function by defect in glycosylation pathways may cause neuronal damage and contribute to the pathophysiology of the disease seems justified. Altogether, these data reveal that modulation of ion channels by glycosylation occurs at different levels and displays a significance of the interaction between the pore-forming and auxiliary subunits.

[email protected] The “Sweet” Side of Ion Channels 97

5.2 Klotho-Dependent Regulation of TRPV5

The mammalian hormone klotho (Kuro-o 2012), abundantly expressed in distal renal tubules, presents glycosidase activity that can stimulate TRPV5-dependent Ca2+ uptake (Table 2) (Chang et al. 2005). Biochemical and functional studies revealed that klotho removes sialic acid moieties from carbohydrate chains, thus exposing N-acetyllactosamine (LacNAc), allowing the binding of galectin-1 with the channel (galectins are carbohydrate-binding proteins with high afﬁnity to LacNAc disaccharides) – formation of TRPV5/galectin-1 complexes stabilising expression of the channel in the plasma membrane (Cha et al. 2008). Interestingly, klotho-dependent stimulation of TRPV5 can be mimicked by sialidase and is dependent on proper N-glycosylation of the channel (Leunissen et al. 2013). How- ever, increased TRPV5 activity after sialidase treatment is independent of galectin and is rather caused by inhibition of lipid raft-mediated internalisation of the channel (Leunissen et al. 2013).

5.3 SP1 Induces CLC-2 Gene Expression

Enhanced transcriptional activity of the RNA polymerase II transcription factor SP1 has been shown upon glycosylation (O-GlcNAc) of the protein (Du et al. 2000). The chloride channel-2 (CLC-2) promoter has SP1 (and SP3) domains that are important for gene regulation and channel expression in foetal airway epithelia. Interestingly, preventing glycosylation of SP1 in lung cell lines induces decreased CLC-2 expression, whereas hyperglycosylation induced by high dose of glutamine, in contrast, enhanced channel expression (Table 2) (Vij and Zeitlin 2006). In addition, consistent with a high expression level of CLC-2 at early developmental stages, SP1 was found hyperglycosylated in 6-week-old lung compared to 16-week-aged tissue (Vij and Zeitlin 2006). Hence, glycosylation- dependent gene expression represents another level of regulation of ion channels, which may be particularly important for organogenesis and tissue maturation during development.

6 Modulation of Ion Channel Glycosylation by External Factors

6.1 Modulation by External Sugar

Zhu et al. (Zhu et al. 2012) investigated the inﬂuence of external monosaccharides on the expression of Kv channels (Table 2). While supplementation of CHO cells expressing Kv1.3 channel with D-mannose, D-glucose and D-galactose had minor

[email protected] 98 J. Lazniewska and N. Weiss

Fig. 15 Glycosylation-dependent glucose concentration mediates upregulation of Cav3.2 T-type Ca2+ channels and contributes to painful diabetic neuropathy. (a) While external glucose elevation has a significant influence on wild-type Cav3.2 channel activity, preventing N-glycosylation of the channel (Cav3.2 N192A/N1466Q) is sufficient to prevent glucose-induced potentiation of channel activity. (b) In vivo desialylation of N-glycan by topical injection of neuraminidase (Neu) in ob/ob mice prevents Cav3.2-dependent painful diabetic neuropathy influence, addition of N-acetylglucosamine (GlcNAc) resulted in a substantial increase in channel surface expression due to decreased internalisation. Interest- ingly, glycan analysis revealed a reduced sialic acid content and increased branching degree (but decreased branch size) of Kv1.3 glycans produced under GlcNAc supplementation. This result highlights the functional importance of external monosaccharides on ion channel function and may have important implication in (patho)physiology, for example, during restricted or supplemented diet. The influence of external glucose concentration on ion channel expression and activity was also investigated. For instance, glucose-dependent potentiation of K2P3.1 channel was documented and relies on the glycosylation status of the channel (Mant et al. 2013). Similarly, a glycosylation-dependent glucose stimula- 2+ tion of Cav3.2 Ca channels was reported (Fig. 15a) (Weiss et al. 2013), with potential implication in peripheral painful diabetic neuropathy (PDN) (Fig. 15, and see Sect. 7).

6.2 Modulation by Temperature

Protein glycosylation has been shown to be markedly sensitive to temperature (Hossler et al. 2009) (Table 2). For instance, Kv1.5 channel expressed in cells

[email protected] The “Sweet” Side of Ion Channels 99 grown under relatively low-temperature condition (28C) is glycosylated more efﬁciently than those expressed in cells maintained under standard environment (i.e. 37C) (Ding et al. 2014). This has a profound impact on channel activity since surface expression, stability and activity were shown to increase, likely due to increased glycosylation state of the channel. Similar results were also obtained with ASIC1a (Jing et al. 2011), suggesting that glycosylation-related temperature may affect many other ion channel subtypes. However, since limited information is available on stress-responsive or condition-induced glycosylation, the relevance of temperature on mammalian physiology of ion channels remains to be explored.

7 Glycosylation-Mediated Channelopathy

Considering the essential implication of ion channels in health and disease (Kim 2014) and that most of ion channel subtypes are virtually glycosylated, alteration of ion channel glycosylation could conceivably cause numerous diseased states, and in some cases a direct causal connection exists.

7.1 Long QT Syndrome

Long QT syndrome (LQT) is a rare congenital cardiac disorder characterised by a prolonged QT interval and a propensity to ventricular tachyarrhythmias, which may lead in some circumstances to syncope, to cardiac arrest or more dramatically to sudden death (Schwartz and Ackerman 2013). Mutations in the gene HERG that + encodes the cardiac voltage-gated K channel Kv11.1 have been linked to LQT syndrome (Satler et al. 1996; Benson et al. 1996), likely caused by defective channel trafﬁcking to the plasma membrane (Thomas et al. 2003). Interestingly, while glycosylation of Kv11.1 is essential for proper surface expression of the channel (Petrecca et al. 1999) and cardiac electrical activity (Du et al. 2014; Norring et al. 2013), two missense mutations (N629S and N629D) found in patients with LQT syndrome disrupt a glycosylation locus (Satler et al. 1998). In addition, LQT-associated Kv11.1 mutations Y611H and V822M, although not affecting glycosylation sites per se, result in an incompletely glycosylated form of the channel and altered surface expression (Zhou et al. 1998; Thomas et al. 2003). Altogether, these results strongly suggest that abnormal glycosylation of Kv11.1 caused by LQT-associated mutations alters proper channel expression that in turn contributes to LQT syndrome. Alteration of ion channel glycosylation in cardiac arrhythmogenesis was also + reported for the voltage-gated Na channel Nav1.5 (Ufret-Vincenty et al. 2001a). / Indeed, biochemical and functional analysis of Nav1.5 in MLPÀ À cardiomyocytes (a genetic model of heart failure) revealed altered channel gating caused by aberrant glycosylation. This includes decreased Na+ current amplitude, altered voltage

[email protected] 100 J. Lazniewska and N. Weiss dependence of activation and inactivation and slower channel inactivation kinetics. Although the arrhythmic phenotype in those genetically modiﬁed animals can be caused by various cellular defects, these results also suggest that defects in glycosylation of voltage-gated Na+ channels can possibly contribute to the development of cardiac arrhythmias.

7.2 Diabetic Neuropathic Pain

2+ It is well documented that Cav3.2 voltage-gated Ca channels contribute to peripheral neuropathic pain by sensitising primary nociceptors (Dogrul et al. 2003; Bourinet et al. 2005; Gadotti et al. 2013; Francois et al. 2014). In addition, an increased expression of Cav3.2 channels in primary nociceptive neurons has been documented in response to hyperglycaemia during diabetes, which in turn causes painful peripheral neuropathy (Todorovic and Jevtovic-Todorovic 2014). Interestingly, elevation of external glucose concentration in a range consistent with diabetic hyperglycaemia induced increase expression of Cav3.2 channels expressed in mammalian cell lines and depends on proper N-glycosylation of the channel (Fig. 15a) (Weiss et al. 2013). In addition, the application of neuraminidase on DRG neurons from ob/ob mice, an animal model of peripheral neuropathy of type 2 diabetes and obesity mouse model of obesity/diabetes (Drel et al. 2006), showed reduced T-type Ca2+ currents, and topically the application of neuraminidase in vivo in ob/ob mice was shown sufﬁcient to prevent T-type-induced painful diabetic neuropathy (Fig. 15b) (Orestes et al. 2013). Hence, it is unambiguous that hyperglycaemia-related Cav3.2 hyperactivity is mediated by channel glycosylation and has profound implication in the pathogenesis of diabetic neuropathic pain.

7.3 Cystic Fibrosis

Cystic fibrosis, also known as mucoviscidosis, is an autosomal recessive disorder caused by mutations in the gene encoding for the CFTR channels, affecting essentially the lungs but also the pancreas, liver and intestine (Guggino and Stanton 2006). Although more than 1,000 mutations in the CFTR channel have been identified in patients with cystic fibrosis, the deletion of the phenylalanine residue at position 508 (ΔF508) is the most common one (Van Goor et al. 2006). While the wild-type CFTR exists in three different glycosylation forms – non-glycosylated, core glycosylated or fully mature complex glycosylated (Gregory et al. 1990) – the mutant CFTR ΔF508 is predominantly expressed as non-glycosylated and core- glycosylated channel and is unable to traffic efficiently to the Golgi apparatus when expressed at physiological temperature (37C) (Cheng et al. 1990; Gregory et al. 1991). However, decreasing temperature (26C) is sufficient to restore expression of fully mature, complex glycosylated channels and partial surface

[email protected] The “Sweet” Side of Ion Channels 101 expression (Denning et al. 1992). In addition, determination of the oligosaccharide structures of both mutant and wild-type CFTR also revealed signiﬁcant differences (O’Riordan et al. 2000; Scanlin and Glick 2001). Altogether, alteration of CFTR glycosylation by cystic ﬁbrosis mutation likely contributes to the pathogenesis of the disease and possibly represents a potential target for the development of new therapies.

7.4 Varitint–Waddler Phenotype

Mutations in the transient receptor potential channel TRPML3 cause the varitint– waddler phenotype, which is characterised by pigmentation defect, hearing loss, circling behaviour and embryonic lethality (Di Palma et al. 2002; Atiba-Davies and Noben-Trauth 2007; Cuajungco and Samie 2008). Interestingly, the TRPML3 A419P mutation, associated with a severe phenotype, induces massive cell death when expressed in mammalian cells (Kim et al. 2007). While membrane expression of the mutant channel is not signiﬁcantly altered, it shows a constitutive activity and altered glycosylation state. Although the exact molecular mechanism by which the A419P mutation alters channel glycosylation remains elusive, it is likely that constitutive TRPML3 activity arises from aberrant channel glycosylation. More- over, aberrant glycosylation may also alter proper targeting of TRPML3 into intracellular organelles that may contribute to diseased state.

Conclusion and Perspectives Protein glycosylation is rapidly emerging as a fundamental post-translational mechanism to control the properties and function of ion channels. To date, up to 30 different ion channels and auxiliary subunits have been biochemically and/or functionally documented at the glycosylation level. At the time we are writing this review, glycosylation of new channels is being documented, like the new volume-regulated anion channel (VRAC) LTTC8A (Voss et al. 2014), and it is expected that the catalogue will grow significantly in the next years. Few other channels including the IP3 receptor (Michikawa et al. 1994) and TMEM16 (Hartzell et al. 2009) not discussed in the current review have also been identified as glycosylated, and further analysis will certainly reveal the physiological importance. It is now explicit that protein glycosylation controls multiple aspects of ion channel life cycle from gene expression, sorting and trafficking, gating and function (Fig. 16). However, there are no general rules regarding the functional influence of glycan on the channel behaviour, and even closely related channel subtypes can be differentially affected by glycosylation. Moreover, although it is unambiguous that glycosylation controls various aspects of ion

(continued)

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Fig. 16 Glycosylation controls ion channel life cycle. The control of ion channel function by glycosylation can occur at various levels: i) modulation of ion channel gene expression by glycosylation of transcription factors; (ii) control of channel maturation, sorting and trafficking and cellular localisation; (iii) channel activity (gating, stability); (iv) modulation of channel function by glycosylation of auxiliary subunits; and (v) modulation by cellular signalling pathway (e.g. Klotho and other glycosidases). Moreover, various external factors including level and type of sugar and temperature also influence glycosylation and glycan composition and have a significant impact on ion channel function

channels, the molecular mechanisms by which addition of a glycan on the polypeptide chain eventually contributes to the localisation and function of the mature channel embedded in the cell membranes remain largely unknown. Although it is believed that the presence of terminal sialic acid residues in the glycan tree contributes to the surface electrical potential that in turn affects the channel voltage sensor in the case of voltage-gated channels, the chemical composition of the glycan itself is also of fundamental importance in modulating channel properties and localisation. Hence, glycan composition appears to be an incredible molecular structure where information relative to ion channel expression and function is encoded. Structural insight into channel glycosylation, combined with functional analysis, will certainly provide important information on how glycan trees control speciﬁc channel features.

(continued)

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Protein glycosylation also represents a powerful mechanism to ﬁne-tune channel activity and function in a cell-type- and loci-dependent manner. A second level of complexity arises from the emerging concept that protein glycosylation not only contributes to deﬁne the cellular behaviour of the channel but also represents an important determinant of channel regulation by other signalling pathways. On the other way around, it is likely that channel glycosylation contributes to crosstalk regulations with other signalling pathways. This reveals the existence of regulation networks related with channel glycosylation, and a major goal is now to elucidate the many players to fully apprehend all aspects of ion channel regulation and physiology by glycosylation. Finally, while the importance of glycosylation in controlling diverse aspects of a single ion channel molecule is unambiguous, very little is known about the functional consequences at the cellular and body levels. It is clear that alteration of protein glycosylation can lead to sever disorders. However, the implication of ion channels remains largely unexplored. More- over, protein glycosylation shares many features with protein phosphorylation, such that proteins can be both glycosylated and deglycosylated. Obtaining information on the cellular signals and enzymes that spatiotempo- rally regulate channel glycan trees will be critical to fully decipher the (patho) physiological role of glycosylation. Certainly, the most important challenge of the next few years will be to reveal the functional consequence of ion channel glycosylation from the single molecule to the whole organism. It will not only contribute to our general understanding of ion channel physiology but also possibly uncover new aspects of channelopathies and glycopathies in general that may lead to novel therapeutic avenues.

Acknowledgements This work was supported by internal funding from the Institute of Organic Chemistry and Biochemistry (IOCB). JL is supported by a postdoctoral fellowship from IOCB.

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