Gap junction channel structure in the early 21st century: facts and fantasies
Introduction
Gap junctions are specialized regions of cell-to-cell contact at which hexameric oligomers, called connexons, dock end-to-end noncovalently across a narrow extracellular gap. Hundreds to thousands of channels cluster in so-called plaques, and the individual channels allow exchange of nutrients, metabolites, ions, and small molecules of up to ≈1000 Da [1]. Coupling by gap junctions is a fundamental mechanism for cell-to-cell communication in higher organisms. More than 20 connexin isoforms have been identified to date in deuterostomes, from sea urchins to humans [2, 3].
Each connexon, or hemichannel, is an annular assembly of six individual connexins that forms a pore through the plasma membrane. The different connexin isoforms can interact structurally in various ways. Connexons may be homomeric or heteromeric, and junctional channels may be formed by connexons having the same or different compositions. The expression of multiple connexins in the same cell type, the multiplicity of isoforms, as well as their different structural combinations, probably provides exquisite ‘functional tuning’ of this unique family of membrane channels.
The primary tools for structure analysis of gap junction channels include electron microscopy and image analysis [4, 5, 6, 7, 8, 9••], X-ray diffraction [10, 11, 12], nuclear magnetic resonance (NMR) spectroscopy [13•, 14, 15] and atomic force microscopy (AFM) [16, 17•, 18•]. Mutagenic, biochemical, and electrophysiological approaches have also been used to elucidate the structure–function relationships of gap junction channels. This review focuses on recent studies that illuminate the structure of connexin channels, drawing on maps derived by electron cryo-crystallography and on structurally focused mutagenesis and electrophysiological studies. The reader is also referred to reviews by Yeager and Nicholson [19], Harris [20], Sosinsky and Nicholson [21], and Kovacs et al. [22].
Section snippets
The connexon contains a ring of 24 α-helices
Hydropathy and topological analyses of various connexins suggest that each contains four transmembrane domains, referred to as M1, M2, M3, and M4, proceeding from the N-terminus to the C-terminus [23]. Connecting the transmembrane domains are two extracellular loops (E1, connecting M1–M2 and E2, connecting M3 to M4) and one cytoplasmic M2-M3 loop. Both the N-termini and C-termini reside in the cytoplasm [23, 24, 25]. The transmembrane domains and the extracellular loops display the highest
Regions in NT, M1, E1, and/or M3 have been implicated in lining the pore
Several domain swap studies showed that the single channel conductance of connexin pores is a property that can be transferred between channels by exchange of M1, particularly its second half (Cx46, Cx37, and Cx32 [38, 39]). Other domain swap studies showed that the charge selectivity of connexin pores can be controlled by E1 (Cx46 and Cx32 [40]), suggesting that E1 contributes to the pore wall. Point mutations in the NT produced changes in the single channel current–voltage relations
A Cα model suggests that mutations that disrupt helix–helix packing interfere with channel function
Clearly, an essential challenge is to utilize the existing 3D cryoEM map and the existing mutagenesis, physiological, and amino acid sequence data to reach a consenus about which parts of which domains line the pore. The key difficulties are that the map is of necessity a snapshot of a single structural state, and it may not correspond to the dominant state probed by the mutagenesis/physiological studies. For these reasons, it is perhaps unrealistic to expect the two sets of data to be entirely
The N-terminus may form a plug that blocks Cx26 channels
NMR spectroscopy of a 13-residue peptide corresponding to the N-terminal domain of Cx26 displayed a two-turn α-helix, which then unraveled into a flexible loop-like structure [35]. It was hypothesized that this short NT helix is oriented parallel to the transmembrane helices lining the entrance to the pore, thus forming part of the conduction path and contributing to the voltage dependence of the channel. Support for this model is suggested by recent cryoEM studies of the M34A mutant of Cx26 [9
Conclusions
The last decade has seen impressive progress in the analysis of several classes of membrane proteins, including reaction centers, porins, ligand-gated channels, voltage-gated channels, transporters, and aquaporins [50]. By comparison, the tempo of discovery in the gap junction channel field has been slower. Possible reasons include difficulties with expression of engineered connexins with sufficient stability and quantity to allow detailed biochemical and biophysical analysis, difficulties in
References and recommended reading
Papers of particular interest, published within the annual period of review, have been highlighted as:
• of special interest
•• of outstanding interest
Acknowledgements
This work was supported by NIH grants RO1HL48908 (MY), RO1GM36044 (ALH), and RO1NS056509 (ALH). We thank Julio Kovacs, Kenton Baker, and Michael E Pique for preparation of Figure 1, Figure 2, Figure 3, respectively. We thank Atsunori Oshima and Yoshinori Fujiyoshi for providing Figure 4.
References (51)
- et al.
Connexin disorders of the ear, skin, and lens
Biochim Biophys Acta
(2004) - et al.
Formation of the gap junction intercellular channel requires a 30° rotation for interdigitating two apposing connexons
J Mol Biol
(1998) - et al.
A Cα model for the transmembrane α helices of gap junction intercellular channels
Mol Cell
(2004) - et al.
Gap junction structures. II. Analysis of the X-ray diffraction data
J Cell Biol
(1977) - et al.
pH-dependent dimerization of the carboxyl terminal domain of Cx43
Biophys J
(2004) - et al.
Conformational changes in surface structures of isolated connexin 26 gap junctions
EMBO J
(2002) - et al.
Aminosulfonate modulated pH-induced conformational changes in connexin26 hemichannels
J Biol Chem
(2007) - et al.
Structure and biochemistry of gap junctions
- et al.
Structural organization of gap junction channels
Biochim Biophys Acta
(2005) - et al.
Topology of the 32-kD liver gap junction protein determined by site-directed antibody localizations
EMBO J
(1988)