α-Helical transmembrane peptides: A “Divide and Conquer” approach to membrane proteins

Chemistry and Physics of Lipids 163 (2010) 1–26

Contents lists available at ScienceDirect

Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip

Invited review

␣-Helical transmembrane peptides: A “Divide and Conquer” approach to membrane proteins Natalie Bordag, Sandro Keller ∗ Leibniz Institute of Molecular Pharmacology FMP, Robert-Rössle-Str. 10, 13125 Berlin, Germany

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Article history: Received 19 March 2009 Received in revised form 21 July 2009 Accepted 21 July 2009 Available online 13 August 2009 Keywords: Detergents Helix–helix interactions Membrane-mimetic systems Peptide–lipid interactions Two-stage model Vesicles

a b s t r a c t ␣-Helical membrane proteins fulfill many vital roles in all living cells and constitute the majority of drug targets. However, their relevance is in no way paralleled by our current understanding of their structures and functions. This is because membrane proteins present a number of experimental obstacles that are difficult to surmount by classical methods developed for water-soluble proteins. Moreover, membrane proteins are not only challenging on their very own but, when embedded in a biological membrane, also reside in an outstandingly complex milieu. These difficulties have fostered a “divide and conquer” approach, in which a membrane protein is dissected into shorter and easier-to-handle transmembrane (TM) peptides. Under suitable conditions, such peptides fold independently and retain many of the properties displayed in the context of the full-length parent protein. This contribution reviews some of the most notable insights into ␣-helical membrane proteins gleaned from experiments on protein-derived TM peptides. We recapitulate some peculiar properties of lipid bilayers that render them such a complex and unique environment and discuss generic features pertaining to hydrophobic peptides derived from ␣-helical membrane proteins. The main part of the review is devoted to a critical discussion of particularly interesting examples of TM peptides studied in membranemimetic systems of increasing complexity: isotropic solvents, detergent micelles, lipid bilayers, and biological membranes. The unifying theme is to explore to what extent TM peptides in combination with different membrane-mimetic systems can aid in advancing our knowledge and comprehension of ␣-helical membrane proteins as well as in developing new pharmacological tools. © 2009 Elsevier Ireland Ltd. All rights reserved.

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lipid bilayers and ␣-helical membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Structural hallmarks of lipid bilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abbreviations: A2 aR, adenosine A2 a receptor; ATR-FTIR, attenuated total reflection Fourier transform infrared spectroscopy; AUC, analytical ultracentrifugation; ␤2 AR, ␤2 adrenergic receptor; BLM, black lipid membrane; CR, conductance recordings (of vesicles, lipid bilayers, or cells); CD, circular dichroism; CMC, critical micellar concentration; DHepPC, 1,2-diheptanoyl-sn-glycero-phosphocholine; DLS, dynamic light scattering; DMPC, 1,2-dimyristoyl-sn-glycero-phosphocholine; DMSO, dimethyl sulfoxide; DMT1, divalent metal transporter 1; DOPC, 1,2-dioleoyl-sn-glycero-phosphocholine; DOSY, diffusion-ordered spectroscopy; DPC, dodecylphosphocholine; DPPC, 1,2-dipalmitoylsn-glycerophosphocholine; DSC, differential scanning calorimetry; eggPC, phosphatidylcholine extracted from hen egg; EPR, electron paramagnetic resonance; ErbB or EGFR, epidermal growth factor receptor; FGFR3, fibroblast growth factor receptor 3; Fl, fluorescence-based methods; FRET, fluorescence (or Förster) resonance energy transfer; GALLEX, TM-heteromer-sensitive assay based on LexA operator; GlyR, glycine receptor; GpA, glycophorin A; GPCR, G-protein-coupled receptor; HFIP, 1,1,1,3,3,3hexafluoro-2-propanol; ITC, isothermal titration calorimetry; KcsA, K+ -channel from Streptomyces lividans; CLSM, confocal laser scanning microscopy; MCP13, major coating protein 13; MCP-II, Escherichia coli methyl-accepting chemotaxis protein II; MD, molecular dynamics; MDCK, Madin–Darby canine kidney; Mi, microscopy; NMR, nuclear magnetic resonance; NOE, nuclear Overhauser effect; Nramp1, natural resistance-associated macrophage protein 1; OCD, oriented CD; pHLIP, pH low insertion peptide; PAGE, polyacrylamide gel electrophoresis; PLB, phospholamban; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-phosphocholine; POPE, 1-palmitoyl-2-oleoyl-sn-glycero-ethanolamine; POPG, 1-palmitoyl-2-oleoyl-sn-glycero-phosphoglycerol; POSSYCAT, selectable TOXCAT-based assay; RMSD, root mean square deviation; RTK, receptor tyrosine kinase; SCoVE, protein E from Coronaviridae; SDS, sodium dodecyl sulfate; SEC, size-exclusion chromatography; Slc11a1, solute carrier family 11 member 1; SPR, surface plasmon resonance; Ste2p, pheromone ␣-factor receptor from Saccharomyces cerevisiae; SUV, LUV, GUV, small, large, giant unilamellar vesicle, respectively; TFE, trifluoroethanol; TM, transmembrane; TOXCAT, ToxR with chloramphenicol acetyl transferase; ToxR, dimerization-dependent membrane-spanning transcriptional activator of ctx promoter from Vibrio cholerae. ∗ Corresponding author. Tel.: +49 30 9406 3064; fax: +49 30 9406 3065. E-mail address: [email protected] (S. Keller). 0009-3084/$ – see front matter © 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.chemphyslip.2009.07.009

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2.1.1. Distribution probability of chemical moieties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. Charge density profile . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. Lateral pressure profile . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Folding of ␣-helical membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1. Two-stage model of membrane protein folding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2. Structure induction in lipid membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.3. Forces driving helix–helix interactions in membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . TM peptides derived from ␣-helical membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Generic features of TM peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Peptide sources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Peptide length and hydrophobic mismatch . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Flanking residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1. Types and purposes of flanking residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2. Effects of flanking residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isotropic solvents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Structure and oligomerization of TM peptides in organic solvents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.1. Secondary structure induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.2. Structures of bacteriorhodopsin fragments in DMSO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.3. Influence on helix–helix interactions in solute carrier family 11 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Solubility of TM peptides in aqueous solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1. Difficulties due to ␤-sheet aggregation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2. pHLIP in aqueous solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Partitioning scales based on isotropic solvents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1. Whole-residue partitioning scale . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.2. Suitability of octanol for determining partitioning scales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Detergent micelles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Structural impact of detergent micelles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1. TM helix 2 of ␣1 glycine receptor: influence of micellar environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.2. TM helix 2 of ␣1 glycine receptor: comparison with other membrane-mimetic systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.3. TM helix 2 of ␣1 glycine receptor: pentamer model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Oligomerization of TM peptides in detergent micelles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.1. Influence of detergents on glycophorin A dimerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.2. Oligomerization of TM helix 4 of solute carrier family 11 member 2 in SDS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lipid bilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. TM peptide dimers in lipid vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.1. Helix–helix interactions in the glycophorin A dimer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Energetics of TM peptide self-insertion into lipid vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.1. Stoichiometric binding model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.2. Partitioning model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Planar bilayers as hosts for TM peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3.1. Bicelles in TM peptide research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3.2. TM peptide orientation in multibilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3.3. TM helix 2 of ␣1 glycine receptor: channel activity in spanned bilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Quantification of helix–helix interactions in cell culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.1. Cell-based quantification of oligomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Pharmacological applications of TM peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.1. Interference with membrane protein function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.2. Reconstitution of channel activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.3. Delivery of pharmacological tools using TM peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References cited in Table S1 (Supplementary Information) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix A. Supplementary data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction ␣-Helical membrane proteins are of paramount interest to both basic and applied research. They fulfill numerous and diverse functions, including water and solute transport, signal transduction and integration, enzymatic activity, and anchorage of cytoskeletal or extracellular components. Membrane proteins account for about one third of all proteins encoded in the human genome (Wallin and von Heijne, 1998) and make up more than half of all current drug targets (Overington et al., 2006). Consequently, enormous efforts have been exerted for decades to deepen our knowledge and understanding of the structures and functions of membrane proteins.

4 4 4 4 4 5 5 6 6 6 6 6 6 6 7 7 7 8 8 8 9 9 9 9 9 10 11 11 12 12 12 12 12 13 13 13 14 14 14 15 15 15 15 15 15 16 16 16 17 17 17 18 18 18 18

Notwithstanding some remarkable successes, overall progress in the field has remained disappointingly slow, especially when compared with the tremendous advances in the study of water-soluble proteins. This disparity is best exemplified by the fact that, as of 4 September 2009, only 203 unique high-resolution structures of membrane proteins have been deposited in the Protein Data Bank (see http://blanco.biomol.uci.edu/Membrane Proteins xtal.html), whereas the structures of several 10,000 water-soluble proteins are known to date. The dearth of information on membrane proteins is due to two major obstacles. On the one hand, many membrane proteins are cytotoxic when overexpressed in cell culture (Junge et al., 2008).

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Thus, they are notoriously difficult to obtain in quantities and purities sufficient for biophysical, biochemical, or structural methods. On the other hand, membrane proteins are hydrophobic and usually require the presence of colloidal additives like detergents or lipids during purification and experimentation. This renders them hard to handle and even unsuitable for many techniques. Most importantly, however, many membrane proteins will fold into their native structures and exert their biological functions only in specific, often complex membranes or membrane-mimetic environments. The present review is devoted to what is sometimes referred to as a “divide and conquer” approach to membrane proteins (White et al., 2001), which is one among numerous strategies devised to circumvent the above problems. The underlying rationale is to dissect an ␣-helical membrane protein into its constituent transmembrane (TM) domains, which can normally be produced more easily and be investigated in a wider range of membrane-mimetic systems than their full-length parent protein. For the sake of simplicity, we will henceforth refer to peptides that correspond to or have been derived from TM sequences of single- or multispanning ␣-helical membrane proteins as TM peptides, domains, or sequences. Some important peptide classes comprise TM peptides but are not dealt with here because they have been the subject of excellent recent reviews. These include model peptides with artificial sequences (Wimley and White, 2000; Killian and Nyholm, 2006; Nyholm et al., 2007; Marsh, 2008), antimicrobial peptides (Som et al., 2008; Khandelia et al., 2008), peptidic venoms (Bechinger and Lohner, 2006; Raghuraman and Chattopadhyay, 2007), fusion peptides (Epand, 2003; Lins et al., 2008), and cell-penetrating peptides (Herbig et al., 2007; Lins et al., 2008). Table S1 in the Appendix provides an extensive, though certainly not exhaustive, compilation of over 300 publications devoted to TM peptides. However, the goal of this review is not to reiterate all of these findings. Instead, we focus on few examples that are particularly well-suited to demonstrate what studies on TM peptides have contributed to our understanding of ␣-helical membrane proteins, thereby putting special emphasis on the roles played by various membrane-mimetic systems. We start with brief summaries of the complex nature of lipid bilayers (Chapter 2) and of some generic properties of TM peptides (Chapter 3). Then, selected examples of TM sequences are discussed in order of increasing complexity of the membrane-mimetic systems used, ranging from isotropic solvents (Chapter 4) and detergent micelles (Chapter 5) to lipid bilayers (Chapter 6) and biological membranes (Chapter 7). The review concludes with a discussion of the potential and limitations of the “divide and conquer” approach and suggestions for future research efforts using TM peptides (Chapter 8). 2. Lipid bilayers and ␣-helical membrane proteins Biological membranes are intricate supramolecular assemblies of lipids and (glyco)proteins, whose protein content may vary from <20% to >90% (w/w) (Helms, 2002). This chapter highlights two aspects that are fundamental to understanding and interpreting experiments on TM peptides in membrane-mimetic systems: the structural complexity inherent to lipid bilayers (Section 2.1) and the folding process of ␣-helical membrane proteins in such a complex environment (Section 2.2). 2.1. Structural hallmarks of lipid bilayers The structures and dynamics of lipid bilayers have been reviewed in depth elsewhere (e.g., Wiener and White, 1992; White and Wimley, 1998; White et al., 2001; van Meer et al., 2008).

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Fig. 1. Heterogeneity and anisotropy of a lipid bilayer as reflected in transbilayer profiles of (A) distribution probabilities of chemical moieties; (B) charge density; and (C) lateral pressure. (A) Gaussian probability distribution functions representing phospholipid moieties and water in a hydrated 1,2-dioleoyl-snglycero-phosphocholine (DOPC) bilayer. The area under each curve is proportional to the number of constituent groups per DOPC molecule (e.g., one phosphate, two carbonyls, and so on). Figure courtesy of Stephen White, adapted from White and von Heijne (2008), with permission of Annual Reviews. (B) Charge density profile, given as absolute partial charge density across a DOPC bilayer computed from atomic partial charges and group volumes (Wiener and White, 1992; White and Wimley, 1998, 1999). Figure courtesy of Stephen White, adapted from White and von Heijne (2008), with permission of Annual Reviews. (C) Lateral pressure profile of a fluid 1-palmitoyl-2-oleoyl-sn-glycerophosphocholine (POPC) bilayer calculated from a 10.7-ns MD simulation of 128 POPC molecules at 310 K. Figure courtesy of Justin Gullingsrud, adapted from Gullingsrud and Schulten (2004), with permission of Elsevier.

Here we restrict ourselves to emphasizing the complex, anisotropic architecture of even the simplest lipid bilayers by illustrating the transbilayer profiles of the distribution probabilities of phospholipid moieties, the charge density, and the lateral pressure (Fig. 1). On a quantitative scale, such profiles will, of course, depend on parameters such as temperature, pressure, and, most importantly, lipid composition. For example, lipid headgroups vary in size, charge, extent of hydration, and number of hydrogen bond donors and acceptors, whereas acyl chains differ in length, degree of saturation, and branching.

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Fig. 2. Two-stage model of membrane protein folding. In step 1, a hydrophobic peptide sequence forms an independently stable ␣-helical TM domain upon release from the translocon or self-insertion from a non-TM state. In step 2, membrane-embedded helices interact with each other to establish tertiary and quaternary structure. Helical backbones and final quaternary structure taken from MD simulations (Sansom et al., 2008; PDB file downloaded from http://sbcb.bioch.ox.ac.uk/cgdb/ , with permission of Mark Sansom) based on the crystal structure of the homotetrameric TM domain of influenza A virus protein M2 (PDB 3bkd; Stouffer et al., 2008a). 1,2-Dipalmitoyl-sn-glycerophosphocholine (DPPC) bilayer taken from MD simulations (Tieleman and Berendsen, 1996; PDB file downloaded from http://moose.bio.ucalgary.ca , with permission of Peter Tieleman). This figure is not meant to imply that assembly of the M2 proton channel proceeds as depicted but merely serves to illustrate the generic features of the two-stage model. M2 helices are colored in different shades of blue. DPPC acyl tails are depicted in black, headgroup glycerols in slate gray, phosphate groups in red and orange, and choline N+ (CH3 )3 moieties in blue and white.

2.1.1. Distribution probability of chemical moieties Using X-ray and neutron diffraction, White and co-workers (Wiener and White, 1992; White and Wimley, 1998; White et al., 2001) determined the distribution probabilities of different chemical moieties of DOPC and water along the bilayer normal (Fig. 1A). Each of the two headgroup regions is ∼15 Å thick, whereas the hydrophobic core is ∼30 Å in thickness. However, the boundaries separating the headgroup regions from the hydrocarbon core are not sharp. Instead, the distribution probabilities are wellrepresented by rather broad Gaussian curves that largely overlap, reflecting the gradual change in chemical composition along the bilayer normal. Water penetrates the bilayer to the level of the carbonyl groups, which form the boundary between the headgroup regions and the hydrocarbon core. The large thermal disorder of a fluid-state lipid bilayer allows the double bonds and the methyl groups of the acyl chains to explore the hydrated interface. 2.1.2. Charge density profile From the above distribution probabilities, it follows that also the charge density and the relative static permittivity (or relative dielectric constant) change gradually as a function of bilayer depth (Fig. 1B). At room temperature, the relative static permittivity amounts to ∼80 in bulk water but drops to ∼12–26 in the headgroup region (Ohki and Arnold, 1990; Ohki and Zschörnig, 1993) and further to ∼2 in the center of the hydrocarbon core (Popot and Engelman, 2000; Jayasinghe et al., 2001). This results in decreased charge density as well as stronger electrostatic forces, which are inversely proportional to the permittivity of the medium, within the bilayer core. 2.1.3. Lateral pressure profile The term “lateral pressure profile” was coined by Xiang and Anderson (1994), and its implications to membrane proteins were later discussed by Cantor (1997). With the aid of molecular dynamics (MD) simulations, Gullingsrud and Schulten (2004) calculated the lateral pressure profile of a POPC bilayer (Fig. 1C). Strong lateral pulling forces act on both sides of a lipid bilayer such as to minimize the contact area between the hydrophilic headgroup regions and the hydrophobic hydrocarbon core. These pulling forces are counteracted by steric and electrostatic repulsion and loss of hydration water in the headgroup regions as well as steric repulsion and

reduced acyl chain flexibility in the hydrocarbon core. At mechanical equilibrium, that is, when the bilayer neither expands nor contracts, the pressure integral along the bilayer normal vanishes, but considerable positive or negative local lateral pressures are exerted as a function of bilayer penetration depth. The lateral pressure profile has been suggested to affect helix–helix interactions and membrane protein conformational transitions that are accompanied by shape changes (Cantor, 1997; van den Brink-van der Laan et al., 2004b; Marsh, 2007, 2008). However, experimental evidence supporting such a fundamental role of the lateral pressure profile has thus far remained scarce and ambiguous (van den Brink-van der Laan et al., 2004a; Duong-Ly et al., 2005). 2.2. Folding of ˛-helical membrane proteins A major driving force in the folding of water-soluble proteins is the hydrophobic effect (or hydrophobic collapse). Obviously, such a mechanism cannot be at play in the folding of membrane proteins, as they are largely embedded in a hydrophobic environment. However, this is only one out of several peculiarities of membrane protein folding, some of which are discussed in the following sections. 2.2.1. Two-stage model of membrane protein folding According to the two-stage hypothesis (Popot et al., 1987; Jacobs and White, 1989; Popot and Engelman, 1990), folding of multispanning ␣-helical membrane proteins proceeds in two separable steps (Fig. 2). In a first step, hydrophobic peptide sequences form independently stable ␣-helical TM domains on bilayer insertion. In a second step, membrane-embedded helices interact with each other (see Section 2.2.3 and Fig. 2) to establish the protein’s tertiary structure. Although simplistic, this hypothesis (Popot et al., 1987) has found support from both protein fragmentation studies (e.g., Furthmayr and Marchesi, 1976; Kovacs et al., 1988; Kahn and Engelman, 1992; Ridge et al., 1995; Corbin et al., 1998) and thermodynamic arguments (Popot et al., 1987; Jacobs and White, 1989; Popot and Engelman, 1990). Refinements of the two-stage model include extensions to account for cofactor binding and structural rearrangement on helix–helix interactions (Engelman et al., 2003) or self-insertion of water-soluble TM sequences (White and Wimley, 1999). In the latter scenario, an initially unstructured peptide first parti-

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tions from the aqueous solution into the membrane interface to form an ␣-helix, which then flips into a TM orientation. Most TM sequences, however, are neither water-soluble nor capable of selfinsertion; instead, their bilayer insertion depends on the activity of a sophisticated membrane protein complex called translocon (reviewed by White and von Heijne, 2005). Briefly, a nascent polypeptide chain partly folds within the ribosome exit tunnel and the translocon channel, from which single TM helices are released sideways into the bilayer (reviewed by White and von Heijne, 2005; Bowie, 2005). Although the situation in vivo is far more complex than self-insertion of some TM peptides observed in vitro, both processes have in common that formation of independently stable TM ␣-helices precedes interactions among bilayer-spanning sequences. The significance of the two-stage model lies in the conceptual and experimental disconnection of TM helix formation and helix–helix interactions within the bilayer. Accordingly, an isolated TM helix constitutes a distinct folding domain that can be separated from the remainder of its parent protein without losing its inherent structural properties. This hypothesis rationalizes the “divide and conquer” approach to membrane proteins and thus founds the theoretical basis of all studies employing TM peptides (White et al., 2001). 2.2.2. Structure induction in lipid membranes The distribution probabilities of lipid moieties and the permittivity profile along the bilayer normal (see Fig. 1 and Section 2.1) are paralleled by preferential occurrence of different amino acid residues in a TM peptide. Whereas the central, hydrocarbonexposed part of a TM sequence contains predominantly apolar amino acids (Ala, Ile, Leu, Val, Phe), polar or charged (Arg, Asp, Asn, Glu, Gln, Lys) residues are located preferentially in flanking regions, where they interact with lipid headgroups and water. Residues comprising both polar and apolar moieties (Trp, Tyr, His) are often located in the boundaries between the two headgroup regions and the hydrophobic core and thus help anchoring the peptide in a TM orientation (Yau et al., 1998; Ulmschneider et al., 2005; von Heijne, 2006). The question then remains as to why an initially unstructured peptide adopts an ␣-helical structure on bilayer insertion. This is explained by the low permittivity and the strongly reduced number of solvent hydrogen bond donors and acceptors in the hydrophobic membrane core as compared with an aqueous environment. Structure induction and stabilization result from the requirement that all backbone carbonyl oxygens and amide protons be intrahelically hydrogen-bonded to reduce the energetic cost of exposing unsatisfied hydrogen bond donors and acceptors to the apolar membrane core (White and Wimley, 1999). This requirement can best be met by satisfying all backbone hydrogen bond donors and acceptors in an ␣-helical secondary structure (or, for some proteins, in a ␤-sheet barrel). For instance, disrupting all backbone hydrogen bonds of a 20-residue TM domain would be accompanied by a prohibitively high free energy penalty of ∼330 kJ/mol (White et al., 2001).

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Fig. 3. Helix–helix interactions mediated by tight knobs-into-holes packing in membrane proteins exemplified by a monomeric subunit taken from the crystal structure of the tetrameric potassium channel KcsA from Streptomyces lividans (PDB 1K4C; Zhou et al., 2001). Van der Waals forces contribute considerably to such helix–helix interactions because of the large contact surface. The position of the lipid bilayer is indicated by a yellow slab.

2.2.3. Forces driving helix–helix interactions in membranes Helix–helix interactions within lipid bilayers dictate the stability and specificity of both intramolecular interactions in multispanning membrane proteins (i.e., tertiary structure) and intermolecular interactions in membrane protein complexes (i.e., quaternary structure; for an example see Fig. 3). In the absence of a noticeable contribution from hydrophobic collapse, other forces gain more importance:

knobs-into-holes packing of side chains at interhelical interfaces (Fig. 3). Amino acid motifs containing small side chains (Gly, Ala, Ser, Thr) separated by three other, often ␤-branched, residues play an important role in this context (reviewed by Senes et al., 2004; White, 2005; MacKenzie, 2006; Rath et al., 2007, 2009; MacKenzie and Fleming, 2008). (ii) Interhelical hydrogen bonds of the type O–H···O or N–H···O between amino acid side chains occur less frequently in membrane proteins than in water-soluble proteins. However, numerous, though substantially weaker, C␣ –H···O hydrogen bonds involving the peptide backbone have been observed (Senes et al., 2001; Adamian and Liang, 2002; Choi et al., 2005; Bocharov et al., 2007; Rath et al., 2009). Although the strength of individual C␣ –H···O bonds is still under debate, their occurrence in literally interhelical hydrogen bond networks implies that they contribute to stability and specificity of helix–helix interactions (Senes et al., 2001; Adamian and Liang, 2002; Dawson et al., 2003). Amino acids often involved in such hydrogen bond networks include Gly, Ser, Thr, Tyr, His (Senes et al., 2001; Dawson et al., 2002; Adamian and Liang, 2002) as well as Asp, Asn, Glu, Gln (Dawson et al., 2003; Tatko et al., 2006). (iii) Salt bridges occur sparingly in membrane proteins, supposedly because they contribute modestly to stability (White and Wimley, 1999) and lack specificity (Schneider, 2004). Thus, in the relatively rare cases where charged side chains are found within the hydrophobic core, these residues are usually involved in helix–helix interactions (Ulmschneider et al., 2005; von Heijne, 2006), cause bilayer distortion and water penetration (MacCallum et al., 2007), or undergo snorkeling into the headgroup region (Senes et al., 2007). Nevertheless, buried polar and charged residues play important functional roles in membrane proteins such as some ion channels (e.g., influenza A virus protein M2, see Section 5.2).

(i) Van der Waals forces contribute substantially to both stability and specificity. They are enhanced by geometrically complementary surfaces of interacting TM domains and tight

Overall, the tertiary and quaternary structures of many membrane proteins are only marginally stable because they need to strike a fine balance between structural stability and conforma-

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tional flexibility in order to carry out their biological functions. Parameters to which membrane proteins may respond by conformational adaptations include interactions with other proteins, peptides, or small-molecule ligands, changes in membrane electrostatics or lipid composition, and alterations in the lateral pressure profile (see Section 2.1). 3. TM peptides derived from ␣-helical membrane proteins Before looking into specific examples in the following chapters, we first outline some generic features of TM peptides (Section 3.1) and then discuss the influence of hydrophilic flanking residues on hydrophobic sequences (Section 3.2). 3.1. Generic features of TM peptides 3.1.1. Peptide sources TM peptides can be obtained either by proteolytic cleavage of naturally occurring or recombinantly engineered proteins or by means of chemical synthesis. Some of the first TM peptides were produced by tryptic fragmentation of full-length glycophorin A (GpA; Furthmayr and Marchesi, 1976). Chemically synthesized peptides followed soon thereafter (Galardy and Kortylewicz, 1985) and, thanks to the tremendous advances in solid-phase peptide synthesis achieved during the past decades, dominate nowadays. However, chemical synthesis of exceptionally long or isotopically labeled peptides often remains difficult or prohibitively expensive, respectively. In such cases, an alternative way of obtaining TM peptides is through biosynthesis in cell culture. To minimize cytotoxicity and facilitate expression and purification, TM peptides may be produced as fusion constructs with other proteins, which can later be removed by proteolytic cleavage (e.g., Adair and Engelman, 1994; Sharpe et al., 2000). 3.1.2. Peptide length and hydrophobic mismatch Most TM peptides range in length from around 20 to roughly 30 residues (see Table S1 for examples). However, these numbers represent typical values rather than sharp limits. For instance, the shortest model peptide found to assume a stable TM topology contains only 11 residues (Krishnakumar and London, 2007). In the other extreme, some peptides containing membrane-spanning segments considerably exceed the above range. For the purpose of this review, we arbitrarily restrict ourselves to peptides containing no more than 50 residues. Hydrophobic mismatch designates a situation in which the hydrophobic regions of a TM ␣-helix and a lipid bilayer differ in length. The energetic penalty of exposing hydrophobic residues to water or hydrophilic ones to the bilayer core can be alleviated by adaptations of the peptide, the bilayer, or both. Positive hydrophobic mismatch results when the hydrophobic sequence of the TM peptide is longer than the hydrocarbon core of the lipid bilayer. Peptides may adapt to this situation by tilting, kinking, reorienting side chains, or partitioning into thicker lipid domains (all reviewed in detail by Killian, 2003; Nyholm et al., 2007; Marsh, 2008). Recent results on model peptides (Özdirekcan et al., 2007) suggest that ear-

lier publications underestimated the extent of tilting. Thus, tilting might play an even more important role than anticipated initially. Furthermore, lipid acyl chains adjacent to the peptide may stretch and stiffen to increase local bilayer thickness. However, this phenomenon is more common for multispanning membrane proteins (Mitra et al., 2004; Andersen and Koeppe, 2007) than for isolated TM helices. Under conditions of negative mismatch, that is, when the hydrophobic peptide sequence is shorter than the bilayer core, lipids often adapt to match the peptide hydrophobic length either by local bilayer thinning due to acyl chain disordering, formation of local nonlamellar structures where the lipid headgroups distort and bend toward the peptide, or lateral lipid sorting leading to enrichment of shorter-chain lipids around the peptide (reviewed in detail by Killian and Nyholm, 2006). Importantly, both positive and negative mismatch may have a strong influence on the strength and specificity of helix–helix interactions (Jensen and Mouritsen, 2004). 3.2. Flanking residues Just like their parent proteins, most TM peptides are very hydrophobic and tend to aggregate or precipitate in some environments, particularly in aqueous solutions (see Section 4.2). In addition to a hydrophobic TM sequence, such peptides therefore often contain flanking hydrophilic residues that do not penetrate into the membrane core but serve to improve peptide solubility. 3.2.1. Types and purposes of flanking residues For the sake of a simple nomenclature in the remainder of this review, we will distinguish between two kinds of flanking sequences: loops and tags. Loops are made up of those amino acid residues that flank a TM sequence in its native context, that is, in the full-length protein the peptide has been derived from. Tags, by contrast, consist of artificial sequences that are not found in the parent protein. Both loops and tags are usually rich in polar or charged amino acid residues like glycine, histidine, proline, lysine, aspartate, and glutamate, but tags may additionally contain nonproteinogenic amino acids such as sarcosine (Melnyk et al., 2003). Besides enhancing solubility and thus facilitating peptide handling, loops and tags can help ensure proper TM topology (e.g., Tomich et al., 1998; Melnyk et al., 2001) or serve as attachment points for a wide range of labels or markers. For example, fluorescence (e.g., Adair and Engelman, 1994; Therien and Deber, 2002; Thévenin and Lazarova, 2008) and spin labels (Karim et al., 2000) can be attached to peptides to expand the experimental repertoire. Table 1 provides an example of a series of peptides that have been derived from the same protein but differ in the length and composition of the flanking sequences attached to the TM domain. 3.2.2. Effects of flanking residues One should always keep in mind that flanking residues may affect not only peptide solubility but also many other biophysical properties, including detergent or lipid affinity, membrane insertion, helical tilt, and so on. A number of commonly used fluorophores have been found to induce higher-order oligomerization

Table 1 Series of TM peptides derived from TM helix 6 of Saccharomyces cerevisiae pheromone ␣-factor receptor Ste2p. Peptide Flank-free Truncated +Loops +Loops + lysine tags +Loops + tags

N-terminal flank

Hydrophobic core sequence

QFDSFH KKKSFH KKKFDSFH

ILLIMSCQSLLVPSIIFILAYSLK AQSLLVPSIIFILAYSLK ILLIMSSQSLLVPSIIFILAYSLK ILLIMSSQSLLVPSIIFILAYSLK ILLIMSAQSLLVPSIIFILAYSLK

C-terminal flank

Naa

Reference

PNQ KKK KKS

24 18 33 33 35

Reddy et al. (1994) Arshava et al. (1998) Naider et al. (2005) Naider et al. (2005) Xie et al. (2000)

In all but the flank-free variant, the only cysteine residue was replaced by either alanine or serine (highlighted in bold) to avoid oxidation problems such as formation of cystine-bridged peptide dimers. Naa is the total number of amino acid residues.

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of normally dimeric leucine zippers in aqueous solution (Daugherty and Gellman, 1999), and there is no reason to rule out such effects a priori for TM peptides. Highly charged tags require special caution, as they may interfere through unspecific electrostatic interactions with other charged components like buffer or detergent or lipid headgroups. Lysine tags, for example, exhibit multivalent electrostatic interactions with phosphate buffer, which may lead to aggregation (Melnyk et al., 2003). Attachment of several lysine residues in sequence may gravely hamper peptide insertion into sodium dodecyl sulfate (SDS) micelles by electrostatically restricting the peptide to the interfacial region of the micelle (Melnyk et al., 2003). Insertion into a lipid bilayer is expected to be even more susceptible to flanking charges because one peptide terminus needs to cross the hydrophobic bilayer core before the peptide can assume its correct TM topology (see Section 6.2). The effectiveness of a tag in solubilizing a hydrophobic peptide depends not only on the lengths and compositions of both the TM sequence and the tag but also on the location of the latter. A synthetic peptide corresponding to TM helix 2 of ␣1 glycine receptor (GlyR) was found to be water-soluble up to 1.4 mM even in the absence of tags (Tomich et al., 1998). This exceptional aqueous solubility is due to the modest hydrophobicity of the poreforming peptide but can further be enhanced by attachment of charged flanking residues: a tetralysine tag improved solubility to 27.5 mM when attached to the C-terminus but only to 13.4 mM when attached to the N-terminus of the peptide. Furthermore, N- and C-terminal oligolysine flanks were shown to have distinct effects on the structures, oligomerization behaviors, and bilayer interactions (Tomich et al., 1998; Broughman et al., 2002a) as well as peptide channel formation and ion selectivity in Madin–Darby canine kidney (MDCK) epithelial cells (Tomich et al., 1998; Wallace et al., 2000; Broughman et al., 2001, 2002a; Shank et al., 2006). A C-terminal pentalysine tag, for instance, completely abolished channel activity of GlyR TM helix 2 (Tomich et al., 1998). More examples of effects of flanking sequences on the properties of TM peptides have been reported for peptides corresponding to TM sequences of GpA (Melnyk et al., 2003), major coating protein 13 (MCP13; Melnyk et al., 2003), fibroblast growth factor receptor (FGFR3; Iwamoto et al., 2005; Merzlyakov et al., 2006), protein E from Coronaviridae (SCoVE; Parthasarathy et al., 2008), and pheromone ␣-factor receptor from Saccharomyces cerevisiae (Ste2p; Naider et al., 2005; Cano-Sanchez et al., 2006). In summary, solubility-mediating flanking residues are often required to render TM peptides amenable to experiments in vitro or applications in vivo (see Section 7.2). However, the influence of flanks on the structure and behavior of a TM peptide can be dramatic and hard to predict. Therefore, whenever possible, flanking sequences should be chosen carefully in terms of length, composition, and location and checked thoroughly with respect to their influence on the application at hand. 4. Isotropic solvents Isotropic solvents are the simplest and crudest membranemimetic systems. Obvious advantages of isotropic solvents are the high solubilities of many TM peptides and the straightforward applicability of a broad spectrum of experimental methods (listed in Table 3 in Chapter 7). The greatest shortcoming of isotropic solvents as membrane-mimetic environments is the lack of chemical and structural heterogeneity as well as anisotropy characteristic of lipid bilayers (see Section 2.1); though seemingly isotropic solvent mixtures can be surprisingly complex (see Section 4.3). Their increased hydrophobicity as compared with water is, in fact, the only property that qualifies organic isotropic solvents as membrane-mimetic systems. For instance, chloroform has a relative static permittivity of ∼5 (Lide, 2009) and thus is thought to

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resemble the hydrophobic core of a lipid bilayer, whereas trifluoroethanol (TFE) with a relative static permittivity of ∼27 is more reminiscent of the lipid headgroup region (Buck, 1998; Hong et al., 1999). Other organic solvents frequently utilized for studying TM peptides include dimethyl sulfoxide (DMSO), methanol, ethanol, 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP), and other hydrogenated or halogenated alcohols as well as mixtures of the above. A few TM peptides are also soluble in water and aqueous solutions, which is of great value for the quantitative description of spontaneous membrane insertion (Hunt et al., 1997b; Wimley and White, 2000; Reshetnyak et al., 2008; see also Sections 4.2 and 6.2) or hydrophobic partition equilibria (see Section 4.3). This chapter summarizes some insights into peptide structure and oligomerization gained from hydrophobic isotropic solvents (Section 4.1), attempts to render TM peptides soluble in aqueous solutions (Section 4.2), and hydrophobicity scales derived from partitioning experiments between aqueous and organic isotropic solvents (Section 4.3). Although these scales were derived using artificially designed peptides, they are covered in this review because they are equally applicable to natural TM sequences. 4.1. Structure and oligomerization of TM peptides in organic solvents In spite of their extreme simplicity and poor membrane resemblance, organic solvents have been applied in peptide research for over 50 years with great success (Goodman and Rosen, 1964). They allow for structure elucidation of hydrophobic TM sequences in the absence of hydrophilic flanks (Tomich et al., 1998; Naider et al., 2003). In at least one case (Arshava et al., 1998), helix formation by an isolated TM segment could hence be shown to be independent of flanking hydrophilic residues (however, see Sections 3.2 and 5.2 for examples where this is not the case). More generally, organic solvents have proven useful in exploring the structural influence of tags (Tomich et al., 1998; Cano-Sanchez et al., 2006), comparing relative helical propensities of different peptides derived from the same membrane protein (Wigley et al., 1998; Lazarova et al., 2004; Bennett et al., 2006), and assessing the impact of disease-related mutations on TM peptide structure (Xie et al., 2000). In the latter case, the authors found that a TM peptide derived from a constitutively active mutant receptor displays a higher helical content than the wild-type peptide. 4.1.1. Secondary structure induction Besides being good media for dissolving hydrophobic peptides, most organic solvents stabilize or induce secondary structure. The exact mechanism underlying this observation was hotly debated for a long time (for a review, see Buck, 1998), but recent reports have shed much light onto this issue (e.g., Roccatano et al., 2002; Duarte et al., 2008a). The most pressing concern with regard to TM domains was whether the dominance of ␣-helical structures observed in various organic solvents (Galardy and Kortylewicz, 1985; Langosch et al., 1991; Reddy et al., 1993; Katragadda et al., 2001a) reflects intrinsic structural propensities of the peptides or merely sequence-unspecific artifacts. Although generic solvent effects can never be ruled out a priori, some circumstantial evidence supports the view that ␣-helical structures of TM domains in organic solvents indeed do, at least to some extent, result from intrinsic structural preferences of the studied peptides (Buck, 1998; Katragadda et al., 2001a,b; Roccatano et al., 2002). MD simulations suggest, for instance, that isotropic organic solvents such as DMSO (Duarte et al., 2008a) and TFE (Roccatano et al., 2002) induce peptide secondary structure by clustering preferentially around and thus limiting water access to the peptide backbone. Simulations of ␣-helical, ␤-sheet, and ␤-hairpin peptides in 30% TFE all revealed high local concentrations of up to

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80% TFE around the peptides (Roccatano et al., 2002). The reduced permittivity and the decreased concentration of water as hydrogen bond donor and acceptor in the vicinity of the backbone favor the formation of intramolecular hydrogen bonds in all three cases. Consequently, formation of any type of secondary structure, not only ␣-helices, will be promoted by TFE as long as requirements for intramolecular hydrogen bonding are satisfied. 4.1.2. Structures of bacteriorhodopsin fragments in DMSO Further support for the usefulness of isotropic solvents for structural investigations of TM peptides comes from the work of Yeagle and co-workers, who characterized peptides derived from rhodopsin (Yeagle et al., 2000; Katragadda et al., 2001b) and bacteriorhodopsin (Katragadda et al., 2001a). In the case of bacteriorhodopsin, the sequence of the full-length protein was split into 12 overlapping fragments, each of which encompassing either a TM domain or a loop sequence. These peptide fragments were synthesized and dissolved in DMSO to determine their structures by solution-state nuclear magnetic resonance (NMR) spectroscopy (with the exception of TM helix 7, which was insoluble in DMSO and some other common organic solvents). The isolated, DMSOsolubilized peptides adopted structures strikingly reminiscent of those observed for the same sequences in two high-resolution crystal structures of the full-length protein (Grigorieff et al., 1996; Pebay-Peyroula et al., 1997), as reflected by an average backbone root mean square deviation (RMSD) < 2.5 Å. The authors (Katragadda et al., 2001a) went on to propose a three-dimensional structure of bacteriorhodopsin by annealing the fragment structures in the overlapping regions and by including additional helix–helix distance restraints taken from a 7-Å crystal structure (Henderson et al., 1990). The structural model thus created had a backbone RMSD of 2.9 Å with respect to the high-resolution crystal structures (Grigorieff et al., 1996; Pebay-Peyroula et al., 1997). Such an approach cannot, of course, substitute for structure determination of full-length membrane proteins as, for example, helix–helix interactions are completely missing. However, the results imply that peptide fragments of ␣-helical membrane proteins possess remarkably stable secondary structure propensities. It is most noteworthy that TM segments and loop sequences may adopt their respective secondary structures even when they are removed from the context of the full-length protein and the anisotropic environment of a lipid bilayer. This speaks in favor of a “divide and conquer” approach to membrane proteins in general and a deliberate use of crude, isotropic solvents in particular. 4.1.3. Influence on helix–helix interactions in solute carrier family 11 Contrary to the enhancing effect they exert on secondary structure, isotropic solvents like TFE (Roccatano et al., 2002) or DMSO (Duarte et al., 2008a) usually disrupt tertiary and quaternary contacts at high solvent concentrations because their solvent shells impair such interactions. Thus, while organic solvents are frequently used in structural studies on single, monomeric peptides, they are widely considered unsuitable for investigations of helix–helix interactions and peptide oligomerization. Using diffusion-ordered spectroscopy (DOSY) NMR, however, Sun and coworkers discovered peptide oligomers even in 100% TFE (Li et al., 2003a, 2004; Xue et al., 2008, 2009) or 40% HFIP (Li et al., 2005b,c, 2008; Xue et al., 2006). The authors determined high-resolution structures of oligomers composed of peptides corresponding to TM helix 4 of solute carrier family 11 member 2 (Slc11a2, formerly known as divalent metal transporter 1 (DMT1)) and TM helix 4 of the homologous protein Slc11a1 (formerly known as natural resistance-associated macrophage protein 1 (Nramp1)). As depicted in Fig. 4, circular dichroism (CD) spectra of TM helix 4

Fig. 4. Concentration dependence of CD spectra of TM helix 4 of Slc11a2 in 100% TFE. The peptide sequence is RVPLYGGVLITIADTFVFLFLDKY. (A) Normalized CD spectra recorded at different peptide concentrations as indicated in the figure. With increasing concentration, signatures of ␣-helical secondary (minima at ∼208 nm and ∼222 nm and maximum at ∼193 nm) become more pronounced, indicating peptide oligomerization.  is the mean residual molar ellipticity and  the wavelength. (B) Plot of the mean residual molar ellipticity at 222 nm,  222 nm , against peptide concentration, c. Figure courtesy of Hongzhe Sun, adapted from Li et al. (2004), with permission of Wiley.

of Slc11a2 in pure TFE clearly depend on peptide concentration, revealing a gain in helicity with increasing concentration. This is indicative of peptide oligomerization accompanied by an increase in ␣-helical structure. The peptide oligomer in 100% TFE consists of four or five monomeric subunits, as determined by DOSY NMR experiments, and has been suggested to exist also in SDS micelles (see Section 5.1). In summary, information gathered from organic isotropic solvents should be interpreted with caution when extrapolating to more complex systems. Nevertheless, because of their ease of use and the broad range of permittivity values covered, isotropic solvents frequently offer a good starting point for further investigations of TM peptides. 4.2. Solubility of TM peptides in aqueous solutions It is not surprising that the vast majority of peptides corresponding to membrane-spanning regions of ␣-helical membrane proteins are too hydrophobic to be reasonably soluble in aqueous solutions. This hurdle can sometimes be overcome by attaching hydrophilic loops or tags to the hydrophobic core sequence (see Section 3.2). Flanking residues add extra polarity or charge to the peptide, thus increasing its aqueous solubility and hampering hydrophobic aggregation or precipitation by electrostatic repulsion. Since, in contrast to more hydrophobic solvents, water can efficiently compete with intrahelical hydrogen bond donors and acceptors, TM peptides are usually largely unfolded and behave like random coils

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when dissolved as monomers in aqueous solutions (e.g., Hunt et al., 1997a,b; Broughman et al., 2002a,b; Lazarova et al., 2004, see also Table S1). 4.2.1. Difficulties due to ˇ-sheet aggregation Oftentimes, TM peptides form extremely stable ␤-sheet aggregates in aqueous solutions. These aggregates may even resist harsh detergents like SDS and therefore are of limited use for further investigations. This was the case for two long TM peptides derived from two receptor tyrosine kinases (RTKs) called Neu and epidermal growth factor receptor (ErbB2) (Jones et al., 2000). At first, both TM domains were produced in good yields by expression as fusion proteins. However, the cleaved and purified TM peptides exhibited CD spectra typical of ␤-structured aggregates that persisted even in SDS–polyacrylamide gel electrophoresis (SDSPAGE). The trick to circumvent this dead end was the choice of an adequate reconstitution procedure. Only after dissolving the lyophilized peptides in a chloroform/formic acid/acetic acid/TFE (2:1:1:1) mixture and evaporating the organic solvents could the TM peptides insert into SDS micelles or lipid bilayers and thereby assume predominantly ␣-helical structures, as evidenced by NMR and CD spectroscopy. Moreover, both TM peptides finally revealed monomer–dimer equilibria in SDS-PAGE, indicating that the isolated TM sequences can induce dimerization in the absence of the water-soluble domains of their respective parent proteins. Furthermore, because of the crucial role of electrostatic forces in keeping TM peptides water-soluble, ionic strength and pH of the buffer solution have to be chosen carefully. The risk of aggregation rises as repulsive electrostatic forces get weakened by increasing ionic strength, as shown by Robinson and co-workers (Lazarova et al., 2004). The authors synthesized seven TM peptides, each corresponding to one of the seven TM helices of the adenosine A2 a receptor (A2 aR). All peptides were soluble and adopted randomcoil conformations in pure water but showed differential behavior in the presence of 10 mM Tris, pH 7. While peptides derived from TM helices 1, 2, 4, and 5 remained soluble and unfolded, those derived from TM helices 3, 6, and 7 aggregated into structures rich in ␤-sheet. In contrast to the example discussed in the preceding paragraph, these peptide aggregates could be disrupted by addition of SDS, in which the TM peptides assumed ␣-helical secondary structures. 4.2.2. pHLIP in aqueous solution Only a few flank-free TM peptides are water-soluble at micromolar or higher concentrations without aggregating or precipitating. A notable example is TM helix 3 of bacteriorhodopsin. In a systematic and extensive “divide and conquer” investigation of a multispanning membrane protein, Engelman and co-workers (Hunt et al., 1997a) synthesized and characterized seven peptides, each corresponding to one of the seven TM domains of the full-length protein. Somewhat surprisingly, they discovered that a peptide derived from TM helix 3 (with glutamine at position 105 in the second cytoplasmic loop replaced by glutamic acid) is soluble in aqueous solutions in the absence of membrane-mimetic systems or denaturants (Hunt et al., 1997b). The peptide later became known as pH low insertion peptide (pHLIP) because of its pH-dependent mode of interaction with lipid bilayers (see Section 6.2). When dissolved in aqueous solution around neutral pH, pHLIP undergoes a concentration-dependent monomer–tetramer equilibrium, as shown by a combination of size-exclusion chromatography (SEC), CD and fluorescence spectroscopy, as well as analytical ultracentrifugation (AUC; Reshetnyak et al., 2007). At a peptide concentration of ∼7.5 ␮M, 94% of the peptide is monomeric and 6% tetrameric, as deduced from sedimentation velocity experiments. The tetramer exhibits CD spectra typical of ␣-helical secondary structure and exciton formation, whereas the

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monomeric form predominant at lower concentrations is largely unstructured and reveals no exciton signatures. Thus, pHLIP obeys a well-defined homooligomerisation equilibrium and does not aggregate unspecifically or irreversibly when dissolved in aqueous solutions. By virtue of its relatively good aqueous solubility, pHLIP is a rare case in which water/bilayer partitioning can be studied experimentally (see Section 6.2). 4.3. Partitioning scales based on isotropic solvents Isotropic solvents have played an indispensable role in the development of hydrophobic partitioning scales, which quantify a peptide’s or protein’s affinity to insert into a lipid bilayer on a per-residue basis. Such scales are widely applied to identify membrane-spanning regions with the aid of sequence information, to design new membrane-anchored peptides, or to quantitatively understand bilayer insertion of peptides and proteins, to name but a few. Hydrophobic partitioning scales can be obtained in three fundamentally different ways: (1) experimental partitioning data (e.g., Jayasinghe et al., 2001); (2) statistical analysis of known membrane protein structures (e.g., Kitsas et al., 2006); or (3) computational approaches like MD or Monte Carlo simulations (e.g., MacCallum et al., 2007). The first option is particularly convincing because it relies exclusively on experimental data and paves the way for rationalizing bilayer insertion in terms of simple physicochemical principles without any evolutionary bias. 4.3.1. Whole-residue partitioning scale Derivation of experimental hydrophobicity scales necessitates accurate determination of the free energies of partitioning from an aqueous phase into a lipid bilayer or a membrane-mimetic environment for each proteinogenic amino acid. Additionally, hydrophobicity scales must also account for the energetically costly dehydration of the peptide backbone upon bilayer partitioning (Wimley et al., 1996; Wimley and White, 1996; White et al., 2001; see also Section 2.2). To quantify side chain partitioning, White and co-workers (Wimley et al., 1996) designed a series of pentapeptides of sequence AcWL-X-LL, with Ac denoting acetylation of the N-terminus and X any of the naturally occurring amino acids. The free energy cost of peptide backbone dehydration was obtained with the aid of model peptides of various hydrophobic lengths of sequence AcWL1–6 . However, as all of the peptides were way too short to insert into a lipid bilayer in a TM topology, octanol was used to mimic the hydrophobic membrane core. On the basis of the partition coefficients from water into water-saturated octanol, the free energies of transfer could be calculated for each side chain and the peptide backbone. The octanol partitioning scale was compiled into a prediction tool called Membrane Protein Explorer (MPEx; Jayasinghe et al., 2001), which was later extended to account for salt bridges, helical hydrogen-bonded peptide backbones, and capped or charged peptide termini (Hristova and White, 2005). A further refinement was achieved by combining the octanol scale with a second scale relating to peptide partitioning into the interfacial region of a lipid bilayer (Jaysinghe et al., 2008; see http://blanco.biomol.uci.edu/mpex). This improved scale has proven remarkably successful in quantitatively predicting the bilayer insertion energetics of peptides and proteins; detailed descriptions and discussions can be found in the literature (Wimley et al., 1996; Wimley and White, 1996; White and Wimley, 1999; White et al., 2001; Jayasinghe et al., 2001; Ladokhin and White, 2001; Fernández-Vidal et al., 2007; White, 2007). 4.3.2. Suitability of octanol for determining partitioning scales From the perspective adopted in the present review, the most striking question emerging from the preceding paragraphs is how

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partitioning experiments relying on a supposedly homogenous and isotropic solvent like octanol can be so astonishingly reliable in mimicking the interactions between TM peptides and the hydrophobic core of a lipid bilayer. Part of the answer is that the refined hydrophobicity scale explicitly accounts for the heterogeneity and anisotropy of a lipid bilayer by resorting to two different scales, one for the interfacial region and another one for the hydrophobic core. Only the latter is based on octanol partitioning, and the hydrophobic bilayer core as such is much less heterogeneous in terms of chemical composition, charge density, and permittivity than the bilayer as a whole (see also Fig. 1). On top of this, however, part of the explanation could lie in the finding that water-saturated octanol exhibits water inclusions reminiscent of inverse micelles (Franks et al., 1993; Jayasinghe et al., 2001), which might rearrange to differentially adapt to polar and apolar parts of a peptide. In conclusion, water-saturated octanol might be less homogenous and isotropic, and thus could represent a better membrane-mimetic solvent, than one might think at first glance.

5. Detergent micelles Detergents are a chemically diverse class of amphiphilic compounds that are of great interest in the study of membraneinteracting proteins and peptides. Like bilayer-forming lipids, detergents consist of a hydrophilic headgroup and a hydrophobic tail driving self-association in aqueous environments. In contrast to bilayer-forming lipids, and with some notable exceptions such as bile salts and related surfactants, detergents are characterized by headgroups that are considerably bulkier than their hydrophobic tails. This molecular shape impedes self-association into planar structures like bilayers and, instead, promotes formation of micelles (Fig. 5) when the detergent concentration exceeds the critical micellar concentration (CMC). CMC values vary tremendously among detergents, ranging from virtually undetectably low concentrations for very hydrophobic detergents to the high millimolar range for hydrophilic ones. CMC values depend on physical parameters such as temperature and, particularly for ionic detergents, on ionic strength and pH. Furthermore, the presence of TM peptides or other micelle-incorporated compounds may drastically lower the apparent CMC. The cross-sectional topology of a micelle mimics that of a lipid bilayer in as far as its hydrophobic hydrocarbon core is shielded from water by an interface formed by detergent headgroups. From this perspective, detergent micelles are expected to be much better membrane-mimetic systems than isotropic solvents discussed in the previous chapter. However, there remain a number of principal differences between detergent micelles and lipid bilayers. Most importantly, both the headgroup region and the hydrocarbon core in a highly curved micelle are packed less orderly (see Fig. 7) and exhibit greater dynamics than in a planar or near-planar lipid bilayer. Thus, the shielding effect of the interfacial headgroup region is less pronounced, and water molecules can penetrate more easily into the micellar core. Furthermore, the good aqueous solubilities of many detergents (as reflected by high CMCs) result in continuous and rapid exchange of detergent monomers between the aqueous and micellar phases as well as straightforward incorporation of other solutes such as peptides into micelles. In brief, detergent micelles offer a valuable compromise for investigating TM peptides in membrane-mimetic systems, combining ease of use and good dissolving properties with an anisotropic environment (see Table 3 in Chapter 7). What should always be kept in mind, however, is that many detergents exert a denaturing effect on membrane proteins and peptides by abrogating helix–helix interactions (e.g., Melnyk et al., 2001; Therien and Deber, 2002). In this chapter, we discuss the influence of detergent micelles on

Fig. 5. Schematic representations of lipid membranes and membrane-mimetic systems: (A) spherical micelle; (B) small bicelle; (C) bilayer fragment; (D) small unilamellar vesicle (SUV); (E) multibilayers on a solid support; and (F) planar lipid membrane. Illustrations are purely schematic and not drawn to scale. This is especially true for the spanned bilayer in panel F: in a real setup, the hole spanned by the bilayer is typically ∼100 ␮m in diameter and thus contains ∼1010 lipid molecules per leaflet (assuming a circular cavity and an average area requirement of 70 Å2 per lipid molecule). Moreover, the septum is at least 1000 times thicker than the bilayer. The annulus of the planar lipid membrane contains alkanes or other non-surface-active hydrocarbons providing stabilization. Figure created using Art of Illusion (Peter Eastman, v2.7).

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Table 2 Sequences of ␣1 GlyR TM helix 2 used in different membrane-mimetic systems. Model systems SDS, DPC, DMPC SDS 30% TFE, SDS, DPC, DMPC eggPC POPC/POPE 30% TFE 40% TFE

Peptide sequence NH2 NH2 NH2 NH2 NH2 NH2 NH2 NH2 -

PARVGLGITTVLTMTTQSSGSRA APARVGLGITTVLTMTTQSSGSRASLPK KKKKPARVGLGITTVLTMTTQS ARVGLGITTVLTMTTQSSGSRA PARVGLGITTVLTMTTQSSGSRA KKKKPARVGLGITTVLTMTTQSSGSRA PARVGLGITTVLTMTTQSSGSRAKKKK KKKKPARVGLGITTVLTMTTQS

-COOH -COOH -COOH -COOH -COOH -COOH -COOH -CONH

Methods

Reference

NMR, MD NMR MD BLM Conductane recordings NMR, MD

Tang et al. (2002) Yushmanov et al. (2003a) Johnston et al. (2006) Langosch et al. (1991) Reddy et al. (1993) Broughman et al. (2002a)

NMR

Cook et al. (2004)

eggPC, phosphatidylcholine extracted from hen egg; BLM, black lipid membrane; POPE, 1-palmitoyl-2-oleoyl-sn-glycero-ethanolamine.

both the structure (Section 5.1) and the oligomerization behavior (Section 5.2) of TM peptides. 5.1. Structural impact of detergent micelles Detergent micelles can induce and stabilize ␣-helical secondary structure by embedding apolar amino acid residues in their hydrophobic core, whose low permittivity promotes intrahelical hydrogen bonding. At the same time, hydrophilic flanking residues interact with detergent headgroups and water in the interfacial region and may help stabilize a transmicellar orientation of a TM peptide. Detergent micelles are popular model systems for structural studies of TM peptides using NMR or CD spectroscopy because, owing to their small sizes as compared with lipid vesicles, they exhibit fast tumbling and reduced light scattering, respectively. Likewise, micelles with relatively low aggregation numbers are particularly easily amenable to MD simulations as they can be handled with reasonable computational effort. 5.1.1. TM helix 2 of ˛1 glycine receptor: influence of micellar environment Among the numerous hydrophobic peptides explored in micellar systems, we picked TM helix 2 of GlyR as an illustrative and debatable example. Several groups have scrutinized different variants of this peptide in all kinds of membrane-mimetic systems using various techniques (Table 2) and have arrived at quite contradictory conclusions.

First of all, it is instructive to compare structural models of GlyR TM helix 2 in SDS micelles obtained by experimental and computational means (Fig. 6). Using NMR spectroscopy, Xu and co-workers (Yushmanov et al., 2003a) found an ␣-helix containing four helical turns flanked by a moderately flexible N-terminus and a highly flexible C-terminus. By contrast, MD simulations of Sansom and co-workers (Johnston et al., 2006) suggested a much shorter helix consisting of only two turns sandwiched between an extremely flexible N-terminus and an almost rigid C-terminus. Moreover, while NMR data were consistent with a transmicellar topology, MD simulations implied a drastic deviation of micellar shape from spherical geometry upon peptide incorporation and a more peripheral location of the peptide helix in SDS micelles. The peptide sequence employed in the MD simulations lacked nine C-terminal residues present in the experimental study but contained an additional tetralysine tag at its N-terminus (Table 2). However, it is questionable if these discrepancies result solely from differences in the hydrophilic flanking sequences. In particular, it is counterintuitive, though certainly not impossible, that attachment of a tag destabilizes the N-terminus, whereas removal of a loop stabilizes the C-terminus, as inferred from MD simulations. A more likely explanation is that, in spite of the relatively small size and simplicity of SDS micelles, some essential features of this membrane-mimetic system are not fully captured in the MD simulations. It is interesting to note in this context that when similar calculations were performed in DPC rather than SDS micelles (Johnston et al., 2006), the peptide was observed

Fig. 6. Structures of GlyR TM helix 2 in different membrane-mimetic systems: (A), (B), (D), and (E) Superpositions of 10 snapshots taken every 2 ns during a 20-ns MD simulation in (A) 30% TFE; (B) SDS; (D) DPC; and (E) DMPC. Figures courtesy of Jennifer Johnston, adapted from Johnston et al. (2006), with permission of Elsevier. (C) Superposition of 10 lowest-energy NMR structures in SDS. Figure created from PDB file 1mot (Yushmanov et al., 2003a). The central TM sequence delimited by residues depicted in black is identical in all peptides (see Table 2 for peptide sequences).

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to assume a transmicellar orientation and leave the micellar shape almost unaffected. Furthermore, the authors found considerably more headgroup–peptide interactions in DPC than in SDS micelles. 5.1.2. TM helix 2 of ˛1 glycine receptor: comparison with other membrane-mimetic systems In addition to MD simulations in different micellar environments, Sansom and co-workers (Johnston et al., 2006) also calculated the structures of GlyR TM helix 2 in 30% TFE and bilayers made up of the zwitterionic phospholipid 1,2-dimyristoyl-snglycero-phosphocholine (DMPC), thus allowing for a comparison of TM peptide structures in isotropic solvents, detergent micelles, and lipid bilayers (Fig. 6). The helical content of the TM peptide was suggested to be highest in DMPC bilayers, followed by DPC, SDS, and finally 30% TFE. Notably, interactions between polar peptide residues and detergent or lipid headgroups appeared similar in micellar DPC and bilayer-forming DMPC, which both contain a phosphocholine moiety in their headgroups. 5.1.3. TM helix 2 of ˛1 glycine receptor: pentamer model Finally, Xu and co-workers (Tang et al., 2002) derived a model of GlyR structure and function from their NMR experiments on TM helix 2. The peptide structure determined in SDS micelles was used as starting point in subsequent MD simulations performed in DMPC bilayers. According to this model, the TM peptide self-assembles to form a pentameric channel that fluctuates between open and closed conformations by a concerted 10◦ tilt of all five helices. The authors conclude that, while anion selectivity appears to be conferred by TM helix 2 itself, channel gating is likely to be controlled by other domains of full-length GlyR. Ion conductance studies performed on pentamers of GlyR TM helix 2 (Langosch et al., 1991; Reddy et al., 1993; see Table 2) are discussed in greater depth in a later chapter (see Section 6.3), whereas more examples of TM peptides oligomerizing in detergent micelles are detailed in the next section. 5.2. Oligomerization of TM peptides in detergent micelles Helix–helix interactions in micelles are susceptible to the physical and chemical properties of the detergent in which they take place. They can be strong and specific in a “mild” detergent and completely absent or rather unspecific in a “harsh” one. Unfortunately, the choice of a suitable detergent for a TM peptide or membrane protein is normally based on cumbersome trialand-error approaches. SDS, the most common representative of denaturing detergents, has been suggested to directly bind to helix–helix interfaces by van der Waals forces, thereby inhibiting interhelical interactions (Renthal, 2006). However, if due care is given to the choice of detergent, micelles may provide a rewarding membrane-mimetic system for studying peptide oligomerization (e.g., Lemmon et al., 1992; Wigley et al., 1998; Melnyk et al., 2001). For example, detergent micelles were used to demonstrate that TM peptide oligomerization may occur in the absence of other protein domains (Bormann et al., 1989), and the common dimerization motif GXXXG (reviewed in Moore et al., 2008) was first observed in detergent micelles (Lemmon et al., 1994). Point mutations in a peptide derived from the tetrameric M2 proton channel of influenza A virus dissolved in DPC micelles led to tetramer stabilization and concomitant loss of proton channel function, suggesting that the wild-type M2 peptide sacrifices structural stability for channel function (Stouffer et al., 2005). 5.2.1. Influence of detergents on glycophorin A dimerization A comprehensive account of the influence of detergents on peptide–peptide interactions was obtained from fluorescence res-

onance energy transfer (FRET) experiments performed on the TM domain of GpA in the presence of various concentrations of seven different detergents (Fisher et al., 1999; Fisher et al., 2003). The authors found that, while the TM domain of GpA is largely ␣-helical in micelles irrespective of the detergent used, peptide dimerization strongly depends on the choice of detergent. However, no obvious correlations with headgroup type or hydrophobic chain length became apparent. Likewise, simple models in which peptide dilution within the hydrophobic phase (i.e., micelles) would drive dimer dissociation with increasing detergent concentration were found incapable of explaining the experimental data. Curiously, raising the concentration of the harsh detergent SDS seems to have a smaller effect than predicted by ideal dilution, whereas some supposedly milder detergents exert a greater influence. For SDS, van’t Hoff analysis of measurements taken at different temperatures suggests large enthalpic and entropic contributions to the free energy of dimerization. As the SDS concentration is raised, enthalpy increasingly favors dimerization, whereas the entropic term changes from favorable to unfavorable. The dependencies of the enthalpic and entropic contributions on SDS concentration cancel each other to a considerable extent, so that the free energy of dimerization is less susceptible to detergent concentration. The detailed molecular origins of the complex interplay between oligomerizing peptides and self-associating surfactants remain to be elucidated. A deeper understanding of such interactions might represent a major advance in the search for detergents that do not abrogate membrane protein stability and activity. 5.2.2. Oligomerization of TM helix 4 of solute carrier family 11 member 2 in SDS A less clear-cut case of peptide oligomerization in micelles was reported by Sun and co-workers (Li et al., 2003a, 2004), who extended their studies of TM domain 4 of Slc11a2 (see Section 4.1) to SDS micelles, where the peptide assumes ␣-helical secondary structure. NMR line broadening experiments revealed that every third to fourth residue in the helix is accessible to the paramagnetic probe Mn2+ , indicating that one side of the helix and part of the termini are exposed to water. As the peptide oligomerizes in a concentration-dependent fashion in 100% TFE or 40% HFIP (see Section 4.1), the authors rationalized the Mn2+ accessibility pattern in SDS micelles by postulating the existence of an oligomeric peptide pore permeable to divalent metal ions. Changes in ␣ H and N H chemical shifts with increasing peptide/detergent ratio were taken as a further indicator of homooligomerization. The lack of interhelical nuclear Overhauser effects (NOEs) in the peptide pore was attributed to a high degree of symmetry in the oligomeric structure. Although the above conclusions appear plausible and consistent with the biological function of Slc11a2, alternative interpretations should also be considered. The periodicity of Mn2+ accessibility and the lack of interhelical NOEs can just as well be explained by a peripheral or interfacial orientation of the amphipathic peptide helix with partly inward-bent termini, as found for GlyR TM helix 2 in SDS micelles (Johnston et al., 2006; see Section 5.1). The rather modest dependence of the chemical shifts on the concentrations of peptide and detergent could as well be attributed to other factors that depend on the peptide/detergent ratio and cause minor structural rearrangements of the peptide, such as changes in micellar shape and aggregation number. This view is also supported by the absence of Slc11a2 TM helix 4 oligomers in SDS-PAGE (Li et al., 2003a). That SDS-PAGE is, in principle, capable of detecting TM peptide oligomerization has been demonstrated for GpA (Fisher et al., 1999), ErbB1 (Jones et al., 2000), MCP13 (Melnyk et al., 2003), and many other proteins (e.g., Melnyk et al., 2001;

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Tang et al., 2002; Rath et al., 2006; Afara et al., 2006; see also Table S1). Even if more convincing evidence of peptide oligomers in SDS micelles can be found, it still remains to be shown that they possess pore activity. To settle the uncertainties surrounding Slc11a2 TM helix 4, and a similar case involving TM helix 4 of Slc11a1 (Xue et al., 2006, 2008, 2009), further experiments are urgently needed. 6. Lipid bilayers Lipid bilayers consist of two apposing leaflets of lipids and constitute an indispensable part of all biological membranes. The latter, however, additionally contain a multitude of different proteins and glycoproteins and form numerous connections with other cellular components such as the cytoskeleton. Pure lipid bilayers lack these features, but they can nevertheless attain a considerable degree of complexity in terms of lipid composition and leaflet asymmetry. While the simplest bilayers consist of a single, synthetic lipid species, more complex ones can be built from well-defined lipid mixtures or less well-defined lipid extracts obtained from natural sources. By varying the lipid composition in a systematic fashion, lipid-specific effects can be distinguished from the influence of generic physical parameters such as headgroup charge, hydrophobic thickness, or lateral pressure profile. Lipid bilayers offer an environment to proteins and peptides that more closely resembles natural membranes than do isotropic solvents or detergent micelles (see Table 3 in Chapter 7). Lipids in a bilayer are more orderly packed and less dynamic than detergents in a micelle (Fig. 7). On the other side of the coin, this means that reconstitution of proteins and peptides into lipid bilayers is oftentimes a more tedious procedure than their solubilization in isotropic solvents or detergent micelles. Using membrane curvature as a criterion, lipid bilayer systems can be divided into two broad classes: vesicles (also known as liposomes) and planar bilayers (Fig. 7). This chapter reviews two studies on peptide–vesicle interactions that are of outstanding interest from a structural (Section 6.1) or thermodynamic (Section 6.2) viewpoint and gives a brief overview of information about TM peptides gained using various kinds of planar lipid bilayers (Section 6.3). 6.1. TM peptide dimers in lipid vesicles Lipid vesicles are very popular and versatile model systems for the study of TM peptides. Unilamellar vesicles consist of a single lipid bilayer, whereas multilamellar vesicles are made up of an onion-like shell comprising several stacked bilayers. Vesicles range in size from ∼20 nm to several micrometers. Unilamellar vesicles are categorized according to their size as small (<50 nm), large (50–500 nm), or giant (>500 nm) unilamellar vesicles (SUVs, LUVs, and GUVs, respectively; Torchilin and Weissig, 2003). Bilayer curvature is small in LUVs and negligible in GUVs; in SUVs, by contrast, the curvature is so pronounced that the outer leaflet contains considerably more lipid than the inner leaflet. In a liposome having a diameter of 30 nm, for instance, ∼60% of the total lipid resides in the outer leaflet (Seelig, 1997). A second consequence of strong bilayer curvature is that lipid headgroups in the outer leaflet of an SUV are packed more loosely than they would be in a planar bilayer, whereas headgroups in the inner leaflet are packed more tightly (see Fig. 7). Several laboratories have successfully applied lipid vesicles for quantifying helix–helix interactions between TM peptides (e.g., Gerber et al., 2004b; Stouffer et al., 2008b) or for determining highresolution structures of TM peptides by NMR (e.g., Smith et al.,

Fig. 7. Snapshots of MD simulations of (A) a DPC micelle; (B) a DPPC SUV; and (C) a planar DPPC bilayer. The SUV structure was generated by coarse-grained simulation, whereas the micelle and planar bilayer structures were obtained from full-atom simulations. Water and hydrogen atoms are not shown for clarity (see Fig. 2 for color code). (A) Full-atom simulation of a DPC micelle. Figure adapted from Tieleman et al., 2000 (PDB file downloaded from http://moose.bio.ucalgary.ca , with permission of Peter Tieleman). (B) Coarse-grained simulation of a DPPC SUV with a diameter of ∼20 nm. Lipids are represented by 12 coarse-grained superatoms: N+ (CH3 )3 in white, PO4 − in red, Glyc1 and Glyc2 in slate grey, and C1, C2, C3, and C4 (for each fatty acyl tail) in black. Figure courtesy of Siewert-Jan Marrink, adapted from Marrink and Mark, 2003, with permission of American Chemical Society. (C) Full-atom simulation of 128 DPPC molecules in a planar bilayer. Figure adapted from Tieleman and Berendsen, 1996 (PDB file downloaded from http://moose.bio.ucalgary.ca , with permission of Peter Tieleman).

2001; Valentine et al., 2001). Here, we limit ourselves to the discussion of a single case that combines both of the above aspects and, furthermore, enables comparison with structural data collected in a micellar environment. 6.1.1. Helix–helix interactions in the glycophorin A dimer Aimoto and co-workers (Smith et al., 2001) compared the dimer structure of the GpA TM domain they obtained in lipid vesicles by solid-state NMR with the corresponding structure in detergent micelles determined by Engelman and co-workers (MacKenzie et al., 1997) using solution-state NMR. In DPC micelles, the dimer interface reveals a tight knobs-into-holes packing with a helix crossing angle of ∼40◦ . Interhelical van der Waals interactions are seen only between residues 1 I76 ↔ 2 L75, 1 V80 ↔ 2 G79/2 G83, and 1 V84 ↔ 2 G83/2 T87, where, for example, 2 G79 denotes a glycine residue at position 79 of helix 2 and residue numbers refer to fulllength GpA (nomenclature according to Smith et al., 2001). This

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contrasts with the dimer structures determined in vesicles composed of DMPC or POPC (Smith et al., 2001). Although the peptide dimer again exhibits a tight knobs-into-holes packing, the helix crossing angle is somewhat lower at ∼35◦ . This change in crossing angle might seem moderate at first glance, but it entails a major shift in the dimerization interface, which now encompasses van der Waals contacts between residues 1 I76 ↔ 2 G79, 1 V80 ↔ 2 G79/2 G83, 1 G79 ↔ 2 G79, 1 V84 ↔ 2 G83, 1 G83 ↔ 2 G83, 1 T87 ↔ 2 I88, and 1 I88 ↔ 2 T87. Although it is conceivable that the differences in GpA dimer structure between micelles and vesicles are, at least in part, due to an insufficient number of distance constraints in the micellar system (MacKenzie et al., 1997), three observations deserve explicit mention: First, in lipid bilayers, but not in detergent micelles, glycine–glycine (G79, G83) contacts are obvious, which are abundant in helix–helix interfaces of membrane proteins (Javadpour et al., 1999; Eilers et al., 2000). Second, the bilayer-embedded dimer is stabilized by an interhelical hydrogen bond (1 T87 ↔ 2 I88), which does not show up in the micelle-suspended dimer (Smith et al., 2001, 2002a). Thus, van der Waals interactions obviously suffice to stabilize the dimer, and it would be interesting to know to what extent the additional hydrogen bond further contributes to dimer stability in the membrane. Third, the dimer structures obtained in DMPC and POPC bilayers are indistinguishable in spite of marked differences in hydrophobic bilayer thickness. Thus, although the dimer structures in micelles and bilayers share the same overall fold, they substantially differ in their structural details. By contrast, the choice of lipid constituting the bilayer is less relevant to this dimer structure, at least for the two phospholipids used in the present example. 6.2. Energetics of TM peptide self-insertion into lipid vesicles Elucidating the bilayer insertion energetics of an isolated TM peptide would greatly contribute to a quantitative picture of the first step of the two-stage model of membrane protein folding (see Section 2.2). pHLIP appears extraordinarily suited for this purpose because of its exceptional aqueous solubility (see Section 4.2) and pH-controllable self-insertion (Hunt et al., 1997b): at neutral pH, the peptide adsorbs to the membrane surface as an unstructured monomer. At lower pH, by contrast, pHLIP spontaneously and reversibly self-inserts into the lipid bilayer in an ␣-helical TM orientation. This process has an apparent pKa of 6.0 and is thought to result from protonation of two aspartate residues residing in the TM sequence, which renders the peptide more hydrophobic and thus allows translocation of the C-terminus across the hydrophobic bilayer core. 6.2.1. Stoichiometric binding model The Engelman group (Reshetnyak et al., 2008) titrated pHLIP with POPC LUVs at pH 8 to follow superficial membrane adsorption or at pH 5 to study TM insertion with the aid of fluorescence spectroscopy and isothermal titration calorimetry (ITC). Scatchard plots derived from these titrations revealed pronounced upward curvature and supposedly biphasic behavior. The authors took this as evidence that lipid interactions of pHLIP do not obey a simple membrane binding equilibrium and, consequently, invoked a more complex scenario. They found that the Scatchard plots at both pH values may be fitted reasonably well by assuming two distinct sets of binding sites that differ in their peptide affinities by about an order of magnitude. At 37 ◦ C, the standard Gibbs free energy of superficial adsorption amounts to −30 kJ/mol for the high-affinity site and −25 kJ/mol for the low-affinity site. The corresponding values for TM helix insertion from the aqueous phase are −38 kJ/mol and −30 kJ/mol for high- and low-affinity sites, respectively. Transition from the superficially adsorbed, unstructured conformation

to the ␣-helical TM conformation hence is characterized by a Gibbs free energy change between −5 kJ/mol and −8 kJ/mol, which is in agreement with a value of −7.5 kJ/mol obtained from ITC acid-titration experiments (Reshetnyak et al., 2008). However, these values should be regarded with caution for several reasons. On the one hand, according to the above Scatchard analysis, the numbers of lipid molecules per binding site suggested by ITC are considerably higher than those indicated by fluorescence data. The authors ascribed this discrepancy to the sensitivity of ITC to perturbations of additional lipid molecules, whereas fluorescence spectroscopy reports the number of lipid molecules in the binding site proper (Reshetnyak et al., 2008). While this as such is certainly conceivable, the size of a binding site returned from a Scatchard analysis should, nevertheless, not depend on the method used. The number of lipid molecules per binding site is inversely proportional to the number of lipid-bound peptide molecules when the membrane is saturated, which must be independent of the technique employed to quantify membrane binding. On the other hand, the reported binding constants increase with increasing temperature, whereas negative binding enthalpies observed in ITC runs would, according to the van’t Hoff equation, indicate the opposite. Finally, application of a stoichiometric binding model assumes binding of one peptide molecule to a binding site made up of a well-defined number of lipid molecules, which is a rather unlikely process. 6.2.2. Partitioning model Interactions of a wide range of peptides with lipid membranes have been shown to be more adequately described by a partition equilibrium rather than a stoichiometric binding model (Seelig, 1997, 2004). Accordingly, a solute such as a peptide partitions between two phases, namely, the lipid bilayer phase and the aqueous phase adjacent to the bilayer. The peptide concentration in the buffer in immediate vicinity of the membrane differs from the bulk aqueous peptide concentration because adsorption of charged peptide causes charge accumulation at the membrane surface and repulsion of like charges, including peptide in the aqueous phase. If passed unnoticed, such effects might be misinterpreted as signs of biphasic behavior. Even small solutes such as tryptophan (Gómez et al., 2002) or SDS (Keller et al., 2006) give rise to apparently biphasic membrane binding isotherms. However, on correcting for electrostatic forces, the isotherms turn linear, can be described by a membrane partition equilibrium, and are in accord with the van’t Hoff equation. When this approach is extended to peptide–lipid interactions (McLaughlin, 1977, 1989; Seelig, 1997, 2004), it is important to realize that electrostatic and hydrophobic effects at the membrane surface are usually not additive for polyionic solutes (Ladokhin and White, 2001). This should be accounted for by including an effective peptide valence as an additional fitting parameter rather than using the formal valence (Beschiaschvili and Seelig, 1990). For these reasons, it appears likely that a scenario combining a simple membrane partition equilibrium with surface electrostatics is an alternative to, and maybe a more appropriate choice than, a two-site stoichiometric binding model (Reshetnyak et al., 2008) to describe lipid interactions of pHLIP. This does not discount the possibility that multiple surface-adsorbed and bilayer-embedded conformations exist in parallel, as was also suggested by kinetic experiments (Tang and Gai, 2008). It means, however, that our quantitative understanding of the thermodynamics of pHLIP selfinsertion is currently based on weak ground and awaits further experimentation. Beyond deepening our insights into a fundamental issue of membrane protein folding, progress in this direction is also expected to shed light on the ability of pHLIP to drag cargo molecules attached to its C-terminus across the membrane core, which makes this peptide interesting for the intracellular deliv-

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ery of therapeutic or diagnostic tools (Reshetnyak et al., 2006; see Section 7.2). 6.3. Planar bilayers as hosts for TM peptides Planar lipid bilayers appear as bicelles, supported bilayers or multibilayers, and immersed (or spanned) bilayers, all of which have been employed to study TM peptides. 6.3.1. Bicelles in TM peptide research Bicelles are lipid discs formed by mixing a long-chain lipid (e.g., DMPC) with a short-chain lipid (e.g., 1,2-diheptanoyl-sn-glycerophosphocholine (DHepPC)) at a specific ratio (van Dam et al., 2004). While the long-chain lipid constitutes the central bilayer part of the disc, the short-chain lipid seals its rims (Fig. 5). Unlike vesicles or spanned bilayers, bicelles are not suitable for transbilayer transport studies since they do not separate two unconnected aqueous compartments. However, bicelles are the only bilayer system in which a self-inserting peptide can adopt a TM orientation without the need of dragging one of its termini across the hydrophobic membrane core. On top of that, owing to their small size and, in some cases, their spontaneous alignment in an external magnetic field, bicelles hold some promise for structure determination by solution-state NMR (Poget and Girvin, 2007). Nevertheless, only a few reports on the application of this rather new membrane-mimetic system to TM peptides have been published so far (e.g., De Angelis et al., 2004; Sato et al., 2006; see also Table S1). In the latter report, the secondary structure of a TM peptide derived from ErbB1 was determined by CD spectroscopy in DHepPC/DMPC/DMPG (13:10:3) bicelles. 6.3.2. TM peptide orientation in multibilayers Using attenuated total reflection Fourier transform infrared spectroscopy (ATR-FTIR), the same authors (Sato et al., 2006) confirmed membrane insertion of the EGRF TM peptide using another kind of planar bilayers, that is, supported multibilayers (sometimes also referred to as stacked bilayers). This approach is excellently suited for the determination of peptide topology because it allows for a macroscopic orientation of the bilayer sample with respect to laboratory coordinates. Other methods that serve the same purpose are oriented CD (OCD; Wu et al., 1990; Iwamoto et al., 2005) and, if isotopically labeled peptide is available, solid-state NMR (e.g., Marassi et al., 1999a; Karp et al., 2006; see also Table S1). Multibilayers can be formed by deposition of vesicles containing the peptide of interest or by evaporation of organic solvent from a dispersion containing both lipid and peptide, followed by rehydration of the lipid/peptide film. Low hydration levels can change bilayer thickness and restrict water or lipid diffusion in multibilayers compared with vesicles in bulk aqueous solution (Chen and Hung, 1996; Högberg and Lyubartsev, 2006; Binder, 2007), which, in turn, might influence diffusion, insertion, and folding of the reconstituted peptide (De Planque and Killian, 2003; Karp et al., 2006). The structure of a TM peptide derived from phospholamban (PLB), for example, is controlled not only by the presence of the negatively charged phospholipid 1-palmitoyl-2-oleoyl-sn-glycero-phosphoglycerol (POPG) but also by the hydration level of the host multibilayers, as maximum ␣-helical content is achieved only upon full bilayer hydration (Karp et al., 2006). 6.3.3. TM helix 2 of ˛1 glycine receptor: channel activity in spanned bilayers Immersed or spanned bilayers represent the third version of planar bilayers and are the system of choice for studying transbilayer transport. In this setup, a planar bilayer separates two aqueous compartments so that solute transport between two compartments

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can be monitored by spectroscopic (e.g., fluorescence correlation spectroscopy (FCS)) or microscopic (e.g., confocal laser scanning microscopy (CLSM)) techniques or by impedance or current recordings. Planar lipid bilayers are incompatible with high-resolution structural methods but offer unparalleled advantages for functional investigations of channels and transporters. For instance, Betz and co-workers (Langosch et al., 1991; see also Section 5.1) reconstituted peptides corresponding to GlyR TM helices 2 and 4 into spanned bilayers and demonstrated channel-like conductance profiles for TM helix 2 but not for TM helix 4. Channel activity of TM helix 2 was corroborated and it was furthermore shown that TM helix 1 of GlyR is not conductive (Reddy et al., 1993). Tomich and co-workers (Reddy et al., 1993; Wallace et al., 1997; Shank et al., 2006) systematically dissected channel function by analysis of more than 200 different TM helix 2 sequences. These efforts were crowned by the discovery of peptide variants that are better water-soluble than the wild-type peptide but still give rise to functional, anion-selective channels when embedded in lipid bilayers. This could represent a major step toward the ultimate goal of applying designed TM peptides as therapeutic channel replacements in fighting channelopathies like cystic fibrosis (Shank et al., 2006; see also Section 7.2). 7. Biological membranes Biological membranes are the most complex and challenging environments for investigating TM peptides. In addition to an asymmetric lipid bilayer matrix, native membranes contain many different proteins and glycoproteins which, depending on cellular compartment and cell type, can literally outweigh their lipidic components. Cellular membranes exist in an environment crowded by macromolecules, which favors reactions that decrease the occupied volume, such as oligomerization (reviewed in Zhou et al., 2008) or adoption of a TM orientation (Aisenbrey et al., 2008). Also, cellular membranes are spatially confined by both intracellular components such as the cytoskeleton and extracellular structures such as cell walls in plants. Moreover, membranes in live cells are exposed to the action of a huge arsenal of enzymes and trafficking systems that continuously work to maintain or change membrane composition, curvature, fluidity, permeability, and so forth. For these reasons, biological membranes are more difficult to study than any of the membrane-mimetic systems treated so far. Assays performed in cellular membranes are inherently more susceptible to experimental fluctuations and poor reproducibility because seemingly constant external conditions like incubation temperature, solution pH, etc. cannot guarantee identical intracellular conditions. Also, changing a single external parameter inevitably entails multiple, and often intricately interdependent, responses at the cellular or organismic level. Notwithstanding these difficulties, experiments in the context of intact biological membranes can furnish insights into TM peptide functions otherwise not available (Table 3). This is particularly true and relevant whenever TM peptides are envisioned for pharmacological applications. In this chapter, we recapitulate some assays specifically designed for looking into TM domain oligomerization in cell-based assays (Section 7.1) and touch on some promising pharmacological applications of TM peptides (Section 7.2). 7.1. Quantification of helix–helix interactions in cell culture Many of the methods described so far are not easily applicable to cell-based systems. Instead, a wide spectrum of microscopic techniques and enzyme-based assays are available. Also, quantification

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Table 3 Overview of membrane-mimetic systems of various complexities. Techniques used to study TM peptides Advantages Isotropic solvents Very easy to use High solubility of TM regions Cheap

Detergent micelles Easy to prepare and use High solubility Moderate light scattering Fast tumbling Lipid bilayers Native-like environment Bilayer asymmetry possible Transmembrane transport measurable

Biological membranes Native environment Organismic level Transmembrane transport measurable

Disadvantages

Applicable

Not applicable

No anisotropy Poor solubility of hydrophilic flanks Disruption of tertiary structure

AUC, CD, cross-linking, DSC, IR, ITC, MD, NMR, SEC (Fl, DLS)

CR, cell-based assays, enzymatic assays, EPR, Mi, PAGE, SPR

Poor detergent packing Pronounced dynamics Changes in micellar size and shape

AUC, CD, cross-linking, enzymatic assays, EPR, Fl, IR, MD, NMR, PAGE (DLS, SEC, SPR)

CR, cell-based assays, Mi

Reconstitution necessary Substantial light scattering Slow tumbling Expensive

CD, cross-linking, enzymatic assays, EPR, Fl, IR, ITC, MD, NMR, CR, SPR (AUC, DLS, DSC, Mi)

Cell-based assays, PAGE, SEC

High complexity Difficult to control Interference with other cell components/functions No high-resolution structures

CR, cell-based assays, cross-linking, enzymatic assays, Fl, Mi (NMR)

AUC, CD, DLS, DSC, IR, ITC, EPR, MD, PAGE, SEC, SPR

Handling

Disturbance of tertiary structure (denaturation)

Complexity Methods enclosed in parentheses are, in principle, applicable but are not applied routinely. CR, conductance recordings (of vesicles, lipid bilayers, or cells); cell-based assays: ToxR, TOXCAT, GALLEX, POSSYCAT (selectable TOXCAT-based assay); DLS, dynamic light scattering; DSC, dynamic scanning calorimetry; EPR, electron paramagnetic resonance; Fl, fluorescence-based methods; Mi, microscopy; SPR, surface plasmon resonance.

of translocon-driven insertion of TM domains into biological membranes is possible with the aid of an in vitro assay based on leader peptidase (Hessa et al., 2005, 2007). Yet other approaches, such as NMR spectroscopy (Selenko and Wagner, 2007), are just being adapted to suit the needs of in-cell experiments. 7.1.1. Cell-based quantification of oligomerization Cell-based oligomerization assays like ToxR, TOXCAT or GALLEX are well-established tools to screen for helix–helix interactions in live cells (see also Table S1). They enable comparison of oligomerization affinities among various TM sequences (Gerber et al., 2004b) or quantification of binding specificity (Yin et al., 2006; Go et al., 2006). Common to all three assays is the incorporation of a guest TM sequence into a host protein containing a reporter moiety that responds to TM-driven dimerization. ToxR (Kolmar et al., 1995; Langosch et al., 1996; Gerber and Shai, 2001; Sal-Man et al., 2005) exploits the dimerization-dependent activity of the membranespanning transcriptional activator of ctx promoter. TOXCAT (Russ and Engelman, 1999; Melnyk et al., 2004; Duong et al., 2007) is a modification of ToxR that relies on the enzymatic activity of chloramphenicol acetyl transferase (CAT) rather than transcriptional activation. GALLEX (Schneider and Engelman, 2003; Finger et al., 2006) is based on ␤-galactosidase repression upon TM heterodimerization of two chimeric proteins containing LexA DNA binding domains with different DNA sequence specificity. Once established, systematic point mutations or scans of randomized sequence libraries can be performed with reasonable effort (Russ and Engelman, 1999), as isolation and purification steps are omitted. In a second round of experiments, the affinity and specificity of TM-driven dimerization can be tested using synthetic TM peptides (Gerber et al., 2004a; Yin et al., 2006; Go et al., 2006). For instance, binding of an all-d peptide (Gerber and Shai, 2002) or a peptide with two d-amino acids (Gerber et al., 2004a) derived from GpA to an all-l GpA TM helix was shown possible. Despite the

mirror image structure of the all-d peptide, the helix–helix contact surface of the heterodimer differs surprisingly little from that of the wild-type homodimer, as suggested by MD simulations (Gerber and Shai, 2002). All-d peptides are much less susceptible to proteolytic degradation and therefore exhibit prolonged stability in the body, thus overcoming a major bottleneck in the pharmacological application of peptides (see next section). 7.2. Pharmacological applications of TM peptides Potential pharmacological applications of TM peptides include modulation of and interference with membrane protein function, replacement of channel activity by synthetic peptide pores, and delivery of therapeutic or diagnostic agents to specific tissues, to name but a few. 7.2.1. Interference with membrane protein function Bouvier and co-workers (Hebert et al., 1996) used a peptide corresponding to TM helix 6 of ␤2 -adrenergic receptor (␤2 AR) to inhibit receptor dimerization and function in a stably transfected Chinese hamster fibroblast cell line. These experiments demonstrated that TM domains of ␣-helical membrane proteins constitute potential drug targets that can be tackled by competitively binding TM peptides. Thereafter, several other peptidic inhibitors of membrane protein oligomerization and function were reported, including peptides against ErbB2 (Gerber et al., 2004b), ErbB1 (Bennasroune et al., 2004), and Escherichia coli methyl-accepting chemotaxis protein II (MCP-II, formerly known as aspartate receptor Tar; Sal-Man et al., 2004, 2005; see also Table S1). There are also a few examples of TM peptides tested in animal models. O’Dowd and co-workers (George et al., 2003) used several TM peptides derived from different G-protein-coupled receptors (GPCRs) and other multispanning membrane proteins. Peptides corresponding to TM helices 7 of D2 dopamine receptor,

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␣1 -adrenergic receptor, ␤1 -adrenergic receptor, and vasopressin V2 receptor as well as to TM helix 12 of sodium-dependent dopamine transporter exhibited ␣-helical secondary structures when initially dissolved in DMSO. The peptides were then supplemented with the detergent digitonin and subsequently diluted into buffer. The resulting peptide solutions, containing 8.3% DMSO and 0.4% digitonin, were applied as intracerebral or intravenous injections to male Wistar rats (Rattus norvegicus). All peptides selectively disrupted the functions of their respective parent proteins, opening the way for the development of potent and nontoxic peptide variants possessing higher aqueous solubilities. 7.2.2. Reconstitution of channel activity Another application of TM peptides arose from the observation of pentameric anion-selective pores formed by TM helix 2 of GlyR (see Sections 5.1 and 6.3). These pores were found not only in artificial lipid bilayers but also in MDCK cell culture (Wallace et al., 1997; Tomich et al., 1998; Broughman et al., 2001, 2002b; Cook et al., 2004; Shank et al., 2006). A water-soluble, channel-forming, and ion-selective variant of this peptide holds some promise for the pharmacological use of TM domains to cure channelopathies like cystic fibrosis (Shank et al., 2006). 7.2.3. Delivery of pharmacological tools using TM peptides A further pharmacological application of some TM peptides arises from their ability to self-insert into lipid bilayers under specific conditions. Such peptides could eventually be employed to deliver drugs or biomarkers in a controlled manner and thus may fulfill important therapeutic or diagnostic functions. Promising results in this direction were obtained by applying pHLIP (see Section 6.2) to mice (Mus musculus; Andreev et al., 2007). Owing to its pH-dependent membrane insertion capability, this peptide specifically targets tissues characterized by low pH such as arthritic or tumor tissues. As peptide accumulation in the kidney might pose problems, renal pH was elevated by supplying the mice with water rich in bicarbonate. Further improvement was achieved with an all-d pHLIP analogue, which is most likely due to its prolonged metabolic stability. 8. Conclusions The “divide and conquer” approach to ␣-helical membrane proteins draws its justification from the central tenet of the twostage model of membrane protein folding, which poses that TM ␣-helices are independently stable protein domains (see Section 2.2). Indeed, a considerable body of evidence underpins this view, as many peptides derived from TM domains of single- or multispanning membrane proteins retain a number of characteristic properties on isolation from their tertiary and quaternary structural contexts. The potential of this approach is illustrated by the numerous and diverse questions that have been answered with the aid of TM peptides, some of which are highlighted in this review. However, it must not be concealed that many questions remain to be answered, especially on a quantitative level, and that several findings challenge the simplistic picture of the two-stage model and, consequently, the “divide and conquer” approach to membrane proteins. In view of the examples discussed in this contribution, the following issues emerge as focal points for future research on protein-derived TM peptides: (i) Already in one of the first applications of the “divide and conquer” approach, Engelman and co-workers (Hunt et al., 1997a) discovered that TM helices 6 and 7 of bacteriorhodopsin are not able to adopt independently stable ␣-helical TM conformations. By contrast, a helix–loop–helix motif encompassing

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both TM domains and the connecting loop assumes largely ␣-helical secondary structure in lipid bilayers (Barsukov et al., 1992). This exemplifies that the two-stage hypothesis is not fulfilled for all TM domains or, in other words, that some TM helices, particularly in multispanning membrane proteins, may depend on interactions with other TM domains, loop regions, or cofactors (Schneider et al., 2007; see Section 2.2). Characterizing a peptide derived from such a marginally stable TM helix in isolation might appear little enlightening, but studying its interactions with other protein domains could contribute substantially to a more comprehensive understanding of membrane protein folding (Engelman et al., 2003). (ii) A point mutant of TM helix 3 of bacteriorhodopsin, dubbed pHLIP, offers hitherto unique opportunities to quantitatively scrutinize the first step of the two-stage model using a natural TM sequence (Hunt et al., 1997b; Reshetnyak et al., 2007). However, the energetics of bilayer insertion of pHLIP seem poorly understood and deserve closer inspection (see Section 6.2). Factors like lipid composition, lateral pressure profile (see Section 2.1), hydrophobic mismatch (see Section 3.1), or hydrophilic flanking residues (see Section 3.2) affect peptide–lipid interactions but have not yet been assessed in much detail in natural TM domains. Moreover, it would be desirable to extend this approach to other proteinderived TM peptides, which have yet to be found, in order to assess to what extent lessons learned from pHLIP can be generalized. Deciphering peptide self-insertion not only will provide a more detailed understanding of the physicochemical principles underlying the first step of the two-stage model but might also open new therapeutic avenues (see Section 7.2). (iii) Homo- or heterooligomerizing TM domains derived from various membrane proteins allow for a quantitative analysis of the second step of the two-stage model, that is, helix–helix interactions within membranes or membrane-mimetic systems. For instance, dimerization of the GpA TM domain has been explored in detergent micelles (see Section 5.2), lipid bilayers (see Section 6.1), and biological membranes (see Section 7.1). However, even in supposedly simple model systems such as micelles, the thermodynamic characteristics of dimerization have thus far evaded straightforward explanations (Fisher et al., 1999, 2003; see Section 5.2). A more detailed comprehension of the second step of the two-stage model would not only be of fundamental interest to membrane protein folding but might also pave new ways in the rational selection of more suitable (“milder”) detergents for the isolation, purification, characterization, and reconstitution of ␣-helical membrane proteins. (iv) Finally, peptides derived from TM helices of single- or multispanning membrane proteins have recently entered areas of chemical biology and medicinal chemistry. They are being used as tools to probe, modulate, or mimic membrane protein function, as vehicles to deliver drugs or biomarkers, or as lead compounds in drug development (see Section 7.2). However, in order to establish a new generation of drugs that would target helix–helix interactions mediated by TM domains, a number of high hurdles have to be overcome. On top of the usual drawbacks encountered with peptidic lead compounds, such as fast metabolic clearance or potential immunogenecity, the vast majority of TM peptides suffer from extremely poor aqueous solubility and thus pose additional formulation problems (Moore et al., 2008). Although some of these challenges have been resolved for some TM peptides (see Sections 3.2 and 7.2), it remains to be seen how these approaches translate to nonpeptidic small molecules targeting interhelical interactions in biological membranes.

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