Like all cellular proteins, membrane proteins are synthesized by ribosomes. But unlike their soluble counterparts, highly hydrophobic membrane proteins require ancillary proteins to prevent aggregation in aqueous cellular compartments. The principal ancillary protein is the translocon that works in concert with ribosomes to manage the orderly insertion of α-helical membrane proteins directly into the endoplasmic reticulum membrane of eukaryotes or the plasma membrane of bacteria. In the course of insertion, membrane proteins come into thermodynamic equilibrium with the lipid membrane where physicochemical interactions determine the final three-dimensional structure. Much progress has been made during the past several years toward understanding the physical chemistry of membrane protein stability, the structure of the translocon, and the mechanisms by which it selects and inserts transmembrane helices. This progress is reviewed in these pages, which are based upon recent reviews by White & von Heijne (2008) and Cymer, et al (2015).



Figure 1

Figure 1. Components of membrane protein assembly.

(a) The components of membrane protein assembly. (a1) A ribosome translating the mRNA of a protein targeted for secretion across or insertion into membranes and a signal of a recognition particle (SRP), which is a GTPase. The structures of ribosomes are reviewed in (18, 62) and the structure of SRP in (54).
(a2) The ribosome and SRP recognize a signal peptide as it emerges from the ribosome exit tunnel, bind together, and cause arrest of elongation (27, 28, 82).
(a3) The ribosome-SRP complex binds to the membrane-bound SRP receptor (SR), another GTPase, that associates dynamically with the translocon. Prokaryotes use a simplified SRP (Ffh) and SR (FtsY) that associate to form a quasi-two-fold symmetrical dimer (20). The binding of SRP to SR causes reciprocal stimulation of their GTPase activities, causing transfer of the signal peptide to the translocon and resumption of elongation, a process that is just beginning to be understood (2).
(a4) Proteins targeted for translocation are secreted into the periplasm (bacteria) or ER lumen (eukaryotes) whereas the stop-transfer signals of MPs are transferred to the membrane bilayer.

(b) Electron cryo-microscopic image of the canine ribosome-translocon (Sec61) complex. The small and large ribosome subunits are indicated by S and L, respectively. Modified with permission from Figure 4 of Ménétret et al. (52).

(c) Structure of a single SecYEβ closed-state translocon heterotrimer from Methanococcus jannaschii (70) that has been embedded in a POPC lipid bilayer using molecular dynamics methods. SecY is viewed from the ribosome along the bilayer normal. The presumed lateral exit from the TM7/TM2b lateral gate is indicated. Red, phospholipid headgroups; white, acyl chains.

(d) SecYEβ viewed along the bilayer plane toward the lateral gate through which nascent TM helices are believed to move into the bilayer. The TM2A 'plug helix' apparently seals the translocon in the absence of nascent peptide. The images in c and d were prepared from a molecular dynamics simulation, courtesy of Alfredo Freites at UC Irvine. Both molecular graphics images were produced using Visual Molecular Dynamics (VMD) (36).

Proteins destined for transmembrane export (translocation) or insertion are generally managed by the concerted action of translating ribosomes in the cytoplasm and translocon complexes located in the endoplasmic reticulum (ER) of eukaryotes or the plasma membrane of bacteria. The operating principles for the membrane protein assembly (8, 15, 19, 40, 58, 75) are summarized in Figure 1a.

The critical membrane-protein component of the translocon complex is heterotrimeric Sec61 in eukaryotes or the highly homologous SecYEG in bacteria. Cryo-EM image reconstructions (Figure 1b) of native ribosome-translocon complexes (RTC) (52) suggest that the complex is likely composed of two dimers of the Sec61 heterotrimer and two copies of the tetrameric translocon-associated protein (TRAP). At least three other proteins associate closely with the translocon complex, but do not seem to be part of the RTC seen in the image reconstructions. These are the translocating chain-associated membrane protein (TRAM) (16, 25); the signal peptidase complex (SPC) (21), which cleaves signal sequences; and oligosaccharyl transferase (OST) (11), which N-glycosylates -Asn-X-Ser/Thr- sites on membrane and secreted proteins.

The translocon complex acts as a switching station: Secretory proteins are allowed to pass straight through into the ER lumen or the bacterial periplasm (secretion), while TM segments of membrane proteins are shunted into the membrane bilayer. Deciphering the code that the translocon uses for selecting elongating segments for TM insertion is of fundamental importance for understanding the folding of membrane proteins (see below). But the selection of TM segments is only the first step in the complex process of gathering the TM segments together to form the native protein structure (12, 65, 66).

Structure of the Translocon

The key protein of the eukaryotic translocon complex-the one that acts as the switching station-is heterotrimeric Sec61αβγ (SecYEG in eubacteria; SecYEβ in archaea) (56). The Sec61 α-subunit has ten TM helices whereas β and γ typically have one TM helix (eubacterial SecE has three and SecG has two TM helices). Van den Berg et al. (70) have determined the crystallographic structure of SecYEβ from Methanococcus jannaschii at a resolution of 3.8 Å. It is shown embedded in a lipid bilayer in Figures 1c and 1d. The images are snapshots from a molecular dynamics simulation of the heterotrimer embedded in a palmitoyloleoylphosphatidylcholine bilayer (73).

Comparisons of the SecYEβ crystallographic structure with cryo-EM reconstructions (52) suggested that the heterotrimers form a tetramer arranged as a dimer-of-dimers arranged in a back-to-back configuration ('back' is defined in Figure 1c). No nascent peptide was observed in the crystallographic structure, which is thus assumed to be in a closed state. Disulfide cross-linking experiments (10), however, revealed that elongating chains pass through the so-called hydrophobic collar in the middle of SecY (Figure 1d), suggesting that translocon-mediated protein export and membrane insertion involves at any particular time only one of the SecY/Sec61 heterotrimers in the translocon complex. The broad purpose of the posited tetrameric association of SecY/Sec61 may be to provide an assembly platform enabling the ribosome and other members of the Sec family to secrete or insert nascent chains (55).

Figure 1c shows SecYEβ from the viewpoint of the ribosome and Figure 1d a view parallel to the membrane. The 10 TM helices of SecY are arranged to form an inverted 'U' (Figure 1c) with TM helices 1-5 (colored green, except for TM2B, which is red) forming one leg and helices 6-10 (colored orange, except for red TM7) forming the other. The two sets of helices have a pseudo-symmetric two-fold rotation axis in the plane of the membrane and are connected at the back by an external loop. This loop and the single TM helix of SecE prevent lipids from contacting the interior of SecY from the backside. The only possible opening from the interior into the lipid bilayer is through the so-called lateral gate formed by TM2B and TM7 (Figures 1c, d), which is hypothesized to control passage of nascent TM helices into the bilayer from the hourglass-shaped water-filled interior of SecY (Figure 1d). The two halves of the hourglass are separated by a ring of hydrophobic residues (hydrophobic collar) that are believed to act as seal around the elongating chain.

Sitting just below the hydrophobic collar is a short helix (TM2A) that apparently acts as a 'plug' to block passage of small molecules through the translocon in the closed state. Van den Berg et al. (70) hypothesized that the plug is displaced by nascent protein translocation. But the necessity for the TM2A plug for sealing the hourglass in the absence of a translocating nascent chain was discounted in a study of a so-called plugless Sec61/SecY mutant (41, 49), because excision of TM2A was found to have no effect on the viability of yeast cells. Quite remarkably, however, a crystallographic study of plugless SecY (45) showed that in fact SecY restructures itself in the absence of TM2A to form a new plug!

The image of SecYEβ in a lipid bilayer (Figure 1c, d) is entirely consistent with the idea that TM helices move into the lipid membrane from the water-filled protein conducting channel by a simple partitioning process, as suggested by cross-linking studies of nascent chains (29, 50). In such a scheme, sufficiently hydrophobic helices prefer the bilayer whereas more polar helices favor the translocon, and ultimately the aqueous phase. That is, the translocon and the lipid bilayer work in concert to decipher the code for TM helices embedded in the amino sequence. If this view is correct, then the big question concerns the code for deciphering the process. Answers to this question should lead to major improvements in the prediction of membrane protein structure.


Translocon Recognition of Transmembrane Helices

A Biological Hydrophobicity Scale

Figure 2

Figure 2. Determination of a biological hydrophobicity scale.

Integration of designed TM segments (H-segments) into the endoplasmic reticulum using dog pancreas microsomal membranes. This system was used to explore systematically the hydrophobicity requirements for TM helix integration via the Sec61 translocon (32).

(a) Wild-type leader peptidase (Lep) from E. coli has two N-terminal TM segments (TM1, TM2) and a large luminal domain (P2). H-segments, flanked by glycosylation sites (G1, G2), were inserted between residues 226 and 253 in the P2-domain. For H-segments that integrate into the membrane, only the G1 site is glycosylated (left), whereas both the G1 and G2 sites are glycosylated for H-segments that do not integrate into the membrane (right). Redrawn from Hessa et al. (32).

(b) An example of SDS gels used in the in vitro determination of the extent of glycosylation of Lep/H-segment constructs in the absence (-RM) and presence (+RM) of dog pancreas rough microsomes.

(c) Equations used by Hessa et al. (32) for the analysis of gels of the type shown in panel b.

(d) Mean probability of insertion, ρ, for H-segments with n = 0 - 7 Leu residues. The curve is the best-fit Boltzmann distribution, which suggests equilibrium between the inserted and translocated states of the H-segments.

(e) Biological ΔGaaapp scale derived by Hessa et al. (32) from H-segments with the indicated amino acid placed in the middle of the 19-residue hydrophobic stretch.

(f) Correlation between ΔGaa app and the WW octanol free energy scale (ΔGaaWW). Data in panels b - e replotted from Hessa et al. (32).

Insights into the process of TM helix insertion have been obtained by Hessa et al. (32) using an in vitro expression system (64) that permits quantitative assessment of the membrane insertion efficiency of model TM segments (Figure 2). Specifically, they examined the integration into membranes of dog pancreas rough microsomes (RMs) of designed polypeptide segments (H-segments) engineered into the luminal P2 domain of the integral membrane protein leader peptidase (Lep, Figure 2a). Because glycosylation of the engineered Asn-X-Ser glycosylation sites (G1 and G2, Figure 2a) can occur only in the lumen of the RMs, H-segment TM insertion could be distinguished from secretion by simple gel assays (Figure 2b). The relative fractions of singly (1g) and doubly (2g) glycosylated molecules allow quantitative assessment of insertion versus secretion (Figure 2c). The first experiments, carried out using H-segments of the form GGPG-(LnA19-n)-GPGG with n = 0 to 7, revealed that the probability of insertion, p(n), conformed accurately to a Boltzmann distribution. This showed that translocon-mediated insertion has the appearance of an equilibrium process. Given this key observation, the insertion of H-segments were quantitated using the apparent free energy of insertion ΔGapp (Figure 2c).

A 'biological' hydrophobicity scale (ΔGaaapp) (Figure 2e) could be derived from studies in which each of the 20 naturally occurring amino acids were placed in the middle position of H-segments containing various numbers of Leu and Ala residues chosen to maintain p ≈ 0.5 (ΔGapp ≈ 0), which is the region of maximum sensitivity of the assay (Figure 2e). Considering the complexity of the biological system, the scale correlated surprisingly well (Figure 2f) with the WW octanol scale. Their overall high correspondence implies that the recognition of TM segments by the translocon likely involves direct interaction between the segment and the surrounding lipid (29), which seems reasonable in the light of Figure 1d.


Position-Dependence of Free Energies

Figure 3

Figure 3. Position-dependence of ΔGaaapp and helix-helix interactions in membrane protein assembly.

(a) Scan of a single Arg residue across H-segments of composition 1R/6L/12A. The position of the Arg in the 19-residue hydrophobic stretch is shown on the x-axis. The locations of the Arg in the KvAP S4 helix are indicated by the red circles. Data replotted from (34). This plot reveals the strong position-dependence of ΔGArgapp. Lys, Asp, and Glu residues show a similar position-dependence.

(b) Molecular dynamics simulation of a model S4 voltage-sensor peptide (GGPG-LGLFRLVRLLRFLRILLII-GPGG) in a POPC bilayer. (left panel) Cut-away view of the simulation system, showing bilayer distortion around the peptide and the contacts between phosphate groups, water molecules, and arginine guanidinium groups. (right panel) This space-filling representation of the hydrophilic neighborhood of the S4 helix, represented as Connolly surfaces, reveals a 10 Å gap that is never occupied by the Arg guanidinium groups because of snorkeling to the bilayer interface. Color code: red, water; yellow, phosphocholine headgroups; green, acyl chains; white, GGPG…GPGG flanks; silver, non-Arg S4 residues; blue, guanidinium groups. Images modified from (24) with permission.

(c) H-segment pair-scans for Phe, Trp, and Tyr residues in which pairs of residues are moved symmetrically toward the N- and C-termini of the sequence. The compositions of the H-segments are indicated. The ΔGapp value for the 4L/15A H-segment is indicated by the dashed line. This shows that the Trp residues have the same apparent free energy as Leu when placed near the ends of the H-segment. Data replotted from (32).

Does ΔGaaapp vary with position within the H-segment? To answer this question, Hessa et al. (32) performed position scans of two types: single- and pair-scans. In the simpler single-scan, an amino acid of interest was placed at different positions in the H-segment sequence and ΔGapp determined. The dramatic results from an Arg scan are shown in Figure 3a (34). Similar results were found for Lys, Asp, and Glu scans. The strong dependence on position must be related to the relative ease of snorkeling of the charge group to the wet bilayer interface (46) - the farther the charge is from the interface, the greater the energetic cost. The strong position-dependence of Arg explains why it is possible for Sec61 to insert the KvAP S4 voltage-sensing helix, which contains four Arg residues, across the ER membrane with ΔGapp ≈ 0 (34). A molecular dynamics simulation of S4 across a lipid bilayer (24) showed that the arginines snorkel to the bilayer interface to form salt-bridges with the phospholipid phosphates and hydrogen-bond networks with water (Figure 3b).

In pair-scans, a pair of residues of a given kind were moved symmetrically from the center of the H-segment towards its N- and C-termini to preclude the possibility of a shift in helix position across the membrane. Pair-scans of charged residues were found to be consistent with single scans, suggesting that helix shifts were not significant. Pair-scans of the aromatic residues, which are known to have preferential interactions with the bilayer interface (42, 86, 88), gave another insight into TM helix insertion. The behaviors of Trp and Tyr were quite dramatic (Figure 3c): When placed centrally, they strongly reduced membrane insertion, but became much less unfavorable as they were moved apart. Indeed, Trp was as favorable as Leu when placed in the outermost positions (dashed line, Figure 3c). The position-dependence of Phe was quite different from those of Trp and Tyr (blue curve, Figure 3c), which is interesting, because Phe does not have a strong interfacial preference in membrane proteins (69, 71). The wave-like pattern observed for the Phe pair-scan is a result of variations in the hydrophobic moment (amphiphilicity) of the helices (32). These results provided further evidence supporting the idea that protein-lipid interactions are central to the recognition of TM helices by the translocon.

Prediction of Transmembrane Helices

The strong position-dependence of ΔGaaapp meant that the base biological hydrophobicity scale would be of limited value for predicting TM helices by simple hydropathy plot methods; accurate predictions require accounting for the position-dependence of . In a recent study, Hessa et al. (33) carried out a comprehensive examination of the position-dependence of ΔGaaapp. In addition, they determined how the overall length of the H-segment affected . The data enabled them to derive a simple expression for calculating the expected for H-segments given the amino acid sequence and overall length:

ΔGpredapp= ∑li=1 ΔGaa(i)app+ c0 ⋅ μ + c1 + c2l + c3l2    (1)

where l is the length of the segment, ΔGaa(i)app is the matrix element giving the contribution from amino acid aa in position i, μ is the hydrophobic moment, c0 is the weight parameter for the hydrophobic moment, and the terms c1 + c2 + c3l2 account for the dependence of ΔGapp on segment length. The optimized matrix was derived by minimizing the sum of the squared difference. A web server for calculating ΔGapp-profiles across a protein sequence is available at and is included in MPEx.

Figure 4

Figure 4. Distributions of ΔGpredapp values in cytoplasmic, secreted, and transmembrane proteins computed from Eq. (1). The 17-to-33-residue segment with lowest ΔGpredapp was identified in 670 cytoplasmic (green), 1012 secreted (blue), and 349 single-spanning transmembrane proteins (excluding signal peptides; black), while for 508 TM helices from multi-spanning transmembrane proteins of known 3D structure (red), the 17-to-33-residue segment with lowest within each annotated helix (plus 10 residues on either side) was identified instead. Data points show the relative frequency of proteins with ΔGpredapp within ±0.5 kcal mol-1 of the value given by the x-axis. Data replotted from (33).

Distributions of ΔGapp values obtained for mammalian secreted proteins as well as single- and multi-spanning membrane proteins are shown in Figure 4. The overlap between the ΔGapp distributions for the single-spanning transmembrane proteins (blue curve) and the secreted proteins (black curve) is small, and the two distributions cross close to the zero-point on the scale defined by the experimental analysis of the designed H-segments. A surprisingly large fraction (25%) of the TM helices in the multi-spanning membrane proteins of known 3D structure have ΔGapp > 0 kcal mol-1 (red curve). Such segments would presumably be only inefficiently recognized as TM helices by the translocon if they were the only hydrophobic segment in a protein. This suggests that TM helices in multi-spanning membrane proteins may depend on interactions with neighboring TM helices for proper partitioning into the membrane. Indeed, a number of such cases have been described in the literature (66), though their overall incidence has been unclear.

The results of these studies by Hessa et al. (32-34) suggest that direct protein-lipid interactions are essential for the recognition of TM helices by the translocon, and support models based on a partitioning of the TM helices between the Sec61 translocon and the surrounding lipid. The details of the partitioning process remain to be determined, but presumably the open state of the translocon is a highly dynamic one that permits rapid sampling of the translocon-bilayer interface by the translocating polypeptide.

Membrane Insertion of Multi-Spanning Proteins

How does the Sec61 translocon handle proteins with multiple transmembrane helices? The most revealing study published so far focused on the 6TM protein aquaporin 4 (65). By a very extensive analysis using site-specific cross-linkers introduced into each of the transmembrane helices, the authors arrived at a detailed picture of when during biosynthesis each TM helix exits the translocon and enters into the lipid bilayer. In general, the helices were observed to follow each other into the membrane in a strict N-to-C-terminal succession. Certain helices, however, would first completely exit the translocon only to revisit it at a later stage when a downstream helix had just entered the translocon channel. One is thus left with a picture of a very dynamic translocon that allows multiple transmembrane helices to interact with each other at early stages of membrane integration. In this way, one may envision a mechanism whereby TM helices that would not by themselves be sufficiently hydrophobic to integrate efficiently into the membrane become embedded in the progressively folding protein.

Helix-Helix Interactions

What kinds of interactions might underlie helix-helix association during translocon-mediated membrane insertion into the lipid bilayer?

Figure 5

Figure 5

(a) In order to examine helix-helix interactions driven by hydrogen bonding, the native Lep H2-segment was replaced by an H2′ segment of the general composition L19-nNn or L19-nDn (n = 0, 1 or 2), and a 19-residue H-segment containing one or two Asn or Asp residues was inserted into the P2 domain. Redrawn from (51).

(b) To measure the effect of Asp- or Asn-mediated interactions between the H2′- and H-segments, two constructs were compared for each H-segment: one with a uniformly hydrophobic 19-Leu H2′-segment (I) and one with an H2′-segment in which one or two of the Leu residues were replaced by Asp or Asn residues (II). The interaction free energy is expressed as the difference (ΔΔGapp) in the apparent free energy of insertion of the H-segment between the two constructs. In the example shown, the H segment contains two Asp residues and the appropriate number of Leu and Ala residues to make ΔGlapp ≈ 0 kcal mol-1. Redrawn from (51).

It is well established that hydrogen bonding between polar residues like Asn or Asp can drive helix-helix interactions in both detergent micelles and biological membranes (13, 26, 89, 90), and can also facilitate the formation of helical hairpins during translocon-mediated insertion (31). Meindl-Beinker et al. recently examined (51) whether and to what extent inter-helix hydrogen bonding could drive the process of translocon-mediated transmembrane helix insertion itself, and whether the separation between the two helices within the sequence may influence any such interaction. To address these questions in a quantitative way, they extended the systematic approach established by Hessa et al. (32) to study the effects of mutual helix-helix interactions on the efficiency of membrane insertion, using the scheme shown in Figure 5a.

The experiments (51) yielded several important results (Figure 5b). First, different Asn- or Asp-containing H2′ sequences did not affect the insertion of a purely hydrophobic H-segment. Furthermore, little effect was seen when a signal peptidase cleavage site was introduced in H2′, or even when the entire H1-H2 region was replaced by the signal peptide from preprolactin. The H2′ sequence thus had little influence on ΔGapp when the H segment was composed only of hydrophobic residues (cf. left-hand side of Figure 5b).

Second, by analyzing model protein constructs in which zero, one, or two Asn or Asp residues were placed in two neighboring hydrophobic segments (H2′ and H), it was found that ΔGapp of a marginally hydrophobic H-segment was significantly reduced only if both the H2′ segment and the H segment contained two Asn or two Asp residues (right-hand side of Figure 5b) with a spacing of three, but not one or five, residues (i.e., when they are spaced one helical turn apart in both H2′ and H). These results suggest that inter-helix hydrogen bonds can form during Sec61 translocon-assisted insertion, and that H2′ remains in close enough proximity to the translocon to offer its hydrogen bond donor and acceptor sites to the incoming H segment even when the intervening loop is 150 residues long (30, 53, 65).

The Biology-Physics Nexus

The ΔGapp measurements of Hessa et al. (32-34) are fully consistent with the simplest model one can propose for how transmembrane helices are recognized by the ribosome-translocon complex: Helices are somehow allowed to partition into the surrounding lipid bilayer based on the free energy of interaction between the transmembrane segment and the lipid. This would explain the correspondence between the biological hydrophobicity scale and biophysical scales like that of Wimley-White, and it would explain why the positional variations in ΔGapp for residues such as Arg, Trp, Tyr, Phe, and Gly (32, 34) match the statistical distribution of these residues across the membrane in the high- resolution X-ray structures (69). The data at hand thus speak strongly in favor of direct protein-lipid interactions as being the main driving force for the integration of single transmembrane helices, although the translocon may affect the ability of pairs or higher order assemblages of transmembrane helices to interact among themselves before partitioning into the bilayer (33, 51).

Although much remains to be done in order to understand fully the results obtained with the Sec61 translocon system, it is notable that the H1 and H2 transmembrane helices present in the model protein (Figure 5a) do not seem to affect the results in any significant way, as they can be replaced by a cleavable signal peptide with little effect on the measured ΔGapp values (51). Moreover, position-specific contributions to ΔGapp obtained by single-scans of a charged or polar residue along an H-segment predict ΔGapp values for H-segments using symmetrical pair scans, or event natural transmembrane helices with multiple charged residues within ~1 kcal mol-1 (32, 34). This suggests that vertical sliding of the H-segments used in the derivation of the 'biological' hydrophobicity scale is not a serious problem.

This is probably not the whole story, however. Many polar and charged residues, Arg included, have rather long and flexible side-chains, making it possible for them to 'snorkel' towards the lipid-water interface region. At the same time, lipid molecules located close to a transmembrane helix can adapt to the presence of polar residues, and water molecules can help solvate polar groups located well within the bilayer plane (17, 24, 39). One upshot of this dynamic picture of protein-lipid interactions is that ΔGapp profiles, such as the one shown in Figure 5a, most likely do not provide an accurate representation of the free-energy profile for moving a charged residue all the way across a membrane (as opposed to inserting it sideways from the translocon as part of a transmembrane helix). Presumably, if a helical peptide is pulled across a lipid bilayer, there is a substantial free-energy barrier (not seen in the ΔGapp profile) at the point when a charged residue has to flip its direction of snorkeling across the 10 Å hydrophobic gap (Fig. 5b) from one membrane surface towards the other (17). Seen from this perspective, one may regard the translocon as a device designed to lower the activation barrier for translocation of polar and charged residues across the membrane. It does so by providing an aqueous channel, while at the same time making it possible for consecutive segments of the nascent polypeptide to make 'lateral excursions' from the channel in order to test whether the free energy of membrane insertion is favorable or not.

Despite these caveats, it seems likely that the biological hydrophobicity scale is a good measure of the energetics of protein-lipid interactions in the true biological context, and as such will help us define the sequence determinants of membrane-protein assembly much more precisely than has been possible so far.


The research discussed on these pages was supported by grants from the Swedish Foundation for Strategic Research, the Marianne and Marcus Wallenberg Foundation, the Swedish Cancer Foundation, the Swedish Research Council, and the European Commission (BioSapiens) to Gunnar von Heijne and the National Institute of General Medical Sciences to Stephen White. We thank Michael Myers for editorial assistance.


References (linked to PubMed)

  1. Allen SJ, Curran AR, Templer RH, Meijberg W. & Booth PJ. (2004). J. Mol. Biol. 342:1279-91.
  2. Angelini S, Boy D, Schiltz E, Koch H-G. (2006). J. Cell Biol. 174: 715-24.
  3. Ash WL, Zlomislic MR, Oloo EO, Tieleman DP. (2004) Biochim. Biophys. Acta 1666: 158-89.
  4. Ben-Tal N, Ben-Shaul A, Nicholls A, Honig B. (1996). Biophys. J. 70: 1803-12.
  5. Ben-Tal N, Sitkoff D, Topol IA, Yang A-S, Burt SK, Honig B. (1997). J. Phys. Chem. B 101: 450-57.
  6. Benz RW, Castro-Román F, Tobias DJ, White SH. (2005). Biophys. J. 88: 805-17.
  7. Benz RW, Nanda H, Castro-Román F, White SH, Tobias DJ. (2006). Biophys. J. 91: 3617-29.
  8. Bibi E. (1998). Trends Biochem. Sci. 23: 51-55.
  9. Bowie JU. (1997). J. Mol. Biol. 272: 780-89.
  10. Cannon KS, Or E, Clemons WM, Shibata Y, Rapoport TA. 2005. J. Cell Biol. 169: 219-25.
  11. Chavan M, Yan A, Lennarz WJ. (2005). J. Biol. Chem. 280: 22917-24.
  12. Cheng Z, Gilmore R. (2006). Nat. Struct. Mol. Biol. 13: 930-36.
  13. Choma C, Gratkowski H, Lear JD, DeGrado WF. (2000). Nat. Struct. Biol. 7: 161-66.
  14. Curran AR, Engelman DM. (2003). Curr. Opin. Struct. Biol. 13: 412-17.
  15. Dalbey RE, von Heijne G. 2002. New York: Academic Press. 1-424 pp.
  16. Do H, Falcone D, Lin J, Andrews DW, Johnson AE. (1996). Cell 85: 369-78.
  17. Dorairaj S, Allen T.W. (2007). Proc. Natl. Acad. Sci. USA 104: 4943-48.
  18. Doudna JA, Rath VL. (2002). Cell 109: 153-56.
  19. Driessen AJM, Manting EH, van der Does C. (2001). Nat. Struct. Biol. 8: 492-98.
  20. Egea PF, Shan S-O, Napetschnig J, Savage DF, Walter P, Stroud RM. (2004). Nature 427: 215-21.
  21. Evans EA, Gilmore R, Blobel G. (1986). Proc. Natl. Acad. Sci. USA. 83: 581-85.
  22. Faraldo-Gómez JD, Smith GR, Sansom MSP. (2004). Eur. Biophys. J. 31: 217-27.
  23. Fleming KG, Ackerman AL, Engelman DM. (1997). J. Mol. Biol. 272: 266-75.
  24. Freites JA, Tobias DJ, von Heijne G, White SH. (2005). Proc. Natl. Acad. Sci. USA 102: 15059-64.
  25. Görlich D, Rapoport TA. (1993). Cell 75: 615-30.
  26. Gratkowski H, Dai Q-H, Wand AJ, DeGrado WF, Lear JD. (2002). Biophys. J. 83: 1613-19.
  27. Halic M, Becker T, Pool MR, Spahn CMT, Grassucci RA, et al. (2004). Nature 427: 808-14.
  28. Halic M, Blau M, Becker T, Mielke T, Pool MR, et al. (2006). Nature 444: 507-11.
  29. Heinrich SU, Mothes W, Brunner J, Rapoport TA. (2000). Cell 102: 233-44.
  30. Heinrich SU, Rapoport TA. (2003). EMBO J. 22: 3654-63.
  31. Hermansson M, von Heijne G. (2003). J. Mol. Biol. 334: 803-09.
  32. Hessa T, Kim H, Bihlmaier K, Lundin C, Boekel J, et al. (2005). Nature 433: 377-81.
  33. Hessa T, Meindl-Beinker NM, Bernsel A, Kim H, Sato Y, et al. (2007). Nature 450: 1026-1030.
  34. Hessa T, White SH, von Heijne G. (2005). Science 307: 1427.
  35. Hristova K, Dempsey CE, White SH. (2001). Biophys. J. 80: 801-11.
  36. Humphrey W, Dalke W, Schulten K. (1996). J. Mol. Graph. 14: 33-38.
  37. Jacobs RE, White SH. (1989). Biochemistry 28: 3421-37.
  38. Jayasinghe S, Hristova K, White SH. (2001). J. Mol. Biol. 312: 927-34.
  39. Johansson ACV, Lindahl E. (2006). Biophys. J. 91: 4450-63.
  40. Johnson AE, van Waes MA. (1999). Annu. Rev. Cell Dev. Biol. 15: 799-842.
  41. Junne T, Schwede T, Goder V, Spiess M. (2006). Mol. Biol. Cell 17: 4063-68.
  42. Killian JA, von Heijne G. (2000). Trends Biochem. Sci. 25: 429-34.
  43. Ladokhin AS, White SH. (1999). J. Mol. Biol. 285: 1363-69.
  44. Lemmon MA, Engelman DM. (1994). Q. Rev. Biophys. 27: 157-218.
  45. Li W, Schulman S, Boyd D, Erlandson K, Beckwith J, Rapoport TA. (2007). Mol. Cell 26: 511-21.
  46. MacCallum JL, Bennett WFD, Tieleman DP. (2007). J. Gen. Physiol. 129: 371-77.
  47. MacKenzie KR. (2006). Chem. Rev. 106: 1931-77.
  48. MacKenzie KR, Prestegard JH, Engelman DM. (1997). Science 276: 131-33.
  49. Maillard AP, Lalani S, Silva F, Belin D, Duong F. (2007). J. Biol. Chem. 282: 1281-87.
  50. Martoglio B, Hofmann MW, Brunner J, Dobberstein B. (1995). Cell 81: 207-14.
  51. Meindl-Beinker NM, Lundin C, Nilsson I, White SH, Von Heijne G. (2006). EMBO Rep. 7: 1111-16.
  52. Ménétret J-F, Hegde RS, Heinrich SU, Chandramouli P, Ludtke SJ, et al. 2005. J. Mol. Biol. 348: 445-57.
  53. Mitra K, Schaffitzel C, Shaikh T, Tama F, Jenni S, et al. (2005). Nature 438: 318-24.
  54. Nagai K, Oubridge C, Kuglstatter A, Menichelli E, Isel C, Jovine L. (2003). EMBO J. 22: 3479-85.
  55. Osborne AR, Rapoport TA. (2007). Cell 129: 97-110.
  56. Osborne AR, Rapoport TA, Van den Berg B. (2005). Annu. Rev. Cell Dev. Biol. 21: 529-50.
  57. Pastor RW, Venable RM, Feller SE. (2002). Acc. Chem. Res. 35: 438-46.
  58. Pfeffer S. (2003). Cell 112: 507-17.
  59. Phillips JC, Braun B, Wang W, Gumbart J, Tajkhorshid E, et al. (2005). J. Comput. Chem. 26: 1781-802.
  60. Popot J-L, Engelman DM. (1990). Biochemistry 29: 4031-37.
  61. Popot J-L, Engelman DM. (2000). Annu. Rev. Biochem. 69: 881-922.
  62. Ramakrishnan V. (2002). Cell 108: 557-72.
  63. Russ WP, Engelman DM. (2000). J. Mol. Biol. 296: 911-19.
  64. Sääf A, Wallin E, von Heijne G. (1998). Eur. J. Biochem. 251: 821-29.
  65. Sadlish H, Pitonzo D, Johnson AE, Skach WR. 2005. Nature Struct. Mol. Biol. 12: 870-78.
  66. Sadlish H, Skach WR. (2004). J. Membr. Biol. 202: 115-26.
  67. Tamm LK, ed. (2005). Weinheim: WILEY-VCH Verlag GmbH & Co KGaA. 444 pp.
  68. Tobias DJ. (2001). In Computational Biochemistry and Biophysics, ed. OM Becker, AD MacKerell, Jr., B Roux, M Watanabe, pp. 465-96. New York: Marcel Dekker.
  69. Ulmschneider MB, Sansom MSP, Di Nola A. (2005). Proteins 59: 252-65.
  70. Van den Berg B, Clemons WM, Jr., Collinson I, Modis Y, Hartmann E, et al. (2004). Nature 427: 36-44.
  71. Wallin E, Tsukihara T, Yoshikawa S, von Heijne G, Elofsson A. (1997). Protein Sci. 6: 808-15.
  72. White SH. (2003). FEBS Lett. 555: 116-21.
  73. White SH, Hessa T, von Heijne G. (2005). In Protein-Lipid Interactions. From Membrane Domains to Cellular Networks, ed. LK Tamm, pp. 3-25. Weinheim: WILEY-VCH
  74. White SH, Ladokhin AS, Jayasinghe S, Hristova K. (2001). J. Biol. Chem. 276: 32395-98.
  75. White SH, von Heijne G. (2004). Curr. Opin. Struct. Biol. 14: 397-404.
  76. White SH, Wiener MC. (1995). In Permeability and Stability of Lipid Bilayers, ed. EA Disalvo, SA Simon, pp. 1-19. Boca Raton: CRC Press.
  77. White SH, Wiener MC. (1996). In Membrane Structure and Dynamics, ed. KM Merz, B Roux, pp. 127-44. Boston: Birkhäuser.
  78. White SH, Wimley WC. (1994). Curr. Opin. Struct. Biol. 4: 79-86.
  79. White SH, Wimley WC. (1998). Biochim. Biophys. Acta 1376: 339-52.
  80. White SH, Wimley WC. (1999). Annu. Rev. Biophys. Biomol. Struc. 28: 319-65.
  81. Wiener MC, White SH. (1992). Biophys. J. 61: 434-47.
  82. Wild UP, Halic M, Sinning I, Beckmann R. (2004). Nat. Struct. Mol. Biol. 11: 1049-53.
  83. Wimley WC, Creamer TP, White SH. (1996). Biochemistry 35: 5109-24.
  84. Wimley WC, Gawrisch K, Creamer TP, White SH. (1996). Proc. Natl. Acad. Sci. USA 93: 2985-90.
  85. Wimley WC, Hristova K, Ladokhin AS, Silvestro L, Axelsen PH, White SH. (1998). J. Mol. Biol. 277: 1091-110.
  86. Wimley WC, White SH. (1996). Nat. Struct. Biol. 3: 842-48.
  87. Wimley WC, White SH. (2000). Biochemistry 39: 4432-42.
  88. Yau W-M, Wimley WC, Gawrisch K, White SH. (1998). Biochemistry 37: 14713-18.
  89. Zhou FX, Cocco MJ, Russ WP, Brunger AT, Engelman DM. (2000). Nat. Struct. Biol. 7: 154-60.
  90. Zhou FX, Merianos HJ, Brunger AT, Engelman DM. (2001). Proc. Natl. Acad. Sci. USA 98: 2250-55.