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Disulfide Bond Formation Promotes the cis- and trans-Dimerization of the E-cadherin-derived First Repeat*

Open AccessPublished:February 20, 2002DOI:https://doi.org/10.1074/jbc.M200916200
      Cadherin is a cell adhesion molecule crucial for epithelial and endothelial cell monolayer integrity. The previously solved x-ray crystallographic structure of the E-CAD12cis-dimer displayed an unpaired Cys9, which protruded away from the Cys9 on the other protomer. To investigate the possible biological function of Cys9 within the first repeat (the E-cadherin-derived N-terminal repeat), E-CAD1 was overexpressed and secreted into the periplasmic space ofEscherichia coli cells. Recombinant E-CAD1 produced a mixed monomer and dimer in an equilibrium fashion. The dimer was linked by a disulfide through Cys9 pairing. Analysis by high pressure liquid chromatography and electron microscopy suggested the existence of oligomeric complexes. Mutation at Trp2 appears to indicate that these oligomeric complexes trans-dimerize. Interestingly, mutation of Cys9 affected not only thecis-dimerization, but also thetrans-oligomerization of E-CAD1. Accordingly, it is plausible that, under oxidative stress, the homophilic interactions of E-cadherin through E-CAD1 may be promoted and stabilized by this disulfide bond.
      MALDI-TOF
      matrix-assisted laser desorption ionization time-of-flight
      HPLC
      high pressure liquid chromatography
      DTT
      dithiothreitol
      SBMD
      stochastic boundary molecular dynamics
      E-cadherin, a Ca2+-dependent member of the cadherin family of cell adhesion molecules, is crucial in providing cell polarity, tightness, and integrity of the intercellular junctions (
      • Takeichi M.
      ,
      • Larue L.
      • Ohsugi M.
      • Hirchenhain J.
      • Kemler R.
      ). In most cancerous tissues, nonfunctional E-cadherin leads to the disturbance of the integrity of the intercellular junctions and consequentially promotes higher mobility and invasiveness of the cancer cells (
      • Beavon I.R.
      ,
      • Perl A.K.
      • Wilgenbus P.
      • Dahl U.
      • Semb H.
      • Christofori G.
      ,
      • Masur K.
      • Lang K.
      • Niggemann B.
      • Zanker K.S.
      • Entschladen F.
      ).
      In recent years, considerable information has been gathered on the adhesion mechanism of classical cadherins from both structural and functional studies. The modular architecture of classical cadherins is characterized by the five repeats in the extracellular domain (
      • Pokutta S.
      • Herrenknecht K.
      • Kemler R.
      • Engel J.
      ). The N-terminal of the extracellular domain is believed to be critical for homophilic cadherin interactions. The structure of the N-terminal fragment derived from the first repeat of the neuronal N-cadherin (N-CAD1) (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ) and the epithelial E-cadherin (E-CAD1) (
      • Tong K.I.
      • Yau P.
      • Overduin M.
      • Bagby S.
      • Porumb T.
      • Takeichi M.
      • Ikura M.
      ) displays a resemblance to the structural fold of immunoglobulin.
      Biochemical analysis of the first extracellular repeat E-CAD1 revealed only the presence of monomers (
      • Tong K.I.
      • Yau P.
      • Overduin M.
      • Bagby S.
      • Porumb T.
      • Takeichi M.
      • Ikura M.
      ). The solution structure of this monomer form was indeed determined by nuclear magnetic resonance (
      • Overduin M.
      • Harvey T.S.
      • Bagby S.
      • Tong K.I.
      • Yau P.
      • Takeichi M.
      • Ikura M.
      ,
      • Overduin M.
      • Tong K.I.
      • Kay C.M.
      • Ikura M.
      ). This is in contrast with the crystal lattice structure of N-CAD1 (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ), which contains a unique mixture of two different populations. One population consists of monomers interacting closely to form cis-dimers (parallel), which are stabilized by the exchange between the N-terminal βA-strand and the intercalation of Trp2 into the partnering hydrophobic core. The other population reveals the pairing between antiparallel-orientedcis-dimers, designated as trans-dimers to reflect the possible head-to-head contacts between two cadherin molecules of the apposing cells. Alternating the cis- andtrans-interactions forms a zipper-like structure (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ). Crystallographic analysis of E-CAD12 also showed cis-dimers linked together by calcium ions with no evidence for adhesivetrans-contact (
      • Nagar B.
      • Overduin M.
      • Ikura M.
      • Rini J.M.
      ). Intriguingly, neither cis-nor trans-dimers were observed in the crystal structure of the extended form of N-CAD12, irrespective of the presence of Ca2+ (
      • Tamura K.
      • Shan W.S.
      • Hendrickson W.A.
      • Colman D.R.
      • Shapiro L.
      ). Thus, the interactions of the molecular repeats of E-CAD and N-CAD remain controversial. Nevertheless, it is basically agreed from in vitro and in vivo studies that lateral cis-dimerization of cadherin molecules is a prerequisite step for the trans-adhesive activity (
      • Yap A.S.
      • Brieher W.M.
      • Pruschy M.
      • Gumbiner B.M.
      ,
      • Alattia J.R.
      • Ames J.B.
      • Porumb T.
      • Tong K.I.
      • Heng Y.M.
      • Ottensmeyer P.
      • Kay C.M.
      • Ikura M.
      ,
      • Murase S.
      • Hirano S.
      • Wang X.
      • Kitagawa M.
      • Natori M.
      • Taketani S.
      • Suzuki S.T.
      ). Further mutational studies of the E-cadherin ectodomain fused with the pentamerization domain of cartilage oligomeric matrix protein (E-CADCOMP) seem to indicate that Trp2 docks into an intramolecular hydrophobic pocket, and this process appears to be critical for the trans-interaction event (
      • Tomschy A.
      • Fauser C.
      • Landwehr R.
      • Engel J.
      ,
      • Pertz O.
      • Bozic D.
      • Koch A.W.
      • Fauser C.
      • Brancaccio A.
      • Engel J.
      ).
      At present, no studies have been performed to determine the role of the conserved Cys9 found in the first repeat of the E-cadherin sequence. Intriguingly, this particular cysteine residue is missing in other classical cadherins. As an initial effort toward understanding the contributory role of Cys9 in the homophilic mechanism of E-cadherin and in contrast to previously published works (
      • Tong K.I.
      • Yau P.
      • Overduin M.
      • Bagby S.
      • Porumb T.
      • Takeichi M.
      • Ikura M.
      ,
      • Nagar B.
      • Overduin M.
      • Ikura M.
      • Rini J.M.
      ), we have overexpressed and secreted the recombinant E-CAD1 fragment into the oxidizing environment of the periplasmic space of Escherichia coli cells. Our study shows that E-CAD1 is capable of forming disulfide-bonded cis-dimers, followed by their presumablytrans-dimerization. These findings have important implications for the proposed mechanism for homophilictrans-interactions occurring at the first repeat of E-cadherin (
      • Tomschy A.
      • Fauser C.
      • Landwehr R.
      • Engel J.
      ,
      • Blaschuk O.W.
      • Sullivan R.
      • David S.
      • Pouliot Y.
      ,
      • Nose A.
      • Tsuji K.
      • Takeichi M.
      ,
      • Kitagawa M.
      • Natori M.
      • Murase S.
      • Hirano S.
      • Taketani S.
      • Suzuki S.T.
      ).

      DISCUSSION

      Previously, the structure of N-CAD1 revealed a dimer (
      • Overduin M.
      • Harvey T.S.
      • Bagby S.
      • Tong K.I.
      • Yau P.
      • Takeichi M.
      • Ikura M.
      ) as opposed to the crystallographic structure E-CAD1 (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ), whereas the cooperative binding of Ca2+ enhanced thecis-dimerization of E-CAD12 (
      • Nagar B.
      • Overduin M.
      • Ikura M.
      • Rini J.M.
      ,
      • Pertz O.
      • Bozic D.
      • Koch A.W.
      • Fauser C.
      • Brancaccio A.
      • Engel J.
      ). It was found that the Ca2+-binding interface located between E-CAD1 and E-CAD2 mediates this dimer arrangement (
      • Koch A.W.
      • Bozic D.
      • Pertz O.
      • Engel J.
      ). In this present work, we further show the existence of a novel E-CAD1 dimer linked by a disulfide bond via its Cys9. The same disulfide bond was also formed when we overexpressed the extended E-CAD12 (the second repeat does not contain cysteine).
      I. T. Makagiansar, P. D. Nguyen, and T. J. Siahaan, unpublished data.
      This peculiar phenomenon was not previously observed with N-CAD1, E-CAD1, or E-CAD12. Thus, dimerization of our E-CAD1 appears to be promoted by the disulfide bond, regardless of the presence of Ca2+. The assumption that dimerization can occur with E-CAD12 only because of the Ca2+-binding interface located between the first and second repeats therefore may not hold true.
      It is interesting to note that the x-ray structure of the E-CAD12 dimer displays each thiol group protruding inwardly with the sulfur atoms separated by 14.8 Å, which appears to sterically hinder the formation of the disulfide bond. Our results suggest that, in solution, the βA-region of E-CAD1dm undertakes a less rigid conformation, rendering the thiol group more exposed for pairing to form the disulfide bond. Indeed, our thermodynamic simulation using the coordinates from the x-ray structure of the E-CAD12 dimer demonstrates that a disulfide bond can be optimally formed at a pairing distance of 2.1 Å without the necessity of large-scale changes in the protein. In relation to this finding, it remains unclear what mechanical force causes N-CAD1 to dimerize in the absence of a disulfide (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ) given the fact that N-CAD1 does not contain cysteine (
      • Reid R.A.
      • Hemperly J.J.
      ).
      The absence of disulfide formation in previous E-CAD1 and E-CAD12 structures may arise because of the expression system used to produce these recombinant fragments. The latter were overexpressed in the cytoplasmic compartment of E. coli cells (
      • Tong K.I.
      • Yau P.
      • Overduin M.
      • Bagby S.
      • Porumb T.
      • Takeichi M.
      • Ikura M.
      ,
      • Nagar B.
      • Overduin M.
      • Ikura M.
      • Rini J.M.
      ), a condition that prevents formation of structural disulfide bonds due to its reducing environment (
      • Skerra A.
      • Pluckthun A.
      ,
      • Derman A.I.
      • Beckwith J.
      ). In contrast, our present E-CAD1 was expressed and exported into the E. coli periplasm, the oxidizing environment of which promotes disulfide formation (
      • Reid R.A.
      • Hemperly J.J.
      ,
      • Skerra A.
      • Pluckthun A.
      ). More importantly, the presence of thiol-disulfide oxidoreductases (i.e. DsbA and DsbC) (
      • Wunderlich M.
      • Jaenicke R.
      • Glockshuber R.
      ,
      • Wunderlich M.
      • Glockshuber R.
      ,
      • Jonda S.
      • Huber-Wunderlich M.
      • Glockshuber R.
      • Mossner E.
      ,
      • Zapun A.
      • Missiakas D.
      • Raina S.
      • Creighton T.E.
      ) in the periplasmic compartment may contribute significantly to the favorable arrangement and pairing of the neighboring thiol side chain from each E-CAD1 monomer to promote dimerization.
      Intriguingly, the E-CAD1 dimer can, in fact, form oligomeric complexes. MALDI-TOF analysis of E-CAD1m/E-CAD1dm using the less acidic matrix condition for noncovalent protein-protein studies revealed masses at m/z 66,585 and 133,257, which are calculated to correspond to a tetramer (66,585/16,910 = 3.94) and an octamer (133,257/16,910 = 7.90), respectively. No higher masses were observed, which was due presumably to the instability of the higher oligomeric complexes under this condition. Indeed, when the dimer band in peak 1 was cross-linked, a smeared band with apparent molecular masses in the range of 180–210 kDa was observed. Furthermore, light scattering of peak 1 revealed a mixture of species of different masses, with a molecular mass distribution ranging from 140 to 300 kDa. This observation strongly suggests that this peak is polydisperse and contains different oligomeric complexes. The E-CAD1oc dissociation rate constant (kd) was calculated to be 23 × 10−4 s−1 at 22 °C, which indicates that the complexes may not be too stable.
      Additionally, the oligomerization of E-CAD1 appears to be stabilized by the formation of the disulfide bond. In the presence of the reducing agent DTT, peak 1 was eliminated. Similarly, when Cys9 was replaced with alanine (E-CAD1C9A), the same chromatographic profile as obtained with DTT was produced.
      Electron microscopic images of E-CAD1m/E-CAD1dmshowed particles of various sizes, consisting mainly of single or longer bead-like molecules. However, these particles did not clearly show whether they adopted a cis-orientation (parallel) or atrans-orientation (antiparallel). Studies with E-CADCOMP demonstrated that docking of the second tryptophan residue (Trp2) into the hydrophobic cavity (79HAV81) of the same molecule is important for adhesive trans-interactions (
      • Pertz O.
      • Bozic D.
      • Koch A.W.
      • Fauser C.
      • Brancaccio A.
      • Engel J.
      ,
      • Koch A.W.
      • Bozic D.
      • Pertz O.
      • Engel J.
      ). Mutation of Trp2 in E-CAD12COMP abolished the trans-dimer interaction, but not the cis-dimerization. In our case, substituting Trp2 with alanine (E-CAD1W2A) led to the partial reduction of oligomeric complexes without affecting dimerization. Full abrogation of the E-CAD1W2A oligomeric complexes was not observed, probably due to other intermolecular forces influencing the trans-dimerization. Among such factors is the presence of the disulfide bond, which may substantially alleviate the mutational effect of tryptophan. Taken together, however, our data support the presumption that the oligomeric complex of E-CAD1 adopts an antiparallel orientation.
      From the x-ray crystallography of N-CAD1, a dimer size of 38 × 38 Å was obtained (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ), whereas the size for the E-CAD1 dimer derived from the x-ray crystallography of E-CAD12 was 46 × 48 Å (
      • Nagar B.
      • Overduin M.
      • Ikura M.
      • Rini J.M.
      ). Given the condition under which two E-CAD1 dimers trans-interact with one another, then total sizes of 2 × 38 = 76 Å for N-CAD1 and 2 × 46 = 92 Å (width) by 2 × 48 = 96 Å (length) for E-CAD1 are predicted. Consequently, a particle size of 80 × 102 Å derived from the electron microscopy is in close agreement with two dimers trans-interacting (hence, a 67-kDa tetramer). The E-CAD1 trans-dimer has a smaller width than that of the predicted E-CAD1 trans-dimer derived from E-CAD12 crystallography, but closer to that of N-CAD1. We speculate that the smaller width may be due to the presence of the disulfide, which brings the interfacial surfaces of the two monomers closer to one another. It is to be noted, however, that neither the monomeric nor the dimeric E-CAD1 particle was detected, probably due to its minute size. Meanwhile, a particle size of 254 × 118 Å appears to correspond to six dimers trans-interacting (hence, a 200-kDa dodecamer). Much longer bead-like structures of different lengths were also detected by electron microscopy. Because of their nonlinear elongated shapes, measurement of these dimensions could not be carried out accurately. Nevertheless, by estimating the tetrameric repeats contained in these long bead-like particles, an apparent molecular mass range of ∼240–340 kDa was deduced.
      Thus, the overall molecular mass range observed by electron microscopy is consistent with both the MALDI-TOF results and the molecular mass distribution of peak 1 obtained by the HPLC/dynamic light scattering method. This information provides strong evidence for the existence oftrans-oligomeric complexes of E-CAD1.
      Despite the fact that molecular force measurements between antiparallel oriented monolayers of the Xenopus C-cadherin extracellular repeats may indicate that cadherin binding involves distinct multiple repeats for trans-interactions (
      • Corada M.
      • Liao F.
      • Lindgren M.
      • Lampugnani M.G.
      • Breviario F.
      • Frank R.
      • Muller W.A.
      • Hicklin D.J.
      • Bohlen P.
      • Dejana E.
      ,
      • Chappuis-Flament S.
      • Wong E.
      • Hicks L.D.
      • Kay C.M.
      • Gumbiner B.M.
      ,
      • Sivasankar S.
      • Gumbiner B.
      • Leckband D.
      ), several lines of evidence, including our present data, suggest the possible homophilic cadherin trans-interaction taking place at the N terminus to form a zipper-like molecule (
      • Shapiro L.
      • Fannon A.M.
      • Kwong P.D.
      • Thompson A.
      • Lehmann M.S.
      • Grubel G.
      • Legrand J.F.
      • Als-Nielsen J.
      • Colman D.R.
      • Hendrickson W.A.
      ,
      • Nose A.
      • Tsuji K.
      • Takeichi M.
      ,
      • Kitagawa M.
      • Natori M.
      • Murase S.
      • Hirano S.
      • Taketani S.
      • Suzuki S.T.
      ,
      • Takeda H.
      • Shimoyama Y.
      • Nagafuchi A.
      • Hirohashi S.
      ,
      • Makagiansar I.T.
      • Hu M.
      • Avery Y.
      • Audus K.L.
      • Siahaan T.J.
      ). Indeed, a cryoelectron microscopy study by rapid-freeze deep-etching of the adherens junction in retinal pigment epithelium showed that the extracellular domain of E-cadherin forms zipper-like molecules within the intercellular space (
      • Miyaguchi K.
      ). These molecules consist of combined rods and globules, the rod being the dimerized E-cadherin and the globule being the enlarged trans-contact regions between the first or second repeat of E-cadherin.
      The biological implication of this disulfide bond suggests an alternative means of regulation of E-cadherin by a redox mechanism. Redox modulation at specific cysteine residues was observed to influence the function of hemoglobin (
      • Jia L.
      • Bonaventura C.
      • Bonaventura J.
      • Stamler J.S.
      ), theN-methyl-d-aspartate receptor (
      • Kim W.K.
      • Choi Y.B.
      • Rayudu P.V.
      • Das P.
      • Asaad W.
      • Arnelle D.R.
      • Stamler J.S.
      • Lipton S.A.
      ), and integrin αIIbβ3 (
      • Yan B.
      • Smith J.W.
      ). Under oxidative stress, it may be possible that a redox switch from an unpaired Cys9 to a paired cysteine affects the conformation of the extracellular domains, stabilizing further E-cadherin cell-mediated adhesion. In retrospect, more work will be required to confirm this possibility.
      In summary, our data provide clear evidence supporting the idea that the first repeat is involved in the trans-dimerization of E-cadherin. It is further demonstrated that a disulfide bond not only appears to be a decisive factor in promoting the dimerization of the first repeat, but also acts as a stabilizing force toward formation oftrans-dimeric complexes of E-CAD1.

      Acknowledgments

      We are grateful to Dr. David Rimm for providing the full-length human E-cadherin cDNA. We also thank Ewa Folta-Stogniew (Yale University) and Dr. Yongbo Hu for their technical assistance and Nancy Harmony for reviewing this manuscript.

      REFERENCES

        • Takeichi M.
        Science. 1991; 251: 1451-1455
        • Larue L.
        • Ohsugi M.
        • Hirchenhain J.
        • Kemler R.
        Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 8263-8267
        • Beavon I.R.
        Eur. J. Cancer. 2000; 36: 1607-1620
        • Perl A.K.
        • Wilgenbus P.
        • Dahl U.
        • Semb H.
        • Christofori G.
        Nature. 1998; 392: 190-193
        • Masur K.
        • Lang K.
        • Niggemann B.
        • Zanker K.S.
        • Entschladen F.
        Mol. Biol. Cell. 2001; 12: 1973-1982
        • Pokutta S.
        • Herrenknecht K.
        • Kemler R.
        • Engel J.
        Eur. J. Biochem. 1994; 223: 1019-1026
        • Shapiro L.
        • Fannon A.M.
        • Kwong P.D.
        • Thompson A.
        • Lehmann M.S.
        • Grubel G.
        • Legrand J.F.
        • Als-Nielsen J.
        • Colman D.R.
        • Hendrickson W.A.
        Nature. 1995; 374: 327-337
        • Tong K.I.
        • Yau P.
        • Overduin M.
        • Bagby S.
        • Porumb T.
        • Takeichi M.
        • Ikura M.
        FEBS Lett. 1994; 352: 318-322
        • Overduin M.
        • Harvey T.S.
        • Bagby S.
        • Tong K.I.
        • Yau P.
        • Takeichi M.
        • Ikura M.
        Science. 1995; 267: 386-389
        • Overduin M.
        • Tong K.I.
        • Kay C.M.
        • Ikura M.
        J. Biomol. NMR. 1996; 7: 173-189
        • Nagar B.
        • Overduin M.
        • Ikura M.
        • Rini J.M.
        Nature. 1996; 380: 360-364
        • Tamura K.
        • Shan W.S.
        • Hendrickson W.A.
        • Colman D.R.
        • Shapiro L.
        Neuron. 1998; 20: 1153-1163
        • Yap A.S.
        • Brieher W.M.
        • Pruschy M.
        • Gumbiner B.M.
        Curr. Biol. 1997; 7: 308-315
        • Alattia J.R.
        • Ames J.B.
        • Porumb T.
        • Tong K.I.
        • Heng Y.M.
        • Ottensmeyer P.
        • Kay C.M.
        • Ikura M.
        FEBS Lett. 1997; 417: 405-408
        • Murase S.
        • Hirano S.
        • Wang X.
        • Kitagawa M.
        • Natori M.
        • Taketani S.
        • Suzuki S.T.
        Biochem. Biophys. Res. Commun. 2000; 276: 1191-1198
        • Tomschy A.
        • Fauser C.
        • Landwehr R.
        • Engel J.
        EMBO J. 1996; 15: 3507-3514
        • Pertz O.
        • Bozic D.
        • Koch A.W.
        • Fauser C.
        • Brancaccio A.
        • Engel J.
        EMBO J. 1999; 18: 1738-1747
        • Blaschuk O.W.
        • Sullivan R.
        • David S.
        • Pouliot Y.
        Dev. Biol. 1990; 139: 227-229
        • Nose A.
        • Tsuji K.
        • Takeichi M.
        Cell. 1990; 61: 147-155
        • Kitagawa M.
        • Natori M.
        • Murase S.
        • Hirano S.
        • Taketani S.
        • Suzuki S.T.
        Biochem. Biophys. Res. Commun. 2000; 271: 358-363
        • Mach H.
        • Volkin D.B.
        • Burke C.J.
        • Middaugh C.R.
        Methods Mol. Biol. 1995; 40: 91-114
        • Farmer T.B.
        • Caprioli R.M.
        J. Mass Spectrom. 1998; 33: 697-704
        • Woods A.S.
        • Huestis M.A.
        J. Am. Soc. Mass Spectrom. 2001; 12: 88-96
        • Jorgensen W.L.
        • Chandrasekhar J.
        • Madura J.D.
        • Impey R.W.
        • Klein M.L.
        J. Chem. Phys. 1983; 79: 926-935
        • Brooks III, C.L.
        • Karplus M.
        • Pettitt B.M.
        Proteins: A Theoretical Perspective of Dynamics, Structure, and Thermodynamics. 2nd Ed. John Wiley & Sons, Inc., New York1988
        • Brooks B.R.
        • Bruccoleri R.
        • Olafson B.
        • States D.
        • Swaminathan S.
        • Karplus M.
        J. Comp. Chem. 1983; 4: 187-217
        • Koch A.W.
        • Bozic D.
        • Pertz O.
        • Engel J.
        Curr. Opin. Struct. Biol. 1999; 9: 275-281
        • Reid R.A.
        • Hemperly J.J.
        Nucleic Acids Res. 1990; 18: 5896
        • Skerra A.
        • Pluckthun A.
        Science. 1988; 240: 1038-1041
        • Derman A.I.
        • Beckwith J.
        J. Bacteriol. 1991; 173: 7719-7722
        • Wunderlich M.
        • Jaenicke R.
        • Glockshuber R.
        J. Mol. Biol. 1993; 233: 559-566
        • Wunderlich M.
        • Glockshuber R.
        J. Biol. Chem. 1993; 268: 24547-24550
        • Jonda S.
        • Huber-Wunderlich M.
        • Glockshuber R.
        • Mossner E.
        EMBO J. 1999; 18: 3271-3281
        • Zapun A.
        • Missiakas D.
        • Raina S.
        • Creighton T.E.
        Biochemistry. 1995; 34: 5075-5089
        • Corada M.
        • Liao F.
        • Lindgren M.
        • Lampugnani M.G.
        • Breviario F.
        • Frank R.
        • Muller W.A.
        • Hicklin D.J.
        • Bohlen P.
        • Dejana E.
        Blood. 2001; 97: 1679-1684
        • Chappuis-Flament S.
        • Wong E.
        • Hicks L.D.
        • Kay C.M.
        • Gumbiner B.M.
        J. Cell Biol. 2001; 154: 231-243
        • Sivasankar S.
        • Gumbiner B.
        • Leckband D.
        Biophys. J. 2001; 80: 1758-1768
        • Yan B.
        • Smith J.W.
        J. Biol. Chem. 2000; 275: 39964-39972
        • Takeda H.
        • Shimoyama Y.
        • Nagafuchi A.
        • Hirohashi S.
        Nat. Struct. Biol. 1999; 6: 310-312
        • Makagiansar I.T.
        • Hu M.
        • Avery Y.
        • Audus K.L.
        • Siahaan T.J.
        Pharm. Res. 2001; 18: 446-453
        • Miyaguchi K.
        J. Struct. Biol. 2000; 132: 169-178
        • Jia L.
        • Bonaventura C.
        • Bonaventura J.
        • Stamler J.S.
        Nature. 1996; 380: 221-226
        • Kim W.K.
        • Choi Y.B.
        • Rayudu P.V.
        • Das P.
        • Asaad W.
        • Arnelle D.R.
        • Stamler J.S.
        • Lipton S.A.
        Neuron. 1999; 24: 461-469