This page describes the energetics of the folding of peptides in lipid bilayer interfaces and why partitioning of peptides into bilayers promotes secondary structure formation.

 

The germinal work of Kaiser and Kézdy (1) first demonstrated that the partitioning of peptides and small proteins into membranes often induces the formation of secondary structure, a process conveniently described as partitioning-folding coupling (2). For that to be true, it must also be true that the overall free energy cost of partitioning the folded peptide is significantly lower than for the unfolded peptide (3). This simple idea focuses attention on the free energy of folding of a peptide partitioned into the interface, ΔGif, and suggests the possibility of quantitative rules for the design of peptides with specified partition coefficients and secondary structure propensities

Given their observation that it is energetically costly to partition peptide bonds into the interface, Wimley and White (2) hypothesized that participation of peptide bonds in H-bonding would dramatically reduce a peptide's partitioning free energy and thus cause partitioning-folding coupling. This hypothesis is born out by two recent measurements of the energetics of α-helix (4) and β-sheet (5) formation summarized in Figures 1 and 2. These measurements suggest that the per-residue free energy reduction, ΔGresidue, accompanying secondary structure formation is typically −0.5 kcal mol-1. This value allows one to estimate ΔGif from the free energy ΔGwiu of partitioning the unfolded peptide from water into the interface (see Energetics of Protein-Bilayer Interactions).
Although the primary driving force for partitioning-folding coupling arises from the free energy reduction associated with H-bonding, ΔGresidue is probably not due to this effect alone. Other interactions must also contribute, including the effects of folding/assembly entropy, side-chain packing, relative exposure of side-chains to membrane and water, and the depth of membrane penetration of secondary structure units. The depth of penetration of helices is likely to be strongly affected by the hydrophobicity-hydrophilicity pattern of the peptide sequence that defines the hydrophobic moment (6). Jacobs and White (3) proposed the existence of a strong free energy gradient for driving peptides into the HC core due to the difference in the non-polar solvation parameters between the interface (−12 kcal mol−1Å−2) and the HC core (−25 kcal mol−1Å−2) (see Equation 2 of Energetics of Protein-Bilayer Interactons). The location of an amphipathic helix in the bilayer interface determined by X-ray diffraction is consistent with that idea. The general amino acid composition of a peptide also appears to be important (7, 8, 9), and there is evidence of important connections between helicity, hydrophobic moment, and lipid charge on the membrane activity of amphipathic helices (10, 11).

Partitioning-Folding Coupling of an a-helix: Melittin

Figure 1

The partitioning-folding coupling of melittin in POPC bilayer interfaces is summarized in Figure 1 [based on (4)]. Melittin, a 26-residue peptide that folds into an amphipathic helix, is monomeric at low concentrations in aqueous solution where it exists in a disordered state with low α-helical content. When bilayer vesicles are titrated into a melittin solution, however, the fraction of melittin partitioned into the vesicles and the average melittin helicity increase concomitantly (12, 13), indicating bilayer induction of secondary structure. The presence of a very distinct isodichroic point in the CD spectra (12, 13) demonstrates a two-state transition in which there are only two populated states: monomeric melittin in water with low helicity and membrane-bound melittin with high helicity. Because of the very tight coupling of folding to partitioning, the energetics of partitioning unfolded melittin are not accessible directly. To access these energetics, the virtual unfolded bound state of melittin is emulated by a melittin diastereomer containing four D-amino acids that inhibit folding (14). The free energies of transfer ΔG of diastereomeric melittin and melittin were determined from mole-fraction partition coefficients and the changes in helicity from CD measurements.

Partitioning-Folding Coupling of a β-Sheet: A Hexapeptide Model System

Figure 2

The hexapeptide AcWL5 avidly forms β-sheet aggregates in a highly cooperative manner upon partitioning into phosphocholine bilayers. Because AcWL5 is strictly monomeric in the aqueous phase, β-sheet formation is induced by the bilayer. The partitioning and aggregation of AcWL5 into phosphocholine bilayer interfaces is summarized in Figure 2 [based on (5)]. Partitioning and other measurements reveal the details of the aggregation process which is temperature dependent and reversible (5). The structures of the monomers and aggregate of AcWL5 are shown for illustrative purposes only, and should not be taken literally. Here the aggregate is shown as containing 8 monomers; the average aggregate size is actually 10 or greater.

 

References (linked to PubMed)

  1. Kaiser ET & Kézdy FJ (1983). Proc. Natl. Acad. Sci. USA 80:1137-1143.
  2. Wimley WC & White SH (1996). Nature Struct. Biol. 3:842-848.
  3. Jacobs RE & White SH (1989). Biochemistry 28:3421-3437.
  4. Ladokhin AS & White SH (1999). J. Mol. Biol. 285:1363-1369.
  5. Wimley WC, Hristova K, Ladokhin AS, Silvestro L, Axelsen PH & White SH (1998). J. Mol. Biol. 277:1091-1110.
  6. Eisenberg D, Schwarz E, Komaromy M & Wall R (1984). J. Mol. Biol. 179:125-142.
  7. Li S-C & Deber CM (1994). Nature Struct. Biol. 1:368-373.
  8. Deber CM & Li S-C (1995). Biopolymers 37:295-318.
  9. Blondelle SE, Forood B, Houghten RA & Pérez-Payá E (1997). Biopolymers 42:489-498.
  10. Dathe M, Schümann M, Wieprecht T, Winkler A, Beyermann M, Krause E, Matsuzaki K, Murase O & Bienert M (1996). Biochemistry 35:12612-12622.
  11. Wieprecht T, Dathe M, Epand RM, Beyermann M, Krause E, Maloy WL, MacDonald DL & Bienert M (1997). Biochemistry 36:12869-12880.
  12. Vogel H (1981). FEBS Lett. 134:37-42.
  13. White SH, Wimley WC, Ladokhin AS & Hristova K (1998). Methods Enzymol. 295:62-87.
  14. Oren, Z. and Shai, Y. (1997). Biochemistry 36:1826-1835.