Critical Structural and Functional Roles for the N-Terminal Insertion Sequence in Surfactant Protein B Analogs
et al. (2010) Critical Structural and Functional Roles for the N-Terminal Insertion
Sequence in Surfactant Protein B Analogs. PLoS ONE 5(1): e8672. doi:10.1371/journal.pone.0008672
Critical Structural and Functional Roles for the N-Terminal Insertion Sequence in Surfactant Protein B Analogs
Frans J. Walther 0
Alan J. Waring 0
Jose M. Hernandez-Juviel 0
Larry M. Gordon 0
Zhengdong Wang 0
Chun-Ling Jung 0
Piotr Ruchala 0
Andrew P. Clark 0
Wesley M. Smith 0
Shantanu Sharma 0
Robert H. 0
Rory Edward Morty, University of Giessen Lung Center, Germany
0 1 Los Angeles Biomedical Research Institute at Harbor, University of California Los Angeles Medical Center, Torrance, California, United States of America, 2 Department of Pediatrics, Leiden University Medical Center , Leiden , The Netherlands , 3 Department of Medicine, University of California Los Angeles , Los Angeles , California, United States of America, 4 Department of Pediatrics, University of Rochester , Rochester , New York, United States of America, 5 Department of Chemistry and Center for Macromolecular Modeling and Materials Design, California State Polytechnic University , Pomona , California, United States of America, 6 Department of Environmental Medicine, University of Rochester , Rochester, New York , United States of America
Background: Surfactant protein B (SP-B; 79 residues) belongs to the saposin protein superfamily, and plays functional roles in lung surfactant. The disulfide cross-linked, N- and C-terminal domains of SP-B have been theoretically predicted to fold as charged, amphipathic helices, suggesting their participation in surfactant activities. Earlier structural studies with Mini-B, a disulfide-linked construct based on the N- and C-terminal regions of SP-B (i.e., ,residues 8-25 and 63-78), confirmed that these neighboring domains are helical; moreover, Mini-B retains critical in vitro and in vivo surfactant functions of the native protein. Here, we perform similar analyses on a Super Mini-B construct that has native SP-B residues (1-7) attached to the N-terminus of Mini-B, to test whether the N-terminal sequence is also involved in surfactant activity. Methodology/Results: FTIR spectra of Mini-B and Super Mini-B in either lipids or lipid-mimics indicated that these peptides share similar conformations, with primary a-helix and secondary b-sheet and loop-turns. Gel electrophoresis demonstrated that Super Mini-B was dimeric in SDS detergent-polyacrylamide, while Mini-B was monomeric. Surface plasmon resonance (SPR), predictive aggregation algorithms, and molecular dynamics (MD) and docking simulations further suggested a preliminary model for dimeric Super Mini-B, in which monomers self-associate to form a dimer peptide with a ''saposin-like'' fold. Similar to native SP-B, both Mini-B and Super Mini-B exhibit in vitro activity with spread films showing near-zero minimum surface tension during cycling using captive bubble surfactometry. In vivo, Super Mini-B demonstrates oxygenation and dynamic compliance that are greater than Mini-B and compare favorably to full-length SP-B. Conclusion: Super Mini-B shows enhanced surfactant activity, probably due to the self-assembly of monomer peptide into dimer Super Mini-B that mimics the functions and putative structure of native SP-B.
Funding: The authors gratefully acknowledge the financial support of the National Institutes of Health through grants HL-092158, ES-015330, HL-080775, and
HL-094641. NIH had no role in the design and conduct of the study, in the collection, analysis, and interpretation of the data, and in the preparation, review, or
approval of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Lung surfactant is a complex mixture of lipids (mostly
phospholipids) and proteins that is required for normal breathing,
due to its ability to reduce alveolar surface tension to very low
values. Surfactant is synthesized and secreted into the alveolar
fluid by type II cells, and consists of approximately 80%
phospholipids, 10% neutral lipids and 10% proteins [1,2]. Despite
dipalmitoyl phosphatidylcholine and phosphatidylglycerol
constituting its main phospholipid components, the biophysical activity
of surfactant in the lung largely depends on the presence of the
hydrophobic surfactant protein B (SP-B), and to a lesser degree on
the extremely hydrophobic surfactant protein C (SP-C) .
Hereditary SP-B deficiency is lethal in humans , while
mutations in the SP-C gene may cause interstitial lung disease
and increase susceptibility to infection . Surfactant therapy
using bovine or porcine lung extracts surfactant extracts, which
contain only polar lipids and native SP-B and SP-C, has greatly
improved the therapeutic outcomes of neonates with respiratory
distress (NRDS). Exogenous surfactant replacement therapies are
currently being extended to pediatric and adult patients with direct
pulmonary forms of clinical acute lung injury (ALI) and the acute
respiratory distress syndrome (ARDS) . An important goal
of surfactant researchers is to replace animal-derived therapies
with fully synthetic preparations based on SP-B and SP-C,
produced by recombinant technology or peptide synthesis, and
reconstituted with selected synthetic lipids (SL) .
SP-B is a small (79 amino acids; monomer MW of 8.7 kDa),
lipid-associating protein that is found in the mammalian lung as a
covalently linked homodimer, through a disulfide bridge at
positions Cys-51, Cys-519. Each SP-B monomer contains three
intramolecular disulfide bridges (i.e., Cys-8 to Cys-77, Cys-11 to
Cys-71 and Cys-35 to Cys-46) . SP-B belongs to the saposin
protein superfamily, and earlier X-ray crystallographic or two
dimensional nuclear magnetic resonance (2D-NMR) spectroscopic
studies on saposins other than SP-B showed that the characteristic
saposin fold consists of 45 a-helical domains [i.e., ,residues
1020 (N-terminal helix), 2538, 4152, 5963 and 6875
(Cterminal helix)] joined together by 23 intramolecular disulfide
links . The helical bundle for saposins is folded into two leaves,
with one leaf having a-helices 1 and 45 and the second leaf
composed of a-helices 2 and 3, with flexible hinges between helices
1 and 2 and also between helices 3 and 45. For the saposins
NKlysin, granulysin or saposin C in aqueous environments, the
protein folds in a closed tertiary conformation; here, the two leaves
are in close contact such that the amphipathic a-helices with
hydrophilic (charged, neutral) residues face the solvent, while the
hydrophobic side chains form a core stabilized by intramolecular
disulfide bonds . On the other hand, aqueous dimeric saposin
B  or saposin C bound to submicellar SDS detergent 
(Fig. 1A, B) show opened conformations, in which the leaves of the
V are now far apart having expanded at the flexible joints. The
respective open conformations allow saposin-B to form
noncovalent dimers interacting through their exposed hydrophobic cores
, while saposin C unmasks its hydrophobic core to bind the
fatty acyl chains of SDS detergent  (Fig. 1B). These
observations support an early proposal  that increases in the
hydrophobicity of the saposins environment (e.g., binding to
membranes or lipids) may generally produce a greater splay
between the protein leaves, thereby exposing more hydrophobic
Although experimental analyses have not yet determined the
3D-structure of full-length SP-B , the native protein will
probably share the above saposin fold for several reasons. First, the
primary sequence of SP-B is highly homologous with those of
other known saposins [18,21,23]. Second, the intrachain
disulfidelinkage pattern observed with SP-B  and saposins has been
conserved for ,300 million years . Lastly, circular dichroism
(CD) and Fourier transform infrared (FTIR) spectroscopy of native
SP-B in membrane mimics indicated high a-helix levels similar to
that of other saposins [23,2527]. Molecular models of
homodimeric SP-B, based on templating the primary sequence of SP-B
onto the known 3D-structures of NK-lysin, suggested that SP-B
may assume the closed and/or opened saposin conformations
when interacting with lipid monolayers or bilayers [4,5,28,29]. In
the open conformation, the exposed amphipathic helices of SP-B
would bind to lipid by inserting its hydrophobic residues to
interact with fatty acyl chains, while charged and neutral residues
would associate with the more polar lipid headgroup region [5,30].
Consistent with these proposed SP-B binding models is an early
orientation-dependent FTIR study of native SP-B in lipid
membranes indicating that a fraction of the helices lie parallel to
the lipid surface, while another fraction is slightly embedded in the
bilayers, parallel to the fatty acyl chains . It is of particular
interest that the above SP-B models predict disk-like structures
containing disulfide-linked charged amphipathic helices (i.e.,
Nand C-terminal domains) which may promote surfactant activity
Previous structural and functional studies with synthetic
peptides representing the N- and C-terminal regions of SP-B
further support the hypothesis that these charged amphipathic
helices participate in surfactant activities. SP-B and a synthetic
peptide based on the N-terminal domain of SP-B (i.e., SP-B(125);
residues 125) each increase the collapse pressure of lipid
monolayers containing palmitic acid. The cationic N-terminus of
SP-B may here interact with anionic lipids to remove the driving
force for lipid squeeze-out from the surface film [31,32]. In
additional studies, SP-B(125) and native SP-B each induced a
coexistence of buckled and flat monolayers when added to
surfactant lipids, promoted a low surface tension and increased
respreading of the surfactant monolayer . The above in vitro
surfactant activities of SP-B(125) are also correlated with the
improved oxygenation and lung compliance noted for this peptide
in surfactant-deficient animal models . Extensive domain
mapping experiments have recently confirmed that the N-terminal
helical-domain is required for the fusogenic, lytic and surface
activities of SP-B . Because physical studies indicated high
ahelical levels for N-terminal SP-B peptides in lipids or
membranemimics [30,3841], the N-terminal domain in native SP-B may
participate in surfactant actions as a charged amphipathic a-helix.
Interestingly, several non-natural analogs of SP-B(125), which are
a-helical in lipid environments, also exhibit in vitro surfactant
activities . A positively-charged helical C-terminus of SP-B
may similarly be involved in lung function, as synthetic peptides
representing the C-terminal domain adopt a-helical conformations
[43,44] and promote in vitro [43,4547] and in vivo [46,47]
surfactant activities mimicking those of the native protein.
Because the N- and C-terminal domains are the principal
interaction sites for SP-B with surfactant lipids, earlier experiments
were performed with an artificial 34-residue construct (i.e.,
MiniB) containing these motifs. Mini-B (i.e., MB) incorporates residues
8-25 and 63-78 of native SP-B as a single linear peptide, and was
designed to join the critical N- and C-terminal amphipathic
helixes with a b-sheet-loop domain . MB folds into a
helixhairpin structure when oxidized, and is stabilized by disulfide
connectivity between Cys-8 and Cys-40 and Cys-11 and Cys-34
(residue numbers refer to the MB sequence in Fig. 2B).
Conventional 12C-FTIR spectroscopy indicated that oxidized
MB has elevated a-helical levels in membrane mimics , and it
is likely that the N- and C-terminal regions will be helical in MB
similar to that observed for peptide fragments based on these
domains (see above). Indeed, residue-specific analyses using either
isotope-enhanced 13C-FTIR  or 2D-NMR  spectroscopy
confirmed that MB shares the same three-dimensional saposin fold
(see PDB accession codes: 1SSZ and 2DWF; www.rcsb.org) as the
predicted full-length SP-B protein in the N- and C-terminal
regions [4,28]. MB in model surfactant lipid mixtures showed
marked in vitro activity, with spread films exhibiting near-zero
minimum surface tension during cycling using captive bubble
surfactometry . Using in vivo experiments, surfactant-deficient,
ventilated rats also demonstrated a rapid recovery of oxygenation
(PaO2) and dynamic compliance after rescue surfactant treatment
with MB and surfactant lipids that was comparable to that of
porcine SP-B and lipids [5,48]. High surfactant activity was
additionally determined recently for MB with DEPN-8 (i.e., a
novel diether phosphonolipid) employing various in vitro techniques
. MB had greatly increased adsorption compared to DEPN-8
alone, while MB and DEPN-8 rapidly reached minimum surface
tensions in either pulsating bubble or captive bubble
surfactometry. In the context of developing fully-synthetic lipid/peptide
preparations for treating surfactant deficiencies, it is noteworthy
that MB and DEPN-8 mixtures were fully resistant to degradation
by phospholipase A2 . In the present paper, we perform
similar structural and functional analyses on Super Mini-B or
S-MB, an artificial construct that has native SP-B residues
(residues 1-7; Phe-Pro-Ile-Pro-Leu-Pro-Tyr) attached to the
N-terminus of Mini-B (Fig. 2A), to test whether the N-terminal
insertion sequence is also involved in surfactant activity. This
putative lipid insertion region of SP-B includes a
X-Pro-X-Pro-XPro motif and residues (79; Tyr-Cys-Trp), and may influence
the ability of SP-B to associate with itself [40,50] or lipids
[30,39,51,52]. Consistent with an important function for the
N-terminal insertion domain is the finding that the surface activity
was reduced for N-terminal peptides with replacements at either
tryptophan 9 or prolines 2, 4 and 6 [38,52]. Moreover, a
nonnatural analog of SP-B(125) with a hydrophobic, helical region
substituting for the N-terminal insertion region exhibited more
surface activity than the native peptide . The roles that the
N-terminal insertion sequence may play in SP-B structure and
function were here investigated by comparing the properties of
MB and S-MB using a suite of in vitro and in vivo assays.
Synthesis of Mini-B (MB), Super Mini-B (S-MB) and
MB (34 amino acid sequence:
NH2-CWLCRALIKRIQAMIPKGGRMLPQLVCRLVLRCSCOOH; see Fig. 2B), S-MB
(41 amino acid sequence:
NH2-FPIPLPYCWLCRALIKRIQAMIPKGGRMLPQLVCRLVLRCS-COOH; see Fig. 2A) and
SPB(18) [8 amino acid sequence: NH2-FPIPLPYC-CONH2] were
prepared with either a ABI 431A solid phase peptide synthesizer
(Applied Biosystems, Foster City, CA) configured for FastMocTM
chemistry , a Symphony Multiple Peptide Synthesizer (Protein
Technologies, Tucson, AZ) using standard Fmoc synthesis, or a
Liberty Microwave Peptide Synthesizer (CEM Corp., Matthews,
NC) configured for standard Fmoc synthesis. A low substitution
(0.3 mmole/gm) pre-derivatized Fmoc-serine (tBu) Wang resin
(NovaBiochem, San Diego, CA) or H-Ser(OtBu)-HMPB Nova
PEG resin (NovaBiochem, San Diego, CA) were used to minimize
the formation of truncated sequences with the MB and S-MB
peptide, while a Rink Amide MBHA resin (NovaBiochem, San
Diego, CA) was employed for synthesis of the SP-B(18) peptide.
All residues were double-coupled to insure optimal yield .
After synthesis of the respective linear sequences, peptides were
cleaved from the resin and deprotected using a mixture of 0.75 gm
phenol, 0.25 ml ethanedithiol, 0.5 ml of thioanisole, 0.5 ml of
deionized water and 10 ml trifluoroacetic acid per gram of resin
initially chilled to 5uC, and then allowed to come to 25uC with
continuous stirring over a period of 2 h to insure complete peptide
deprotection . Crude peptides were removed from the resin by
vacuum-assisted filtration, and by washing on a medium porosity
sintered glass filter with trifluoroacetic acid and dichloromethane
to maximize yield. Filtered crude peptides were precipitated in ice
cold tertiary butyl ether, and separated by centrifugation at
20006g for 10 min (23 cycles of ether-precipitation and
centrifugation were used to minimize cleavage-deprotection
byproducts). Reduced crude peptides from ether-precipitation
were verified for molecular mass by MALDI-TOF spectroscopy,
dissolved in trifluoroethanol (TFE):10 mM HCl (1:1, v:v), freeze
dried, and purified by preparative HPLC . Final folding of
HPLC-purified peptides was facilitated by air-oxidation for at least
48 h at 25uC in TFE and 10 mM ammonium bicarbonate buffer
(4:6, v:v) at pH 8.0 . Final oxidized MB and S-MB were
repurified by reverse phase HPLC, verified in molecular mass via
MALDI-TOF, and disulfide connectivity was confirmed by mass
spectroscopy of enzyme-digested fragments (trypsin and
Sodium Dodecyl Sulfate-Polyacrylamide Gel
The S-MB and MB peptides were characterized in a detergent
environment using sodium dodecyl sulfate-polyacrylamide gel
(SDS-PAGE) electrophoresis . Purified MB or S-MB
samples were eluted from reverse-phase HPLC as indicated
above. Without either heating or reducing these peptide samples,
PAGE with SDS was carried out by dissolving dried peptide (4 mg)
into 8 ml of buffer (50 mM MOPS, 50 mM Tris, pH 7.7, 0.1%
SDS, 1 mM EDTA; NovexTM 2X buffer from Invitrogen,
Carlsbad, CA). PAGE is performed by then applying the dissolved
peptide aliquots to precast 16% acrylamide gels (NovexTM Gels,
Invitrogen, Carlsbad, CA), which are used because of their
reproducibility and minimal sample requirements. Homogenous
16% polyacrylamide gels, which are 4365060.4 mm and precast
on a 0.175-mm-thick polyester support, are used not only to
optimize the resolution in the low molecular weight range, but
also to run the two peptides in the same gel under identical
conditions. The respective S-MB and MB bands were stained
using Coomassie blue and silver to enhance the sample contrast
as described previously , and bands corresponding to
monomeric and oligomeric peptide were identified using tracking
molecular weight markers (MW range 2.516.9 kDa, Invitrogen,
Surface Plasmon Resonance (SPR) Measurements
The respective binding affinities of S-MB and MB, both to
themselves (self-association) and to each other (cross-association),
were measured with surface plasmon resonance (SPR)
spectroscopy using a Biacore 3000 system (Biacore, Uppsala, Sweden)
[16,59]. S-MB and MB, each with a cysteine added to its
Nterminus, were chemically linked to a thiol Biacore chip
(BR-100014, research grade, containing a carboxymethylated dextran
matrix covalently attached to a gold film) . Solutions of
peptide in HBS-EP buffer (i.e., 10 mM Hepes, pH 7.4, 150 mM
NaCl, 3 mM EDTA, 0.005% surfactant P20) were then flowed
over the chip-linked peptide at a flow rate of 50 ml/min to
determine binding affinity at 37uC. Assessment of peptide binding
was determined with sensorgrams, in which the arbitrary response
units (RU) were plotted as a function of time. Binding associated
with control medium containing no peptide was subtracted from
final affinity curves, and mean on and off rate constants (kon
and koff) and the dissociation equilibrium constant (KD = koff/
kon) were calculated using BIAevaluation Software Version 4.1
based on curve fitting form measurements at 1 mg peptide/ml
buffer concentration. The sensor surface was regenerated using
10 mM HCl between sample injections.
Prediction of Aggregation-Forming Domains in Peptides
MB and S-MB were each analyzed with PASTA [61,62] and
AGGRESCAN [63,64] to determine those peptide regions most
likely to form b-sheet, particularly when exposed to polar
environments such as in the aqueous buffer or at the lipid-water
interface [59,65]. The PASTA algorithm systematically calculates
the relative energies of the various pairing arrangements by
calculating a pair-wise energy function for residues facing one
another within a b-sheet. With a database of known 3D-native
structures, PASTA computes two different propensity sets depending
on the directionality (i.e., parallel or antiparallel b-sheets) of the
neighboring strands. PASTA assigns relative energies to specific
bpairings of two sequence stretches of the same length, and assumes
that the lower relative energies enhance aggregation by further
stabilizing the cross-b core. The AGGRESCAN algorithm is based
on the prior finding that fusion of the amyloid Ab (142) to the green
fluorescence protein (GFP) inhibited the folding and fluorescence of
GFP by forming Ab(142) aggregates, while Ab(142) mutants that
block this aggregation instead promoted fluorescence [66,67].
Systematic in vivo screens of Ab(142) variants fused to GFP
permitted the assignment of intrinsic aggregation propensities for
natural amino acids, and indicated enhanced aggregation for amino
side-chains with increasing hydrophobicity [66,67]. In conjunction
with the experimentally-determined predispositions of amino acids
to aggregate and earlier reports that very short sequences may either
facilitate or inhibit amyloid fibril formation , AGGRESCAN
predicts local hot spots of aggregation in proteins and peptides
other than Ab(142) [63,64]. Theoretical PASTA and
AGGRESCAN predictions of MB and S-MB aggregation were performed
here by submitting the primary sequences of these peptides (Fig. 2) to
the respective PASTA Version 1.0 (http://protein.cribi.unipd.it/
pasta)  and AGGRESCAN (http://bioinf.uab.es/aggrescan/)
Attenuated Total Reflectance-Fourier Transform Infrared
Infrared spectra were recorded at 25uC using a Bruker Vector
22 FTIR spectrometer (Pike Technologies) equipped with a DTGS
detector, averaged over 256 scans at a gain of 4 and a resolution of
2 cm21 [16,48]. For spectral measurements of MB and S-MB in
hexafluoroisopropanol (HFIP) or methanol (MeOH) solutions,
selffilms were first prepared by air-drying peptide originally in 100%
HFIP onto a 5062062 mm, 45u attenuated total reflectance
(ATR) crystal for the Bruker spectrometer. The dried peptide
selffilms were then overlaid with solutions containing 100% HFIP,
40% HFIP/60% deuterated-10 mM sodium phosphate buffer
(pH 7.4) or 100% methanol (MeOH) before spectral acquisition;
control solvent samples were similarly prepared, but without
peptide. Spectra of peptides in solvent were obtained by
subtraction of the solvent spectrum from that of the
peptidesolvent. For FTIR spectra of MB or S-MB in sodium dodecyl
sulfate (SDS), dipalmitoyl phosphatidylcholine (DPPC) or
1palmitoyl-2-oleoyl phosphatidylglycerol (POPG) environments,
lipid-peptide mixtures were spread from
chloroform:trifluoroethanol (1:1, v:v) onto the ATR crystal, and then air-dried under
nitrogen in the sample chamber to form a multilayer film
(SDS:peptide or phospholipid:peptide of 40:1 or 10:1, mole:mole,
respectively). The peptide:lipid films were then hydrated with
deuterium vapor in nitrogen for 1 h prior to acquiring spectra
. The spectra for peptide in SDS, DPPC or POPG were
obtained by subtracting the lipid spectrum with D2O from that of
peptide in lipid with D2O hydration. The proportions of a-helix,
b-turn, b-sheet and disordered conformations of the resulting IR
spectra were determined by self-deconvolution for band narrowing
and area calculations of component peaks using curve-fitting
programs supplied by Galactic Software (GRAMS/AI 8, version
8.0; Thermo Electron Corp.). The frequency ranges for the
different structures were: a-helix (16621645 cm21), b-sheet
(16371613 cm21 and 17101682 cm21), b-turns (1682
1662 cm21), and disordered or random (16501637 cm21) .
Molecular Modeling of Monomeric MB and S-MB
For molecular modeling and molecular dynamics (MD)
simulations of the MB sequence (Fig. 2B), the initial
threedimensional conformation was previously determined from
13Cenhanced Fourier transform infrared (13C-FTIR) spectroscopy of
the disulfide-linked peptide (PDB Accession code: 1SSZ) . The
starting MB structure was chosen as the model with lowest energy,
least violations of spatial restraints, and the highest number of
residues in core regions of the Ramachandran plot. For the
corresponding modeling of S-MB, the peptide backbones of the
lowest energy conformers of the overlapping SP-B(125) (PDB:
1DFW)  and MB (PDB: 1SSZ) structures were used as
templates for the S-MB sequence (Fig. 2A). The sequences were
aligned and homology modeled with Modeller version 9v4
(http://www.salilab.org/modeller/). Both the MB and S-MB
structures were further refined using the GROMACS suite of
MD programs . The homology structures for MB and S-MB
were each placed in a periodic 65 cubic A box of HFIP:spc water
(4:6, v:v) to emulate the solution environment of the FTIR
measurements (equilibrated HFIP solvent box and topology files
courtesy of D. Roccatano) . The respective ensembles
containing either monomeric MB or S-MB peptides were each
minimized by the steepest descent method as implemented in
the GROMACS version 3.3.3 environment  (http://www.
gromacs.org). Chloride counterions were added to the solvent box
with the peptide to neutralize its charge with constraints on the
peptide; the ensembles were then subjected to 100 psec of MD at
300K using the ffG53a6 force field option that allows the solvent
to equilibrate while restraining the peptides. These 0 nsec
systems for either MB or S-MB were then subjected to 100 nsec
MD simulations at 300K without any experimental constraints,
utilizing Berendsen temperature and pressure coupling and the
Particle Mesh Ewald method for evaluating long-range
electrostatic interactions. The time-dependent evolution of the root mean
square deviations (RMSD) for the peptide a-Cs, radius of gyration
and secondary structure (i.e., analyzed using the DSSP criteria
 for the peptide in the HFIP-water environment indicated
when equilibrium was reached. Molecular model structures were
rendered using Rasmol version 220.127.116.11 (http://www.RasMol.org)
and PyMOL v0.99 (http://www.pymol.org).
Methods for Docking Monomer S-MB to Form
The molecular structure for the S-MB homodimer was derived
from the coordinate set generated by the molecular dynamics run
of the SMB monomer in HFIP-water. The conformer of S-MB at
49.8 nsec was selected to model the homodimer because its
conformation was closest to the b-sheet structure for residues
Tyr7 to Arg-12 as predicted from the PASTA and AGGRESCAN
programs (see Results). Two S-MB monomers were initially
docked to form a homodimer using ZDOCK , similar to that
previously described for the docking of the N-terminal domain of
SP-B . The lowest energy conformer of this initial SMB
homodimeric structure was then further refined by using
RosettaDock (www.rosettacommons.org), as implemented in
CAPRI . The docking method was performed using a
twostep process of rigid-body Monte Carlo searching and parallel
optimization of the backbone displacement and side-chain
conformations. Monte Carlo minimization was then employed to
identify a final lowest energy S-MB dimeric structure .
MD Simulation of the Preliminary S-MB Docked
Homodimer in a SDS-Water Environment
Molecular Dynamics (MD) simulations of the S-MB dimer was
accomplished by inserting the RosettaDocked peptide homodimer
into a pre-equilibrated SDS micelle, which was downloaded from the
National Resource for Biomedical Supercomputing (http://www.
membrane.html). The ratio of SDS to dimeric peptide was adjusted
to 28/1 (i.e., SDS/dimer S-MB) by removing excess detergent.
This peptide-SDS ensemble was then minimized in an aqueous
56 A3 solvent box with sodium counter ions for electronic
neutrality with Hyperchem 7.5, using the CHARMM 27 option
. The coordinate set of this minimized peptide-detergent
construct in the PDB format was then ported to the Gromacs
environment, and the structure refined using molecular dynamics
with the ffG53a6 force field. For the Gromacs environment, the
SDS molecule was parameterized using the formalism of
Sammalkorpi et al. . In this peptide-detergent simulation, the
temperature, pressure, electrostatics and bond length constraint
run parameters for the molecular dynamics of the system were the
same as those used for the monomer S-MB-solvent system (see
Captive Bubble Surfactometry
The captive bubble surfactometer used here was a
fullycomputerized version of that described by Schurch and co-workers
. In brief, the sample chamber of the apparatus was cut
from high-quality cylindrical glass tubing (10 mm inner diameter).
A TeflonH piston with a tight O-ring seal was fitted into the glass
tubing from the top end, with a plug of buffered 1% agarose gel
inserted between the piston and the surfactant solution that was
added through a stainless steel port from the other end of the
sample chamber. The chamber and piston were vertically
mounted in a steel rack, the height of which was regulated by a
precision micrometer gear. In a typical experiment, the chamber
was filled with a buffered salt solution (140 mM NaCl, 10 mM
HEPES, 2.5 mM CaCl2, pH 6.9) containing 10% sucrose. One ml
of surfactant solution containing 35 mg of lipid was added to this
subphase, which was stirred by a small magnetic bar at 37uC.
The subphase volume in the sample chamber averaged 0.7 ml
(0.51 ml), resulting in a final average surfactant lipid
concentration of 50 mg/ml (3575 mg/ml). An air bubble approximately
7 mm in diameter (,200 ml in volume) was then introduced
within the sample chamber and subjected to cyclic volume (surface
area) changes by systematically varying the height of the steel rack
following a 5 min pause to allow adsorption to the air-water
interface. The ionic composition of the buffered agarose plug
minimized bubble adhesion to the plug during cycling, so that an
uninterrupted bubble interface was maintained. Surface studies
utilized a compression ratio of approximately 5:1 (maximum area/
minimum area) and a rate of 20 cycles per min. Bubble images
were continuously monitored during compression-decompression
using a digital video camera (PULNIX Model TM-200, Pulmix
America Inc, Sunnyvale, CA) and a professional video recorder
(Panasonic AG-1980P, Secaucus, NJ) coupled to a computer with
an Intel Pentium 4 processor. Selected single frames stored in
RAM were subsequently subjected to image processing and
analysis . Bubble areas and volumes were calculated by an
original algorithm relating bubble height and diameter to areas of
revolution, and bubble surface tension was determined by the
method of Malcolm and Elliot .
Ventilated Lung-Lavaged Rat Model
Animal experiments were performed under established
protocols approved by the Animal Care and Use Committee at the Los
Angeles Biomedical Research Institute at Harbor-UCLA Medical
Center. Anesthesia, surgery, lavage, ventilation, and monitoring
methods used have been detailed previously . Briefly, adult
male Sprague-Dawley rats weighing 200225 g were anesthetized
with 35 mg/kg pentobarbital sodium and 80 mg/kg ketamine by
intraperitoneal injection, intubated, and ventilated with a rodent
ventilator (Harvard Apparatus, South Natick, MA) with 100%
oxygen, a tidal volume of 7.5 ml/kg and a rate of 60/min. An
arterial line was placed in the abdominal aorta for measurements
of arterial blood pressure and blood gases. Rats were paralyzed
with 1 mg/kg pancuronium bromide intravenously. Only animals
with PaO2 values .400 torr while ventilated with 100% oxygen
and with normal blood pressure values were included in the
experiments. Airway pressures were measured with a pressure
transducer (Gould Inc., Cleveland, OH) and tidal volume with a
pneumotachometer (Validyne, Northridge, CA) connected to a
multi-channel recorder (Gould Inc., Cleveland, OH). The lungs
were lavaged 812 times with 8 ml of pre-warmed 0.9% NaCl.
After the PaO2 in 100% oxygen had reached stable values of
,100 torr, the rats were treated with 100 mg/kg of experimental
surfactant by intratracheal instillation. Arterial blood gases, tidal
volume and airway pressures were determined at 15 min intervals
throughout each experiment. Dynamic lung compliance was
calculated by dividing tidal volume/kg body weight by changes in
airway pressure (peak inspiratory pressure minus positive
endexpiratory pressure) (mL/kg/cmH2O). Ninety minutes after
surfactant instillation, rats were killed with 200 mg/kg
pentobarFTIR Spectroscopic Analysis of Mini-B (MB) and Super
Mini-B (S-MB) in Lipid Mimic and lipid environments
The secondary structures for MB and S-MB in either lipid
mimics [i.e., 40% HFIP/60% deuterated-sodium phosphate
buffer, pH 7.4 or 100% methanol (MeOH)], lipid-detergent [i.e.,
sodium dodecyl sulfate (SDS)] or lipids (i.e., POPG or DPPC) were
investigated with conventional 12C-FTIR spectroscopy.
Representative FTIR spectra of the amide I band for MB in these
environments (Fig. 3A) were all similar, each indicating a major
component centered at ,16511657 cm21 with a small low-field
shoulder at ,1620 cm21. Because prior FTIR studies of
proteins and peptides [69,84] have assigned bands in the
range of 16501659 cm21 as a-helical, while those bands
,16131637 cm21 reflect b-sheet, MB likely assumes a-helical
and b-sheet structures and possibly other conformations in these
environments. Self-deconvolutions of the Fig. 3A spectra confirmed
that MB is polymorphic, principally adopting a-helix but with
significant contributions from b-sheet, loop-turn and disordered
components (Table 1). Interestingly, the relative proportions of
secondary structure determined for MB (i.e., a-helix . b-sheet ,
loop-turn , disordered) in both lipids and lipid-mimetics of varying
polarity are all comparable (Table 1), suggesting an overall stability
for the MB conformation that is well-maintained. In agreement with
these results, past 12C-FTIR studies of MB in 90% HFIP/10%
water , or in the lipid-mimetic TFE (pH 7.4) and hydrated
multilayers of the synthetic lipid DEPN-8 , all showed similar
spectra with a major a-helical peak at ,16551658 cm21 and a
small b-sheet shoulder at ,1623 cm21.
Figure 3. ATR-FTIR spectra of MB and S-MB in the structure-promoting solvents hexafluoroisopropanol (HFIP) and methanol
(MeOH), the detergent lipid SDS and the phospholipids POPG and DPPC. Panel A: Stacked FTIR spectra of MB in 40% HFIP (i.e., 40% HFIP/
60% deuterated sodium phosphate buffer, pH 7.4), 100% MeOH (i.e., 100% methanol), deuterated SDS, POPG and DPPC. Panel B: FTIR spectra of S-MB
in 40% HFIP, 100% MeOH, SDS, POPG and DPPC. In Panels A and B, the IR spectra for MB and S-MB each show dominant a-helical components
centered at 16571651 cm21 (arrows), with minor bands at ,16371613 cm21 (arrow at 1620 cm21 ) denoting b-sheet. Peptide concentrations were
470 mM for solvent spectra and 10:1 lipid:peptide (mole:mole) for lipid spectra. The abscissa (left to right) is 17401560 cm21, while the ordinate
represents absorption (in arbitrary units). See text for discussion.
FTIR spectroscopy was also performed to assess secondary
structures for S-MB in both lipid-mimic and lipid environments.
The representative FTIR spectra obtained for S-MB (Fig. 3B) were
similar to the corresponding MB spectra in Fig. 3A with
predominate a-helical peaks between 16511657 cm21 and a
low-field shoulder at 1620 cm21. Deconvolution of these S-MB
spectra confirmed elevated levels of a-helix, with smaller amounts
of b-sheet, loop-turn and random structures (Table 1). The relative
proportions of secondary conformations for S-MB and MB in each
of these environments are comparable in Table 1, suggesting that
inclusion of the short N-terminal insertion sequence (i.e., S-MB
residues 17; Fig. 2A) does not grossly perturb the overall
secondary conformation of the disulfide-linked core shared by MB
and S-MB (Fig. 2). However, it should be noted that somewhat
more residues in S-MB were found to participate as b-sheet in
these lipid and lipid mimics than those in MB (i.e., 811 vs. 58
residues, respectively; Table 1). As was observed for MB, the
relative percentages of secondary conformations for S-MB were
well-conserved in various lipid-mimic and lipid environments
Molecular Dynamics (MD) Simulations of Monomeric MB
and S-MB in a Lipid-Mimic Environment
Molecular dynamics (MD) simulations were next conducted to
provide residue-specific information on monomeric Mini-B (MB)
and Super Mini-B (S-MB) in the lipid-mimic 40% HFIP/60%
water. Although the above 12C-FTIR spectroscopic experiments
are useful for determining secondary structures averaged over
the entire peptide, they cannot indicate the conformations of
individual amino-acids. With starting models based on
experimental structures, MD runs using GROMACS force-fields should
provide worthwhile estimates of the 3D-conformations of both MB
and S-MB in lipid-mimics. Here, we performed MD simulations
using the 13C-FTIR-determined structure of MB in 90% HFIP/
10% water (i.e., the 1SSZ structure)  as the starting model in
40% HFIP/60% water. The 40% HFIP/60% water environment
was chosen for MD simulations for several reasons. First, HFIP
(.,35%) tends to form hydrophobic micellar-like clusters
in water mixtures [85,86] that mimic key properties of either
detergent micelles or lipid membranes [71,87,88]. Second, the
FTIR spectra and secondary conformations for MB in 40% HFIP,
DPPC or POPG were all similar (Fig. 3A; Table 1), indicating that
MD simulations of this peptide in 40% HFIP will be pertinent to
its behavior in lipids.
MD simulations were performed on MB by first calculating a
0 nsec by equilibrating MB in a 40% HFIP/60% water box
with chloride counterions (see Methods). This 0 nsec structure
in Figure 4A differs minimally from the 1SSZ structure  on
which it is based. Fig. 4A indicates that the 0 nsec model is
folded as a helix-hairpin-helix when oxidized, and is stabilized by
disulfide linkages between Cys-8 and Cys-40 and Cys-11 and
Cys34 (Fig. 2B). As expected, the axes of the N- and C-terminal helices
in the 0 nsec model are not parallel, but instead are tilted at an
angle (Fig. 4A) comparable to that seen in the1SSZ structure .
The time course of the adaptation of 0 nsec MB structure to the
lipid-mimetic 40% HFIP was then computed for a 100 nsec-MD
simulation, with the final MB model at 100 nsec (i.e., 100 nsec
structure) shown in Fig. 4B. The evolution of the MB structure
may be characterized from the kinetics of the root mean square
deviation (RMSD) of the a-Cs. The RMSD vs. time plot in Fig. 5A
shows that MB in the 40% HFIP environment reaches an
equilibrium plateau at ,40 nsec. The simulations were further
studied by examining secondary conformations as a function of
time. Figure 6A shows a plot of the MB secondary structure versus
time, and indicates that the major conformational elements are
largely conserved. For example, the 100 nsec structure in Fig. 4B
confirms the presence of N-terminal a-helix (residues 1017), loop
region with a mix of random coil and bend conformations (1829)
and C-terminal a-helix (3036). Similar to the 0 nsec MB
structure in Fig. 4A, the 100 nsec MB model in Fig. 4B folds as
a compact N- and C-terminal helical bundle, with considerable
interactions between hydrophobic side chains across the interhelix
interface. On the other hand, the axes of the N- and C-terminal
helices in the 100 nsec model are now parallel, instead of being
tilted at an angle (Fig. 4A) comparable to that seen in either the
0 nsec or 1SSZ structures . The final 100 nsec ensemble
of MB, HFIP and water molecules also demonstrates that the
amphipathic MB peptides sequesters HFIP (not shown),
comparably to that previously observed for melittin in HFIP/water
Partial validation of the 100 nsec MB structure in 40%
HFIP/60% water (Fig. 4B) is provided by our 12C-FTIR
spectroscopic findings and a previous high-resolution, 2D-NMR
structure of MB in detergent micelles. The secondary structures
obtained from the deconvolution of 12C-FTIR spectra of MB in
40% HFIP (Fig. 3A; Table 1) are broadly compatible with those
predicted in the 100 nsec structure (Fig. 4B). For example,
comparably high a-helix was noted in the FTIR spectrum and in
the 100 nsec structure (Table 1). However, the loop-turn and
disordered components in the FTIR spectrum (i.e., ,45%
combined in Table 1) are somewhat lower than the
roughlycorresponding turn, bend and random coil conformations (i.e.,
,56% combined in Fig. 4B) predicted in the 100 nsec model.
Interestingly, the 100 nsec MB structure in Fig. 4B did not
predict any b-sheet conformation, despite significant b-sheet in
FTIR spectra (Fig. 3A and Table 1; see below). The validity of the
100 nsec MB model may be further tested with an independent
2D-NMR structure determined for MB in SDS-detergent micelles
Figure 4. The evolving 3D model of monomeric Mini-B (MB) in 40% HFIP/60% water at the starting (0 nsec) and ending
(100 nsec) times of the molecular dynamics (MD) simulation. Plate A: Snapshot of MB at 0 nsec. DSSP analysis indicated the following
secondary conformation map (residues in parentheses): coil (8, 22, 2627, 29, 3941); turn (2324, 3738); bend (25, 28); and a-helix (921, 3036) (see
text). Plate B: Snapshot of MB at 100 nsec. DSSP analysis indicated the following conformation map (residues in parentheses): coil (89, 1824, 28
29, 3941); bend (2527, 3738); and a-helix (1017, 3036). In Plates A and B, MD simulations were performed in the GROMACS version 3.3.3
environment (see Methods). The protein backbone structure is shown with color-coded ribbons denoting the following domains: N-terminal helix
(red), turn-loop (green), and C-terminal helix (red) rendered with PyMOL v0.99. Appropriately colored sidechains are shown as stick figures attached
to either the helix (red) or loop (green) ribbon backbones. The orientations of MB in Plates A and B are the same as that for MB in Fig. 2B, with the
Nterminal Cys-8 at the far-left bottom. Disulfide linkages between the N-terminal helix in the foreground and C-terminal helix in the background are
highlighted in yellow.
 (PDB: 2DWF). Respective overlays of the 2DWF structure
with either our 0 nsec or 100 nsec MB models indicated
much better overlap between the 2DWF and 100 nsec
structures (not shown).
Analogous MD simulations were carried out on S-MB in 40%
HFIP. The preequilibration 0 nsec model (Figs. 6B and 7A)
shows that S-MB folds as a helix-hairpin, stabilized by disulfide
bonds between Cys-8 and Cys-40 and Cys-11 and Cys-34, with an
extended tail for the additional N-terminal insertion sequence
(Fig. 2A; residues 1-7). DSSP analysis (Figs. 6B and 7A) indicates
that the helical bundle region for the 0 nsec model is
substantially different from that of the original 1SSZ structure,
with the N- and C-terminal a-helices reduced in length and
realigned so that their axes are now parallel. Time-dependent plots
of both RMSD (Fig. 5B) and DSSP (Fig. 6B) indicate the S-MB
simulation reaches equilibrium at ,65 nsec, with a final
100 nsec structure demonstrating a helix-hairpin bundle
(Fig. 7B) which closely matches the secondary structure and
overall topography of the 100 nsec MB structure (Fig. 4B). This
suggests that the N-terminal insertion domain in S-MB has little
influence on the final organization of the helical core, with it
projecting away from the N-terminal helix similarly to that of the
1DFW structure of SP-B(125) . The N-terminal sequence of
S-MB assumes a largely random coil conformation in the
100 nsec model (Fig. 7B), probably due to its repeated proline
motif which prevents back bonding to form intramolecular b-turns
and/or b-sheets. The flexible N-terminal tail also contributes to
the higher plateau RMSD values observed in Fig. 5B for S-MB
Figure 5. Conformational drift indicated as a-C root mean square deviation (RMSD) from the starting structure for the MD
simulation of the monomeric MB and S-MB peptides. Plate A: Time course of the RMSD in nm from the 0 nsec structure of MB (Fig. 4A). Plate
B: Time course of the RMSD from the 0 nsec structure of S-MB (Fig. 7A). See Methods for experimental details.
Figure 6. Secondary structure (determined with DSSP ) as a function of time for the monomeric MB and S-MB peptides in 40%
HFIP/60% water. Plate A: The 34-residue MB peptide. The abscissa range is 0 nsec to 100 nsec, while that for the ordinate is 8-41 residues (see
Fig. 2B for sequence). Plate B: The 41-residue S-MB peptide. The abscissa range is 0 nsec to 100 nsec, while that for the ordinate is 1 to 41
residues (see Fig. 2A for sequence). The secondary structures indicated are: a-helix (blue), 5-helix (purple), 310-helix (grey), turn (yellow), bend (green)
and coil (white).
than those for MB in Fig. 5A which lacks this sequence (i.e., ,0.6
and ,0.4 nm, respectively). Similar to that noted above with the
100 nsec ensemble for MB, the 100 nsec S-MB structure is
preferentially coated with HFIP selected from the
fluorocarbonwater mixture (not shown).
The final 100 nsec S-MB model (Fig. 7B) obtained from MD
simulations was partially corroborated with our 12C-FTIR spectral
results (Fig. 3B; Table 1). The percent secondary structures
obtained from the deconvolution of 12C-FTIR spectra of S-MB in
40% HFIP/60% aqueous buffer (Fig. 3B; Table 1) approximate
those predicted in the 100 nsec structure (Fig. 7B), but there
remain significant differences. For example, similarly elevated
ahelix levels were noted in the FTIR spectrum and in the
100 nsec structure (Fig. 3B and Table 1). However, the
loopturn and disordered conformations in the FTIR spectrum (i.e.,
,44% combined in Table 1) are reduced from the comparable
turn, bend and random coil structures (i.e., ,63% combined in
Fig. 7B) predicted in the 100 nsec model. It also should be noted
that FTIR spectra of S-MB in 40% HFIP indicated ,21% b-sheet
(Fig. 3B; Table 1), while this minor conformation was not
identified in the corresponding 100 nsec structure for S-MB
(Figs. 6B and 7B).
Molecular Weight (MW) and Aggregation Analyses on MB
and S-MB Peptides Using SDS-PAGE
Both the molecular weight (MW) and self-aggregation properties
of MB and S-MB peptides in a lipid-detergent environment were
next assessed in sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE) experiments. MW determinations for
proteins and peptides in SDS-PAGE are possible because SDS
molecules bind with high affinity to both hydrophobic protein sites
 and positively-charged amino acid residues, and also because a
consistent amount of detergent generally binds to proteins .
Maximum levels of SDS bound to proteins (or peptides) occur at
,1.52 g detergent/g protein . Besides MW measurements of
monomer proteins, SDS-PAGE has also been useful for
investigating the self-aggregation of proteins such as native, full-length SP-B
[56,92]. In our present SDS-PAGE study, MB was initially dissolved
in the loading buffer at ,2 g SDS/g peptide ratio (i.e., ,27 SDS/
MB, detergent to peptide molar ratio), which should be sufficiently
high to saturate detergent-binding sites on the peptide. SDS-PAGE
was then performed on MB using a protocol that earlier detected
native SP-B proteins , and Coomassie blue and silver staining
showed a diffuse, broad band centered at ,3.9 kDa (see Figure 8,
Lane 2) when compared to the standard molecular mass markers
(MW range 2.516.9 kDa) (see Figure 8, Lane 1). This MW
determined for MB from SDS-PAGE was in good agreement with
those assessed from either mass spectrometry (MALDI-TOF) or the
known primary sequence, indicating that MB is monomeric on
SDS-PAGE. Also in support of the above findings are those from a
previous 2D-NMR structural analysis, which demonstrated strong
affinity for monomer MB peptide with micellar SDS .
SDS-PAGE was similarly performed on S-MB and yielded a
broad band at ,8.9 kDa and a much fainter band at ,4 kDa
(Fig. 8, Lane 3). It is likely that the weak ,4 kDa band represents
low amounts of monomeric S-MB, as it corresponds well with the
molecular weights determined from mass spectrometry and the
known amino acid sequence. On the other hand, the apparent MW
for the principal S-MB band from SDS-PAGE (Fig. 8, Lane 3) is
considerably higher than that obtained for monomeric S-MB from
mass spectrometry (i.e., ,8.9 vs. ,4.7 kDa), and consistent with
SMB forming dimers on electrophoresis. Interestingly, the S-MB
peptide, which simulates key structural features of the SP-B leaflet
containing the N- and C-terminal a-helical domains and the
Nterminal sequence, may also partly mimic the ability of full-length
SP-B to form non-covalently associated dimers on SDS-PAGE .
Prediction of Aggregation-Forming Domains in the MB
and S-MB Peptides Using PASTA and AGGRESCAN
To further explore this differential aggregation of MB and
SMB, PASTA  and AGGRESCAN  analyses were next
performed on these peptides. PASTA predicted that MB will
aggregate at both the C- and N-terminal regions. Specifically,
PASTA energy calculations showed that the most likely pairing
will be an antiparallel b-sheet for the C-terminal residues 3440
(Fig. 2B), with a relative energy of 25.49. Using as a benchmark a
database of 179 peptides derived from the literature , an
energy threshold of 25.49 indicates that the probability that MB is
amyloid-like (i.e., highly b-sheet promoting) for this C-terminal
segment is ,90% (true positive rate). Similar PASTA
investigations demonstrated that the most likely pairing in the N-terminal
region was an antiparallel b-sheet for residues 811, with a
lower self-aggregation and a higher energy of 25.33 than the
corresponding most likely pairing in the C-terminal domain. To
account for all possible pairings, a PASTA aggregation profile was
next constructed for MB in Fig. 9A , and confirmed that
segments 3440 and 811 may each form b-sheets; however, this
aggregation plot for MB also showed a much lower self-association
propensity for the N-terminal domain than that for the C-terminal
region (Fig. 9A). An independent analysis of the self-associating
domains in MB was similarly conducted with AGGRESCAN, a
predictive algorithm which combines the known aggregation
tendency of amino acids with earlier findings that short sequences
(,5 residues) either promote or inhibit interpeptide b-sheets (see
Methods). The AGGRESCAN aggregation profile of MB is shown
in Fig. 9B, in which the normalized hot spot area (i.e., directly
proportional to the self-association propensity of a residue) is
plotted as a function of each peptide residue (k). The
AGGRESCAN plot for MB in Fig. 9B confirms our PASTA predictions in
Fig. 9A that not only will the N- and C-terminal domains of MB
be self-association zones, but also that the C-terminal region will
exhibit a higher aggregation tendency than the N-terminal
domain. These PASTA and AGGRESCAN computations suggest
that the b-sheet conformations identified in the FTIR spectra of
MB (Fig. 3A; Table 1) are due to interpeptide b-sheets forming
between opposing amino-acids of the N-terminal (, residues 8
11) and/or C-terminal (,residues 3440) domains. Nevertheless,
these interpeptide interactions are apparently too weak to promote
oligomers for MB on SDS-PAGE, as only the monomer was
observed in Fig. 8.
PASTA and AGGRESCAN calculations were next conducted
on S-MB, and indicated that attachment of the N-terminal
insertion sequence to the MB peptide dramatically enhanced the
self-association properties of the N-terminal region. For example,
PASTA analysis of S-MB demonstrated that an antiparallel
bsheet for the N-terminal segment 712 (Fig. 2A) is now the most
likely pairing for the entire S-MB peptide. The PASTA energy for
the antiparallel b-sheet segment for S-MB is 26.07, significantly
lower than that of the most likely pairing of the C-terminal domain
of S-MB (i.e., segment 3440) with a relative energy of 25.49.
Inclusion of the N-terminal insertion sequence increases the
probability that S-MB is amyloid-like (i.e., highly b-sheet) to
,97% from the ,90% value determined above for MB . The
PASTA aggregation plot for S-MB in Fig. 9A confirms that the
Nterminal region (, residues 712) now shows a predominant
selfassociation propensity over that determined for the C-terminal
region (, residues 3440). A similar AGGRESCAN aggregation
profile for S-MB in Fig. 9B also shows that the self-association
properties of the N-terminal region (, residues 716) are
increased over those noted for the truncated MB peptide, although
here the self-association of the C-terminal region (, residues
3238) is predicted to be slightly higher than that of the
Nterminal domain. Taken together, these PASTA and
AGGRESCAN results suggest that the dimer formation observed for S-MB,
but not for MB, in Fig. 8 is due to S-MB possessing the Tyr-7
Figure 9. Propensity for b-sheet aggregation determined for MB (----) and S-MB (____) from PASTA and AGGRESCAN analyses. Plate A:
Plot of the aggregation propensity [i.e., h(k)], calculated from the PASTA algorithm for the relative energies of the various antiparallel and parallel
bsheet pairings . The h(k) values are plotted for each peptide residue (k), and normalized so that the summation of all h(k)s for a peptide equals 1.0.
Plate B: Plot of the normalized hot spot area as a function of each peptide residue (k), determined from the AGGRESCAN algorithm  for the
relative propensity of local peptide domains to form aggregates. The sequence and numbering for MB and S-MB are in Fig. 2.
residue in the N-terminal insertion sequence (17). Nevertheless,
our subsequent MD simulations of dimer S-MB in a SDS-water
environment indicated that the enhanced self-association observed
with S-MB is not simply due to the inclusion of Tyr-7, but is also
the result of the Phe-1 to Pro-6 sequence participating through a
distinct mechanism (see below).
One cautionary note in using the above PASTA and
AGGRESCAN algorithms is that they do not account for the
actual solvent environment of a given protein domain. Thus, these
predictions may not be used to directly calculate association
constants and binding stoichiometries, which require more precise
binding free energies . Instead, relative aggregation
propensities are assessed for protein sequences with PASTA or
AGGRESCAN, each using databanks to benchmark a given
domain [62,64]. These aggregation propensities are most accurate
for peptide regions exposed to polar environments , such as in
aqueous buffer or the membrane lipid-water interface , but are
less applicable for proteins in hydrophobic milieu. In this context,
it is worthwhile analyzing the lung SP-C (see Introduction) with
these predictive algorithms. SP-C is a hydrophobic 35-amino acid
transmembrane protein that not only exhibits lung surfactant
activities, but also is associated with the onset of pulmonary
alveolar proteinosis (PAP). Bronchoalveolar fluid from PAP
patients is rich in insoluble SP-C aggregates, which show amyloid
properties such as Congo red staining and fibril formation on
electron microscopy . PASTA and AGGRESCAN further
support an amyloid classification for SP-C, as both predict
extremely high aggregation with relative PASTA energies and
normalized hot spot areas of 229.2 (SP-C residues 8-28 as parallel
b-sheet) and 128 (SP-C residues 12-35 as b-sheet) (Gordon et al.,
unpublished observations). Consequently, PASTA and
AGGRESCAN successfully predict only the high b-sheet detected with
FTIR spectra of depalmitoylated SP-C in aqueous buffer at
neutral pH , but not the elevated a-helix found in CD spectra
of this peptide in detergent micelles . Because MB and S-MB
are each likely exposed to water when bound to lipids at the polar
headgroup region [30,49,97], we anticipate that our PASTA and
AGGRESCAN results in Fig. 9 will also accurately forecast
aggregated domains in MB and S-MB. These predictions also
permitted us to develop starting dimer S-MB models for MD
Surface Plasmon Resonance (SPR) Measurements of MB
Given the differential self-aggregation observed for S-MB and
MB on SDS-PAGE (Fig. 8), it is important to determine the direct
binding affinities of these peptides, both to themselves
(selfassociation) and to each other (cross-association). Here, we use
surface plasmon resonance (SPR) spectroscopy to make these
peptide-peptide binding assessments. SPR is a surface-sensitive
methodology, in which the ligand is chemically-linked to a sensor
surface, and the solute is then flowed past the chipped molecule.
The binding of the solute to the immobilized ligand produces an
evanescent response, which is measured in response units (RU)
and is proportional to the bound mass. SPR experiments have
recently determined the self-association properties of amyloid
peptides , which bear some similarities to our synthetic
surfactant peptides (see above). In the present studies, S-MB and
MB were each chemically-linked to chips at their respective
Nterminal amine groups, while the solutes S-MB and MB in buffer
were flowed past the immobilized peptides. Molecular
bindingaffinities (associations) between the soluble peptides and
chiplinked films of S-MB and MB were measured at 37uC using a
Biacore apparatus. Representative sensorgrams of S-MB and MB
binding to chipped S-MB and MB peptides are shown in Fig. 10,
and indicate that the self-association of S-MB is considerably
greater than that observed for MB. Specifically, plots of RU as a
function of time indicated a much higher maximal RU value for
1 mg S-MB/ml buffer flowed past chipped S-MB than that for
1 mg MB/ml buffer flowed over chipped MB (i.e., respective
maximal RU values of 325 and 28). Also consistent with a higher
self-association for S-MB than that for MB is the much lower
dissociation constant (KD) observed for S-MB binding to chipped
S-MB than the corresponding KD for MB binding to chipped MB
(Table 2). Control SPR experiments examining the binding of
SMB to chipped MB, and also MB to chipped S-MB, produced not
only low-response sensorgrams which nearly overlap that obtained
for MB binding to chipped MB (Fig. 10), but also higher KD
values than that observed for S-MB binding to itself (Table 2).
These SPR results indicating much higher self-association for
SMB than that noted for MB are supportive of our SDS-PAGE
findings indicating dimer formation for S-MB but not for MB
Prediction of Dimer S-MB Structures Using ZDOCK and
Because S-MB predominately formed dimers in the above
SDSPAGE (see above), potential 3D-interactions between S-MB
monomers were next assessed with several docking algorithms.
aMB and S-MB in running buffer (10 mM HEPES, 150 mM NaCl, 3 mM EDTA,
0.005% Surfactant P20, pH 7.4) were flowed past monolayers of N-terminal
Cys-MB or N-terminal Cys-S-MB, linked via their respective N-terminal thiol
groups to CSM sensor chips in a Biacore 3000 system (Methods).
Mean kinetic rate constants (kon, koff) and equilibrium dissociation constants
(KD = koff/kon) were determined from curve fitting analyses of SPR results at
1 mg peptide/ml buffer.
ZDOCK is a preliminary stage docking program, which optimizes
shape complementarity, desolvation and electrostatics using Fast
Fourier Transform (FFT) based methods [73,99]. Rather than allow
flexibility in the surface side-chains and/or backbones of the two
proteins, ZDOCK uses searches that permit the proteins to interact
only as rigid-bodies . With S-MB, this soft docking approach may
be particularly useful, given that FTIR spectroscopy (Fig. 3; Table 1)
and MD simulations of the monomer (Figs. 47) indicate that the
disulfide-linked N- and C-helical domains are stable in a wide-range
of lipid-mimic and lipid environments. Because PASTA and
AGGRESCAN programs both predict that the S-MB sequence
containing residues Tyr-7 to Arg-12 is the most stable b-sheet
pairing, initial ZDOCK searches were performed with a 49.8 nsec
simulation of the MD run of monomer S-MB in 40% HFIP/60%
water. As indicated in Fig. 6B, the S-MB sequence 712 adopts an
extended conformation that most closely approximates that of
extended b-sheet. In light of PASTA further predicting that the
lowest relative energy will be an antiparallel b-sheet for residues
712, ZDOCK searches were next conducted with the dimer S-MB
folded as an antiparallel b-sheet for residues ,712. The lowest
energy conformer for the resulting S-MB homodimer maintained a
close antiparallel apposition between the two Trp-Leu pairings (i.e.,
Trp-9A to Leu-10B and Leu-10A to Trp-9B; A and B referring to the
two S-MB sequences in the homodimer). An approximate two-fold
axis lies through the center of the two Trp-Leu pairings that relates
the two leaves of the S-MB dimer. Each of these leaves includes
the helical bundle containing the N- and C-terminal a-helices, and
also the N-terminal insertion sequence (1-7) that adopts an extended
conformation near the N- and C-terminal helices (see Fig. 11A).
Last, the leaves appear to splay about the local dyad in a relatively
The lowest energy conformer from the above ZDOCK search
for S-MB homodimers was then further analyzed by using
RosettaDock , as implemented in CAPRI . Although
ZDOCK permits fast global docking searches, its use of
coursegrained representations for proteins or peptides may not be an
accurate model of the binding surfaces . The RosettaDock
methodology uses a two-step process of rigid-body Monte Carlo
searching and parallel optimization of the backbone displacement
and side-chain conformations. Here, a plot of the energies of 1000
S-MB homodimers versus the RMSD from the initial ZDOCK
conformation was produced by the RosettaDock server. This plot
indicated that the top ten scoring (i.e., lowest energy) candidates
reside in a docking funnel near the starting input conformation
(not shown). Thus, RosettaDock converged to final S-MB dimer
models similar to the lowest energy conformer from ZDOCK.
Indeed, the overall folding pattern of the lowest-energy
RosettaDock model for the S-MB dimer closely reproduced that of the
lowest-energy ZDOCK dimer structure (see above and Fig. 11A).
MD Simulations of Dimer S-MB with SDS in Water
It is of considerable interest to determine whether homodimeric
S-MB structures similar to those identified in the above docking
studies are also present in the SDS-PAGE experiments. MD
simulations on dimer S-MB were conducted here by first inserting
the lowest energy conformer from the above RosettaDock
computations into a SDS-water mileu with a detergent/peptide
molar ratio of 28/1. This peptide-SDS ensemble was then
minimized in an aqueous solvent box with Hyperchem 7.5, using
the CHARMM 27 option (see Methods). The environment for the
resulting 0 nsec structure of dimer S-MB thus closely
approximates that of our SDS-PAGE experiments, which similarly use a
loading buffer containing submicellar SDS at a detergent/peptide
molar ratio of ,28/1 (Fig. 8).
The 0 nsec dimer S-MB model in Fig. 11A shows a peptide
structure comparable to those described above for the lowest
energy conformers in either the ZDOCK or RosettaDock studies
of dimer S-MB. Specifically, the 0 nsec homodimer exhibits a
near antiparallel juxtaposition between the two Trp-Leu pairings
of the monomers. An approximate two-fold axis lies through the
center of these Trp-Leu pairings, which relates the two leaves
that contain not only the N- and C-terminal helical bundles, and
also the extended N-terminal insertion sequences (residues 17)
Secondary structure analysis of the 0 nsec dimer S-MB
structure demonstrates that the N- and C-terminal regions largely
maintain their helical conformations, but that there is some fraying
of the two N-terminal helices at residues Arg-12 to Leu-14. The
topographical organization of the S-MB dimer in Fig. 11A
indicates that the two leaves are in an extended, or open,
conformation that is readily accessible from all sides to SDS
molecules and water. In the 0 nsec dimer S-MB structure of
Fig. 11A, anionic SDS closely associates not only with exposed
nonpolar residues through hydrophobic interactions, but also with
positively-charged Arg and Lys residues through electrostatic
interactions. The binding of SDS to the 0 nsec dimer S-MB
appears to be concentrated in two surface cavities near the center
of the dimer peptide in Fig. 11A, and these two shallow
depressions containing SDS are themselves related by the
aforementioned two-fold axis.
The time course of the adaptation of 0 nsec dimer S-MB in
SDS and water was then calculated for a 10 nsec-MD simulation,
with the final dimer S-MB model at 10 nsec (i.e., 10 nsec
structure) shown in Fig. 11B. The 10 nsec model indicates that
the dimer S-MB converts to a oblate spheroid from the original,
more extended 0 nsec model, with a concomitant merger of the
two shallow cavities into a single deep cavity that is exposed to the
external solvent (Fig. 11). The interior of this central cavity in
dimer S-MB is now lined with hydrophobic amino-acid sidechains
capable of interacting with hydrophobic detergents such as SDS.
Fig. 11B shows that the SDS molecules primarily occupy the
central cavity, and are oriented so that their anionic headgroups
either face the exterior water or form ion pairs with the cationic
residues (i.e., Arg, Lys) that line the outer periphery of the cavity.
The fatty acyl groups of SDS in this cavity are also directed
towards the protein interior. Space-filling models (not shown)
confirm that the 10 nsec dimer S-MB folds as a globular
lipoprotein, consisting of a partial micelle of ,25 SDS molecules
in the central cavity and the dimer peptide providing the
remainder of the structure. Analogous to other
thermodynamically-stable, soluble proteins, the surface of the 10 nsec structure
is enriched in charged or polar molecules (e.g., positively-charged
amino-acids or anionic sulfates of SDS), while the protein interior
is dominated by hydrophobic groups (e.g., nonpolar amino acids
or hydrocarbon tails of SDS molecules) (see Fig. 11B). The local
two-fold axis identified in the 0 nsec structure (Fig. 11A) is
conserved in the 10 nsec structure (Figs. 11B and 12A), and the
10 nsec dimer S-MB model also demonstrates secondary
conformations comparable to those seen in the 0 nsec model,
although with additional fraying of several helical elements
A RMSD vs. time plot shows that the MD simulation of the
entire dimer S-MB ensemble reaches rapid equilibrium and a
plateau phase in ,3 nsec, confirming that the dimer S-MB-SDS
complex is stable in water for the 10 nsec run. The reorganization
of bound SDS likely provides the thermodynamic driving force for
the conversion of the dimer S-MB-SDS complex to the oblate
spheroid conformation (i.e., 10 nsec structure) in Fig. 11B. For
example, the separation between the two antiparallel segments
Tyr-7 to Arg-12 in the 0 nsec structure is dramatically increased
in the 10 nsec structure (see top of Fig. 11). Two SDS molecules
insert between these two opposing strands in the 10 nsec model,
thereby increasing by ,10 A the a-C distance between the
Trp9A and Trp-9B residues. This restructuring of bound SDS is also
probably responsible for the lengthwise narrowing of the central
portion of dimer S-MB, in which the distance between the a-Cs of
Phe-1A and Phe-1B decreases from 42.69 A (0 nsec model) to
38.01 A (10 nsec model). Simultaneous with this narrowing is
the creation of the central cavity in the 10 nsec dimer S-MB
model (Fig. 11). In the 0 nsec or 10 nsec models of Figs. 11
and 12A, the two leaves in dimer S-MB each consist of the N- and
C-terminal helical bundle and the N-terminal sequence (17), and
are related to one another by a local two-fold axis. In the
10 nsec model, however, the two leaves form the walls of the
internal cavity that hold the SDS detergent. The effective
clamping down on the bound SDS molecules by the dimer
SMB leaves in the 10 nsec structure is probably due to strong
hydrophobic interactions between the SDS detergent molecules
and the nonpolar sidechains that project from the backbones of the
helical bundle and the N-terminal sequence in the interior of the
cavity (Fig. 12A). Also contributing to the stability of the dimer
SMB complex are the Arg and Lys residues at the surface periphery
of the cavity, which form electrostatic interactions between the
negatively-charged headgroups of SDS and the positively-charged
Arg (residues 12A, 17A, 12B and 17B) and Lys (16A, 24A, 16B and
24B). The space-filling model for dimer S-MB in Fig. 12B shows
more clearly how the hydrophobic residues are clustered on the
surfaces of the interior cavity, while polar and charged residues
line the periphery of this hydrophobic region. Given that the
present 10 nsec model for dimer S-MB in Fig. 11B was
developed with MD simulation conditions approximating those of
our SDS-PAGE experiments, this structure may also be
responsible for the dimer band observed in our electrophoresis studies
(Fig. 8). Last, it is of interest to note that that our final 10 nsec
model for the docked S-MB dimer adopts a saposin-like
conformation (Figs. 11B and 12) that bears a striking resemblance
to the 3D-structure of saposin C in SDS that was determined using
2D-NMR spectroscopy (Fig. 1)  (see Discussion).
In Vitro Dynamic Surface Activity
Synthetic surfactant preparations were formulated by mixing
synthetic lipids (SL), consisting of 16:10:6:1:2 (weight ratio)
DPPC:POPC:POPG:POPE:cholesterol, with 1.5 mol% S-MB,
MB, SP-B(18), or native pig SP-B. S-MB and MB surfactants
had very high surface activity in captive bubble experiments and
reached identical minimum surface tension values ,1 mN/m
during each of ten consecutive cycles of dynamic cycling (rate of
20 cycles/min, Figure 13). Pig SP-B surfactant (positive control)
reached minimum surface tension values ,3 mN/m, whereas
SPB(18) surfactant, based on the SP-B insertion sequence which is
present in S-MB and absent in MB, and lipids alone (negative
control) reached significantly higher minimum surface tension
values of 20 and 16 mN/m (p,0.001 versus S-MB, MB and pig
SPB surfactants) after ten cycles on the captive bubble surfactometer.
In Vivo Activity of Synthetic Surfactants in Ventilated,
Lung-Lavage Rats with ARDS
The pulmonary activity of S-MB and MB surfactant (described
above) was investigated in comparison to native pig SP-B (positive
control) and SP-B(18) surfactants and lipids alone (negative
control) during a 90 min period following intratracheal instillation
of these surfactants into ventilated rats with ARDS induced by in
vivo lavage. Oxygenation and lung compliance (Figure 14)
increased quickly after instillation of S-MB, MB, and pig SP-B
surfactant. Instillation of the negative control of lipids alone or
SPB(18) surfactant had minimal effects on arterial oxygenation or
compliance. The relative order of pulmonary activity in terms of
both oxygenation and compliance was given as: S-MB . MB .
pig SP-B . SP-B(18). lipids alone (negative control) (Figure 14).
The differences in oxygenation and compliance between S-MB,
MB and pig SP-B surfactants were statistically significant (p,0.05)
starting at 30 min after surfactant instillation and consistently
surpassed the performance of SP-B1-8 and lipids only surfactants
A suite of experimental and theoretical techniques was used
here to develop structural models for monomeric MB and S-MB,
and also dimeric S-MB in lipid-mimic environments. With our
molecular dynamics (MD) simulations conducted in 40% HFIP/
60% water, the 100 nsec models for monomeric MB and S-MB
(Figs. 4B and 7B) indicated that each peptide is organized as a
helical bundle in this membrane-interfacial environment, with
S-MB having an additional flexible sequence (residues 17)
projecting from its core. These monomer simulations were
performed using as initial models the residue-specific structures
determined from 13C-FTIR spectroscopic analyses of the
overlapping disulfide-linked MB (PDB: 1SSZ)  and/or SP-B(125)
(PDB: 1DFW)  peptides. This general approach has been
previously used to investigate MD-simulated interactions between
lipids and SP-B(125) . Given the high a-helical levels in
the 12C-FTIR spectra for either MB or S-MB in both lipids and
lipid-mimics (Table 1), the helical bundle structure observed here
for 100 nsec MB and S-MB in 40% HFIP may also be present
in the surfactant lipids used in our functional assays. Our
subsequent finding that S-MB, but not MB, primarily exists as a
dimer in SDS-PAGE (Fig. 8) prompted us to next perform MD
simulations to characterize the 3D-structure of dimeric S-MB
peptide in SDS and water. Using input structures from ZDOCK
and RosettaDock searches, MD simulations produced a 10 nsec
structure (Figs. 11B and 12) confirming the dimer S-MB may bind
SDS to form a stable complex in water. Although this 10 nsec
dimer structure is only preliminary because it has not been
experimentally verified, such MD simulation models are likely to
increase our understanding of S-MB structure and function. In this
context, elucidation of the dimer S-MB structure using
highresolution techniques, similar to that conducted earlier from
2DNMR analysis of monomer MB with SDS micelles (PDB: 2DWF)
, will be of considerable interest.
It is of interest to compare our 10 nsec dimer S-MB model
with previous experimental structures of related saposin proteins.
An approximate two-fold axis lies through the center of the
10 nsec S-MB dimer, which relates the two leaves each
containing the N- and C-terminal a-helices and the N-terminal
sequence (17) (Figs. 11B and 12A). The space-filling
representation in Fig. 12B indicates that the two S-MB monomers join
seamlessly to form the 10 nsec dimer, despite their association
being due to non-covalent interactions. SDS molecules
predominately bind to an exposed central cavity formed by the
hydrophobic residues that line the concave side of the 10 nsec
dimer S-MB model; contrarily, few SDS bind to the opposing
convex side that is enriched with polar and positively-charged
residues (Figs. 11 and 12). Although the 3D-structure of full-length
SP-B has not been experimentally determined, comparisons are
possible between our 10 nsec dimer S-MB and other saposin
family members with residue-specific structures. A survey of the
experimental saposin structures deposited in the PDB (www.rcsb.
org) (see Introduction) indicated that the 2D-NMR spectroscopic
structure of saposin C with submicellar SDS (PDB: 1SN6) 
(Fig. 1) showed several similarities with our 10 nsec dimer S-MB
model (Figs. 11 and 12). For example, each exhibited analogous
saposin-like folds with their respective leaves in similarly open
conformations. The 10 nsec dimer S-MB model (Figs. 11B and
12) and the saposin C structure (Fig. 1) each have an exposed
central cavity formed by the hydrophobic residues that line the
concave side, and an opposing convex side with polar and charged
residues. Moreover, the external hydrophobic cavities of either the
10 nsec S-MB model (Figs. 11B and 12) or saposin C  were
each observed to bind SDS lipid at submicellar detergent
concentrations. These correspondences are noteworthy, given that
the 10 nsec dimer S-MB must first self-assemble non-covalently
from its monomers, while saposin C has only to fold as a single
Figure 14. Arterial oxygenation and dynamic compliance in surfactant-treated, ventilated rats with ARDS induced by in vivo lavage.
Arterial partial pressure of oxygen (PaO2 in torr) and dynamic compliance (mL/kg/cm H2O) are shown as a function of time for groups of 810 rats
treated with synthetic lung surfactants (synthetic lipids +1.5 mol% S-MB, MB, or SP-B(18)), synthetic lipids +1.5% porcine SP-B (positive control), or
synthetic lipids alone as a negative control. Data are shown as mean 6 SEM.
Our MD simulations indicating that S-MB self-assembles into a
stable dimer protein in a SDS-water environment (Figs. 11 and 12)
may explain the present findings that S-MB, but not MB, forms
dimers in SDS-PAGE (Fig. 8). It is important to note that our
SDS-PAGE experiments are not simply conducted in an aqueous
buffer, but one that also contains 3.47 mM SDS (i.e., below its
CMC of 8.20 mM). Indeed, the basic premise behind MW
determinations in SDS-PAGE is that consistent amounts of SDS
(,1.52 g detergent/g protein) will typically bind to proteins .
Using a similar submicellar SDS concentration in a water
environment, our MD simulations indicated that dimer S-MB
will actually incorporate a partial micelle of SDS to create the
final 10 nsec dimer S-MB ensemble (Figs. 11 and 12). The
remarkable formation of a stable, globular lipoprotein from
constituent S-MB and SDS is attributed to the N-terminal residues
(17) in S-MB (Fig. 2), as analogous dimers are not observed in
SDS-PAGE of MB (Fig. 8). Our MD simulations suggest that the
N-terminal insertion sequence is critical for maintaining the dimer
S-MB structure, because this hydrophobic sequence and the
bundle of N- and C-terminal helices together make up the interior
surface of the exposed cavity which binds SDS (see below).
Conceivably, an analogous dimer S-MB model may also account
for the strong binding of S-MB to itself seen in SPR studies (Fig. 10;
Table 2). Comparable to the SDS-PAGE experiments, SPR
binding assays are performed in an aqueous buffer that includes
4.08 mM polysorbate 20 (P20) (i.e., below its CMC of 4.89 mM).
P20 is an amphipathic detergent, which like SDS has been used as
a solubilizing agent to extract membrane proteins . In a
manner similar to that described for SDS (Figs. 11 and 12), P20
molecules may promote the assembly of the dimer S-MB complex
by forming a partial micelle in the central cavity formed by the
two S-MB molecules. It is also of interest that strong interactions
were noted for S-MB with itself, but not for the self-association of
MB or the cross-association of MB with S-MB (Fig. 10; Table 2).
This suggests that both N-terminal insertion sequences must
be present in the cavity of the dimer peptide, followed by
incorporation with the P20 partial micelle, before strong
intermolecular associations will form between homodimers and/
or heterodimers (see below).
Although the 10 nsec MD simulation of dimer S-MB
provides a reasonable framework to account for our SDS-PAGE
and SPR results (Figs. 8 and 10), it is worthwhile to consider other
structural models by which the N-terminal insertion sequence may
promote the self-association of S-MB. One possible alternative is
that oligomeric S-MB may be due to the N-terminal sequence
(residues 1-7) forming b-sheet. The N-terminal SP-B(19) (Fig. 2)
was earlier proposed to act as a biochemical VelcroH, facilitating
in vivo either the aggregation of SP-B or the interactions of SP-B
with SP-C . In support of this model were earlier 13C-FTIR
spectroscopic studies of SP-B(125) in POPG liposomes, which
indicated that residues 15 participated in interpeptide b-sheet
. Nevertheless, most theoretical and experimental evidence
argues against the N-terminal sequence (17) by itself promoting
S-MB oligomers. SP-B(1-7) does not form a standard
PaulingCorey b-sheet on Ramachandran analysis, but instead belongs to a
broader class of extended conformations that includes the b-sheet
. If anything, the repeated proline motif should act as a b-sheet
breaker [97,106], due to each proline residue lacking the amide
hydrogen and structural constraints imposed by its pyrrolidine
ring. Consistent with this are PASTA calculations indicating high
energies (i.e., low aggregation) for the best pairings of SP-B(17)
(i.e., respective PASTA relative energies of 1.94 for parallel
segments 15, and 2.88 for antiparallel segments 47) . Also
in support of the N-terminal residues (17) not participating in
b-sheet is the earlier FTIR spectral result indicating only minor
bsheet (i.e., 26.3% or ,2.4 residues) for the overlapping SP-B(19)
peptide in methanol, a mimic of the lipid-water interface .
Another mechanism by which the N-terminal sequence (17)
might promote S-MB oligomer formation is through relatively
non-specific aggregation of its hydrophobic sidechains, particularly
in water. Interestingly, earlier ESR studies of the overlapping
B(125), spin-labeled at the amino-terminal Phe-1, demonstrated
that this peptide was primarily aggregated in PBS, exhibiting
spectra that were not only exchange-broadened and
motionallyrestricted, but also insensitive to the paramagnetic broadening
agent chromium oxalate . These ESR results indicated that
spin-labeled SP-B(125) formed high MW oligomers, possibly
involving the formation of a peptide-like micelle with the spin-label
reporter group buried in the interior of the peptide aggregate.
However, it should also be noted that addition of SDS micelles
rapidly dissociated these SP-B(125) aggregates, producing a dilute
ESR spectrum indicating insertion of the spin-labeled N-terminal
SP-B(125) peptide into the SDS micelle. The above non-specific
aggregation mechanism is unlikely to account for the dimer
formation of S-MB in SDS-PAGE for several reasons. First, the
SDS-PAGE in Fig. 8 is not performed in an aqueous buffer, but
one that has high SDS levels that will disaggregate any
nonspecific oligomers of S-MB, analogous to that observed for the
overlapping spin-labeled SP-B(125) . Second, S-MB dimers
based on non-specific hydrophobic interactions between the
Nterminal 17 residues are improbable, because such configurations
were not among the ten lowest energy conformers of the 1000
decoy candidates tested with RosettaDock (see above). Last, our
10 nsec MD simulation for dimer S-MB in SDS-water (Figs. 11
and 12) indicated no conformational drift to this alternative model,
which was subsequently confirmed by extending this MD
simulation run to 50 nsec (data not shown).
Synthetic lung surfactant preparations containing synthetic
lipids plus S-MB or MB had high surface activity in captive bubble
surfactometer experiments, and also improved pulmonary
function and mechanics in ventilated lung-lavaged rats with ARDS to
an even greater extent than porcine SP-B surfactant. These
findings with synthetic lipid/peptide surfactants are consistent with
our recent reports detailing not only enhanced surfactant activity
with MB and typical surfactant lipids in in vitro and in vivo
experiments , but also high surface activity and inhibition
resistance of a synthetic surfactant preparation containing MB
peptide combined at 1.5% by weight with a
phospholipaseresistant phosphonolipid compound (DEPN-8) . These earlier
results, coupled with the surface and physiological activity data
found here for S-MB and MB surfactants, strongly support the use
of SP-B-related peptides in synthetic lipid/peptide exogenous
surfactants for treating lung surfactant deficiency (NRDS) or
injury-induced dysfunction (ALI/ARDS). As the S-MB peptide
investigated here had even greater pulmonary activity than MB,
SMB may be preferred over MB in totally-synthetic surfactant
preparations also containing synthetic lipids.
The active components of endogenous surfactant and all current
exogenous surfactants are lipids and peptides, and this also is the
case for the synthetic surfactants of this study. Endogenous
surfactant contains a complex mixture of lipids, with a
predominant phospholipid content of 8590% by weight and
cholesterol content of 45% by weight ( for review). The
composition of synthetic lipids in synthetic surfactants here was a
mixture of five components (DPPC, POPC, POPG, POPE and
cholesterol) that together accounted for the major lipid-based
molecular interactions in native surfactant. In terms of chain-chain
interactions, the four phospholipids in this lipid mixture
incorporated intermolecular biophysical interactions between
either identical C16:0 acyl chains or between C16:0 and C16:1
acyl chains. Endogenous surfactant phospholipids contain a
substantial content of both these fatty acyl moieties [8,107,108].
In addition, the phospholipids in surfactant lipids allowed for
molecular interactions involving both zwitterionic (PC) and
anionic (PG) headgroups that are prominent in native surfactant.
This includes not only headgroup/headgroup interactions among
lipid molecules themselves, but also lipid headgroup interactions
with charged or polar amino acids in peptides. DOPE and
cholesterol were present in surfactant lipid in smaller amounts
than other lipids, but contributed additional molecular features.
Because of the small size of the PE headgroup relative to PC, the
DOPE molecule has a more wedge-shaped cross-section that
affects molecular packing in lipid bilayers and films. Cholesterol
also has the ability to influence local fluidity/rigidity and packing
in lipid membranes and films [8,109,110] and may additionally
increase lipid adsorption .
The biophysical behavior of lipids in endogenous surfactant is
significantly increased by molecular interactions with three active
apoproteins (SP-A, -B and -C). All these apoproteins interact
strongly with lipids at the molecular level [8,111,112], but peptides
related to SP-B are of special interest because it is known to be
particularly active in improving the adsorption and film behavior
of lung surfactant lipids [8,13,17]. In the present studies, MB and
S-MB were designed to maintain key structural features of the
fulllength human SP-B . The N- and C-terminal domains of
native SP-B actively bind lipids [30,34,39,40,44], and MB and
SMB each incorporates residues 825 and 6378 of human SP-B
that participate in these amphipathic helices (Figs. 4,6,7). These
Nand C-terminal regions are joined in either MB or S-MB by means
of a novel loop domain [16,48], which simulates that occurring in
full-length SP-B . Peptide folding during synthesis is facilitated
by specific solvents to produce the necessary helix-hairpin
structure stabilized by oxidation of cysteine residues, allowing
MB and S-MB to form intramolecular disulfide connectivities
analogous to those between Cys-8 and Cys-78 and Cys-11 and
Cys-71 in human SP-B (residue numbers refer to the full-length
sequence of SP-B) . As reviewed previously , earlier
theoretical and physical studies on peptides based on the N- and
C-terminal domains of SP-B suggest that the cross-linked,
amphipathic helical domains of MB partition into the polar
headgroup region of lipids. Consistent with this are SPR results
indicating that MB will bind to either DPPC or the synthetic lipid
DEPN-8 with high affinity . Given that MB and S-MB in
lipids and lipid-mimics show similarly high a-helix on FTIR
analysis (Fig. 3 and Table 1), and also demonstrate comparable
helical bundles for their shared residues in the lipid-mimic 40%
HFIP (Figs. 4,6,7), it is reasonable to propose that the adjacent,
positively-charged helices in S-MB will also be surface-seeking in
Besides promoting the formation of a dimer S-MB that assumes
a saposin-like fold (Figs. 11 and 12), the N-terminal insertion
sequence with its hydrophobic residues (Fig. 2) may also anchor
the S-MB helices onto surfactant lipids, as has been noted in
earlier physical  and MD simulation  studies of the
overlapping SP-B(125) peptide. Such a dual structural role for the
N-terminal insertion sequence may explain the dramatic
enhancement of in vivo surfactant activities observed here for S-MB
(Fig. 14). The present results support the basic concept that the
Nterminal SP-B(19) may act as a biochemical VelcroH, promoting
the in vivo aggregation of SP-B or the interactions of SP-B with
SPC . Based on our PASTA predictions and docking searches
(see above), however, the self-adhesive region was identified as the
overlapping Tyr-7 to Arg-12 sequence, which likely forms an
antiparallel b-sheet in aqueous environments. The extended
conformation for dimer S-MB, obtained from the lowest energy
RosettaDock searches (e.g., the 0 nsec model in Fig. 11A), may
reflect the dimer peptide in polar environments, such as the
aqueous buffer or at the lipid-water interface.
It is of particular interest that the 10 nsec model for the dimer
peptide in SDS (Figs. 11B and 12) may represent membrane-bound
dimer S-MB, where the dimer adopts a relatively open saposin-like
conformation when interacting with membrane lipids. Before
making this assignment, however, it is important to first assess
whether SDS is a useful surrogate for the typical phospholipids in
membrane lipids. To test this hypothesis, control FTIR experiments
were performed to compare the secondary conformations of MB
and S-MB in SDS. Fig. 3 and Table 1 indicate similar % secondary
conformations for these peptides in either SDS or phospholipids,
consistent with SDS being a reasonable substitute. It should also be
noted that previous ESR experiments using the spin-labeled
Nterminus of SP-B(125) indicated comparable incorporation of the
N-terminal Phe of SP-B(125) into either SDS micelles or
phospholipid bilayers . Moreover, the 3D-structure of MB in
POPG liposomes determined from residue-specific 13C-FTIR
spectroscopy  (PDB: 1SSZ) was very similar to that obtained
from 2D-NMR analysis of MB in SDS micelles  (PDB: 2DWF).
These latter findings are consistent with prior 2D-NMR analyses
suggesting that membrane proteins may adopt native conformations
when incorporated into SDS [20,113]. Collectively, these findings
from control experiments indicate that SDS may be a reasonable
first approximation of membrane phospholipids. Our 10 nsec
model (Figs. 11B and 12) shows that dimer S-MB integrates a partial
SDS micelle into its structure, which then separates the antiparallel
b-sheet pairing (i.e., Tyr-7 to Arg-12). Each monomer in the
10 nsec dimer S-MB model optimizes its interactions with SDS,
yet the overall organization of the dimer S-MB and its local two-fold
axis are still retained. Thus, the 10 nsec dimer S-MB model
(Figs. 11B and 12) may be the surfactant-active configuration, which
inserts into the membrane so that its N-terminal sequences (17) are
immediately subjacent to the polar lipid headgroup, and its
amphipathic N- and C-terminal helices are bound at the
lipidwater interface with their hydrophobic faces oriented inward
[30,44,97,103]. In this context, it should be noted that recent
coaxial confocal-AFM imaging and FRET assays  suggest that
saposin C may perturb membrane bilayers by adopting an open
conformation, thereby exposing its inner hydrophobic surfaces for
interactions with lipid acyl chains. Because both saposin C (Fig. 1)
 and the 10 nsec dimer S-MB model (Figs. 11B and 12) share
an open, saposin-like conformation when bound to submicellar
SDS, it is tempting to speculate that dimer S-MB may exert its
surfactant activities by analogously binding to lipid bilayers and
monolayers. The above results also indicate not only that the
extended N-terminal insertion sequence (residues 112) anchors the
dimer S-MB in lipids and promotes the self-association of S-MB, but
also that these two structural properties may be antagonistic to some
extent. Conceivably, the expression of optimal surfactant activities
produced by either S-MB or native SP-B proteins may depend on
the N-terminal sequence (112) achieving a finely-tuned balance
between protein self-association and insertion into lipids.
Functional and structural studies in our laboratory are continuing
to investigate the surfactant properties of additional SP-B peptide
mimics, including covalently-linked dimer forms (e.g., Ref. ),
SPB constructs containing extended regions of the full-human sequence,
and SP-B variants with modified sequences. Here, we report that the
high in vivo surfactant activity for S-MB may be partially due to the
ability of this peptide to form non-covalently associated dimers. These
findings are broadly supportive of previous experiments indicating
that the dimeric full-length SP-B, associated through either covalent
or non-covalent linkages, shows elevated in vitro and in vivo surfactant
activities over those of the monomeric protein [92,115]. Although the
present experiments demonstrated higher in vivo surfactant activity for
S-MB than for native SP-B (Fig. 14), our in vitro results also indicated
that S-MB does not completely reproduce all of the surface-tension
lowering properties of full-length SP-B in captive bubble
surfactometry (Fig. 13). The discrepancies between our in vivo and in vitro
findings suggest that further modifications of S-MB may be required
to obtain a fully-optimized SP-B mimic.
In summary, FTIR spectroscopy of S-MB and MB in lipids and
lipid-mimics showed that these peptides exhibit similar
conformations, with primary a-helix and secondary b-sheet and loop-turns.
With each peptide treated as a monomer in a lipid-mimic
environment, subsequent MD simulations indicated that S-MB and
MB not only share the same bundle of adjacent N- and C-terminal
ahelical domains [48,49], but also that the N-terminal insertion
sequence (residues 17) of S-MB assumes an extended conformation
projecting from its helical core. SDS-PAGE electrophoresis
demonstrated that S-MB was dimeric in submicellar SDS concentrations,
while MB was monomeric. SPR, predictive aggregation algorithms,
and MD and docking simulations further suggested a preliminary
model for dimeric S-MB with SDS, in which monomers
noncovalently associate to form a dimer S-MB peptide with a
saposinlike fold in both aqueous and lipid environments. The external
N-terminal insertion domain (residues 112) may act as a
biochemical VelcroH  to fasten S-MB molecules together into
a dimer peptide that adopts the open saposin-fold when bound to
lipids. These membrane-associated dimer S-MB peptides may
possibly self-associate to form protein-rich networks in lipids,
analogous to those observed for native SP-B and other SP-B peptides
containing the insertion sequence [32,116]. Besides promoting
peptide self-association, these and prior investigations with SP-B
analogs indicate that the hydrophobic N-terminal insertion sequence
may assist in anchoring S-MB into lipid bilayers and monolayers. In
vitro and in vivo experiments also indicated that S-MB and MB each
exhibits a range of surfactant activities, with S-MB showing greater
oxygenation and dynamic compliance in animal models than MB.
Consequently, our functional studies are supportive of earlier results
with SP-B peptides and peptide analogs, which demonstrated that
this N-terminal insertion sequence plays critical roles in the
expression of in vitro surfactant activities [38,52,53].
Conceived and designed the experiments: FJW AJW LMG RHN.
Performed the experiments: FJW AJW JMHJ ZW CLJ PR APC WMS
SS. Analyzed the data: FJW AJW JMHJ LMG CLJ. Contributed reagents/
materials/analysis tools: RHN. Wrote the paper: FJW AJW LMG RHN.
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