Supported Lipopolysaccharide Bilayers - Langmuir (ACS Publications)

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Supported lipopolysaccharide bilayers Stefan Kaufmann, Karin Ilg, Alireza Mashaghi, Marcus Textor, Bernhard Priem, Markus Aebi, and Erik Reimhult Langmuir, Just Accepted Manuscript • DOI: 10.1021/la3020223 • Publication Date (Web): 25 Jul 2012 Downloaded from http://pubs.acs.org on July 30, 2012

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Supported lipopolysaccharide bilayers Stefan Kaufmann1, Karin Ilg2, Alireza Mashaghi1, Marcus Textor1, Bernard Priem3, Markus Aebi2, Erik Reimhult4*

1

Laboratory for Surface Science and Technology, Department for Materials Science, ETH Zurich, Switzerland 2

Institute of Microbiology, Department of Biology, ETH Zurich, Zurich, Switzerland 3

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CERMAV-CNRS, Grenoble, France

Department of Nanobiotechnology, University of Natural Resources and Life Sciences Vienna, Austria

* [email protected]

In this report the formation of supported lipopolysaccharide bilayers (LPS-SLBs) is studied with extracted native and glycoengineered LPS from Escherichia coli (E. coli) and Salmonella enterica sv Typhimurium (S. Typhimurium) to assemble a platform that allows measurement of LPS membrane structure and the detection of membrane tethered saccharide-protein interactions. We present QCM-D and FRAP characterization of LPS-SLBs with different LPS species, having for e.g. different molecular weights, that show successful formation of SLBs through vesicle fusion on SiO2 surfaces with LPS fractions up to 50 wt%. The thickness of the

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LPS bilayers were investigated with AFM force-distance measurements which showed only a slight thickness increase compared to pure POPC SLBs.

The E. coli LPS were chosen to study the saccharide-protein interaction between the Htype II glycan epitope and the Ralstonia solanacearum lectin (RSL). RSL specifically recognizes fucose sugars, which are present in the used Htype II glycan epitope and absent in the epitopes LPS1 and EY2.The E. coli LPS were chosen to study the saccharide-protein interaction between the Htype II glycan epitope and the Ralstonia solanacearum lectin (RSL) that specifically recognizes the fucose unit of Htype II structure. We show via fluorescence microscopy that the specific, but weak and multivalent interaction can be detected and discriminated on the LPS-SLB platform. The detailed chemical structures of the used glycan epitopes can be found in the Supporting Information Figure 1.

KEYWORDS Supported lipid bilayer, lipopolysaccharide, lectin, multivalent interactions, bacteria membrane mimic, glycomimicry.

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INTRODUCTION The outer membrane of Gram-negative bacteria contains approximately 20 - 25 wt% phospholipids, 20 - 30 wt% lipopolysaccharides (LPS) and 45 - 50 wt% proteins1. The dense packing of the acyl chains, lateral interactions of the headgroups and sugar moieties by dipoledipole, water bridges, H-bonds and cations (e.g. Ca2+ neutralizing the negative charges in lipid A2) result in a stable permeability barrier. It prevents diffusion of large hydrophilic and small hydrophobic molecules into the cell and makes Gram-negative bacteria generally more resistant to hydrophobic drugs and detergents than Gram-positive bacteria2-3. Lipopolysaccharides (LPS) are typically comprised of lipid A, an inner core, an outer core and the O specific chain. The LPS has significant importance for Gram-negative bacteria as usually the minimal substructure lipid A and 3-deoxy-D-mannooctulosonic acid (KDO) are required for growth of Gram-negative bacteria (with exceptions found in the strains of Sphingomonas and Neisseria meningitidis)4. The inner core of LPS is composed of KDO, heptose, hexuronic acid5 or phosphate whereas the outer core exhibits branched oligosaccharides of neutral sugars and the O-specific chain repeating oligosaccharide units. Relative to the structure of their LPS bacteria are referred to as "deep-rough" with only lipid A and inner core, as "rough" with an additional outer core and “smooth” if the membrane contains O-antigen repeats.

Recognition of host glycans enables pathogens to attach to their host cells and at the same time recognition of pathogen saccharides can initiate an immune response in higher organisms to defend themselves against foreign invaders. This competition between pathogenicity and immune response is believed to have played a role in the structural diversity of glycan structures6 and renders the O-antigen components attractive drug and vaccine targets7.

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The importance of glycan-binding proteins (lectins) led to the development of glycan arrays to screen for potential saccharide-lectin interactions8. Low affinity and selectivity of monovalent lectin interactions however provide a challenge for array platforms. Lectins frequently offer multiple binding sites allowing for multivalent interactions and potential screening platforms have to match the separation and orientations of saccharide ligands. Current platforms utilize multivalent scaffolds to obtain multivalent binding with e.g. dendrimers, polymers and lipids9. The structure of LPS with branched oligosaccharide and denser packed acyl chains compared to phospholipids results in a different behavior in the cell membrane. For instance, the mobility of LPS in the outer membrane of S. Typhimurium was reported10 to be as low as 2 × 10-2 µm2/s, two orders of magnitude slower than phospholipids in native membranes11. LPS in smooth S. Typhimurium with repeated O-antigens offers more efficient protection from membrane disrupting molecules than e.g. LPS in rough E. coli12 highlighting the importance of the properties of the LPS layer. The mechanical properties are particularly interesting to be examined in bacterial membrane mimics as potential models to study how LPS mediates bacteria-host interactions. A versatile model system to study membrane molecules in a close to native environment is provided by supported lipid membranes, which are extended membranes assembled on solid supports13. Due to their tunable similarity in composition to native cell membranes and ease of combination with surface-based sensing14 and imaging techniques15, such synthetically produced membranes are increasingly used to study (trans)membrane proteins16, lipid-lipid interactions17, lipid-protein interactions18 or even cell-cell contacts19. Addition of native components such as native lipids or membrane proteins, allows the characterization of the behavior of these constituents in a well-defined and controlled environment. Negatively charged LPS make it difficult to form supported LPS membranes and it is not straightforward to obtain various LPS mutants at sufficient purity since naturally derived LPS 4


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generally display substantial structural heterogeneity. This complicates the investigation of the precise interactions with the substrate. Despite these obstacles several studies of SLBs mimicking bacterial membranes can be found in the literature. Merz et al.20 studied the formation of bacterial membrane mimics on solid substrates via vesicle self-assembly with lipids ranging from POPC/POPG to E.coli total lipid extract. Nomura et al.21 investigated the degree of mixing of Re mutant LPS from E. coli strain WBB06 in supported membranes comprising egg-PC, POPE and POPG. The POPG lipid bilayers, meant to mimic bacterial membranes, were deposited with the Langmuir-Blodgett and Langmuir Schäfer techniques. POPG membranes were later used by Nguyen et al.22 to study interaction and orientation of the antimicrobial peptide magainin 2. Tong and McIntosh23 used a positively charged polymer polyethylenimine (PEI) as support to form LPS-SLBs with deep rough (Rd) from Salmonella enterica sv. Minnesota (S. Minnesota) strain R7, rough (Ra) LPS from E. coli EH100 and mixtures thereof with bacterial phospholipid via vesicle fusion. Gutsmann et al.24 used lipid species PS, deep-rough LPS from S. Minnesota strain R595, DPhyPC, PE and SM and formed planar lipid membranes by opposing two lipid monolayers at the aqueous phase (Montal-Müller technique) to study the interaction and orientation of the lipopolysaccharide-binding protein (LBP) in the membrane. Most of the above described approaches circumvent the difficulty of self-assembling bacteria mimetic lipid bilayers at solid interfaces in a near-native state by investigating monolayers at the air-water interface, on hydrophobic substrates or SLBs assembled on strongly positively charged polymers. The strong interaction with the positively charged polymer or the potential expansion on the hydrophobic interface makes these approaches uncertain for investigations of phenomena related to the natural fluidity, integrity, density or the mechanical properties of the membrane. In this paper E. coli strain K-12, wild type and several mutants, and S. Typhimurium LPS were chosen to demonstrate that formation of fluid supported lipopolysaccharide bilayers (LPS5


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SLBs) can be accomplished to study LPS-lectin interactions and the increase in the membrane thickness due to the additional lipopolymers. Several different LPS mutants were used to demonstrate the generality of the assembly and characterization approaches and to serve as controls for the specific recognition. The LPS from the two different species, E. coli and S. Typhimurium differ in their structure: E. coli K-12 LPS are rough and S. Typhimurium LPS are smooth. LPS-lectin interactions between the artificial Htype II LPS and Ralstonia solanacearum lectin (RSL) that specifically recognizes the fucose unit of the Htype II epitope were investigated. The detailed chemical structures of the used glycan epitopes can be found in the Supporting Information Figure 1. AFM force-distance measurements were performed to investigate differences in the mechanical resistance offered by LPS-SLBs relative to SLBs containing no polymer headgroups and relative to PEG-lipopolymer systems25.

EXPERIMENTAL BUFFERS Water used for the preparation of buffers, suspensions and for the cleaning of substrates was obtained by a Milli-Q Gradient A10 (Millipore, USA) purification system. The system is equipped with an Elix 3 device (three step purification) and an ultraviolet lamp for photooxidation. The resistance of the ultrapure water was 18.2 MΩ and at most 4 ppm (TOC). Tris buffered saline (TBS) was prepared by mixing 10 mM Tris(hydroxymethyl)aminomethan (Sigma, Switzerland) and 150 mM sodium chloride in water. The pH was adjusted to pH 7.4 by adding a solution of 2 M hydrochloric acid (HCl). TBS with calcium (Ca2+) was obtained by adding 2 mM (if not otherwise stated in the text) of calcium chloride (CaCl2) from a 1 M stock solution in TBS. TBS with urea was obtained by adding 20 mM urea (Sigma, Switzerland) prior to mixing with water. The buffers were stored at 4 °C and were filtered by 0.22 µm filters (Sartorius, France) before use. 6


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LIPIDS Lipids were purchased from Avanti Polar Lipids (USA), Sigma (Switzerland) or extracted from bacteria (see below) and stored in chloroform at −24 °C or in water at 4 °C. The LPS Rd mutant (rough strain) was purchased from Sigma Aldrich (Switzerland) and was extracted by the phenol:chloroform:petroleum ether method

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from E. coli F583 (Rd mutant) as stated by

the company. For a schematic representation of the extracted lipids see Figure S1 in the Supporting Information. EXTRACTION OF LIPOPOLYSACCHARIDES The extraction was adapted from Galanos et al.

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modified with an acetone precipitation of

the LPS from the phenol phase27. An O-deacylation of the extracted LPS was further performed by a mild hydrazine treatment28 except for the Rd mutant from Sigma Aldrich which was used as-delivered. This O-deacylation also cleaves the O-acetyl group linked to the abequose in the Salmonella-LPS. LPS with the acronyms LPS129, EY2 (see Supporting Information), wild type and Htype II30 were extracted from Escherichia coli K-12 wild type and the respective mutants. The LPS denoted S-LPS was extracted from Salmonella enterica serovar Typhimurium serogroup B. MALDI-TOF spectra of LPS1, EY2 and Htype II and SDS-polyacrylamide gel electrophoresis (SDS-PAGE) of S-LPS are shown in Figure S2 and S3 in the Supplementing Information. The heterogeneous mixture of LPS as indicated by MALDI-TOF for E. coli and SDS-PAGE for S. Typhimurium implies that the nominal wt% used to name the sample is lower than the actual concentration in solution and therefore also in the SLB. LIPOSOMES A clean round bottom flask was thoroughly rinsed with water and dried under a stream of nitrogen. Subsequently a small amount of chloroform (≈ 400 µl) was added to clean the flask with the solvent prior to adding of the lipid solutions. The lipids were mixed in the desired 7


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ratios and were dried under a stream of nitrogen for 1 hour followed by resuspension in the desired buffer solution. For LPS dissolved in water the drying time was extended up to 2 h, as in the case of Htype II and S-LPS. Unilamellar vesicles were obtained by extruding the solution 31 times through two stacked polycarbonate membranes (Avestin, Canada) with a pore size of 100 nm using a Liposofast extruder (Avestin, Canada) 31. The extruded vesicles were checked by dynamic light scattering (DLS, Malvern Zetasizer Nano, United Kingdom) yielding a consistent overall average diameter of 98 ± 8 nm (polydispersity index 0.067 + 0.024). RSL The Ralstonia solanacearum lectin (RSL) was obtained by expression cultures of the strain with the recombinant plasmid carrying the RSL gene (received from Prof. Dr. Anne Imberty32). The protein was purified by affinity chromatography. Biotinylation was performed with the commercially available EZ-link Sulfo-NHS-LC biotinylation kit (Pierce Protein Research Products from Thermo Fisher Scientific, USA). QUARTZ CRYSTAL MICROBALANCE WITH DISSIPATION MONITORING (QCM-D) A Q-Sense E4 (Q-Sense AB, Sweden) quartz crystal microbalance with dissipation monitoring (QCM-D) was used in this work. Quasi-simultaneously, the 3rd, 5th, 7th, 9th, 11th and 13th overtones were recorded and the 5th was used for all comparisons in this paper. SiO2 coated 4.95 MHz QCM-D crystals were purchased from Q-Sense (QSense AB, Sweden). Before the first use they were cleaned in an ultrasonic bath (Bandelin, Germany) in toluene and isopropyl alcohol for 10 min. Before measurements the crystals were cleaned with 2% sodium dodecyl sulfate (SDS; Sigma, Switzerland) and ethanol in an ultrasonic bath for 10 min and treated in a pre-heated UV-Ozone cleaner for 30 min. The sample solutions were pumped into the cells with a high-precision multichannel dispenser (IPC Ismatec SA, Switzerland) exchanging 0.8 ml (QCM-D cell volume 40 µl). Measurements were done in batch exchange if not stated otherwise to keep consumption of 8


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lipid solutions low. The flow rate for inserting the vesicle solutions was set to 579 µl/min (maximum pump speed), which was then switched off for batch experiments. If not otherwise stated the flow rate was 20 µl for continuous flow experiments. FLUORESCENCE RECOVERY AFTER PHOTOBLEACHING (FRAP) A Zeiss LSM 510 (Carl Zeiss AG, Germany) equipped with a 25-mW Argon laser and 63× 1.4NA, 40× 1.3NA, 20× 0.8NA and 10× 0.3NA objectives was used for FRAP measurements. Glass substrates (Nexterion borosilicate glass, class 100 clean-room cleaned) were obtained from Schott (Germany) or from Menzel GmbH (Germany) with dimensions of 24 mm × 24 mm × 0.17 mm. Glass substrates were rinsed with ethanol and water before use and dried with nitrogen. The substrates were treated in a pre-heated UV-Ozone cleaner for 30 min and mounted in a home-built microscopy cell made of polyether ether ketone. The microscopy cell was filled with the desired vesicle solution. Before starting measurements the samples were rinsed ten times with the respective buffer to remove remaining vesicles from the solution. The image size was chosen as 512 × 512 pixels at a zoom factor of 2 (71.3 µm × 71.3 µm) and the time delays used during the measurements were 0 s, 10 s and 1 min and were set manually depending on the recovery rate. The circular bleach spot had a diameter of 22 µm. For the evaluation of the FRAP data we chose the method based on spatial frequency analysis of averaged radial data presented by Jönsson et al.33 for the determination of diffusion coefficients in this work. LECTIN BINDING ASSAYS RSL labeled with fluorescein isothiocyanate (RSL-FITC) that binds specifically to the fucose unit was used for the binding assays with Htype II. The SLBs were rinsed ten times with TBS buffer after the SLB formation to remove excess vesicles in all fluorescence microscopy experiments. Next, the SLBs were incubated with 8 µg/ml solution of RSL-FITC for 1 h after which it was rinsed ten times with TBS to remove excess lectins from the solution. Most 9


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monovalently bound lectins will be removed by this procedure due to the low affinity of RSL34. Multivalently bound lectins will remain bound to the SLB and are detected by fluorescence microscopy. Biotinylated-RSL was used with a streptavidin label (FluoProbes647, Interchim, France) for assays with Htype II SLBs and SLBs lacking Htype II where 8 µg/ml biotinylated-RSL and subsequently 10 µg/ml fluorescently labeled streptavidin was bound to the SLBs. This assay was performed with a different fluorophore to avoid photobleaching as occurred with FITC.

RESULTS SUPPORTED LIPID BILAYER FORMATION WITH E. COLI AND S. TYPHIMURIUM LPS All liposomes used for SLB formation consisted of the main lipid component POPC and varied fractions of LPS. The dilution of the LPS with POPC was chosen due to existing protocols demonstrating that POPC drives liposome fusion to SLBs on silica surfaces35. Figure

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1Figure 1 (a) shows QCM-D curves of LPS-SLB formation with the different E. coli LPS species at a concentration of 10 wt% and a vesicle concentration of 0.1 mg/ml. For the Rd mutant and Htype II 2 mM Ca2+ and 20 mM urea were used respectively, as buffer additives to improve the SLB formation36. The QCM-D curves demonstrate the resonant frequency and dissipation kinetics of successful LPS-SLB formation for all samples37. Figure 1Figure 1 (b) displays the stable ∆f (solid line, ) and ∆D (dashed line, ) values after the SLB formation with 10 wt% of the different types of E.coli LPS. Fig 1 (c) displays the same data for the specific LPS Htype II as function of LPS concentration.

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Figure 1: QCM-D data of LPS-SLB formation. ∆f is displayed as solid line and filled symbols and ∆D with dashed lines and blank symbols. (a): QCM-D curves of the SLB formation for POPC liposomes containing 10 wt% of LPS1 (, ), EY2 (, ), Rd mutant (, ), wild type (), Htype II (, ). (b) – (d): Stable ∆f and ∆D values after the SLB formation of LPS-SLBs with average values and standard deviations. Numbers in brackets indicate number of repeats. (b): 10 wt% Htype II(3), Rd mutant(2), LPS1(2), EY2(4), wild type(4) and as comparison for pure POPC(3). (c): 10 wt%(3), 15 wt%(1), 20 wt%(1) and 50 wt%(4) of HtypeII. (d): 5 wt%(2), 10 wt%(2), 20 wt%(5) of S-LPS.

The concentrations of Htype II used were 10 wt% (), 15 wt% (), 20 wt% (), 50 wt% (). TBS with 20 mM urea was used for low Htype II concentration (10 wt%, 15 wt%, 20 wt%) and in the case of 50 wt% TBS with 2 mM Ca2+ was used. Similarly Figure 1Figure 1 (d) shows the stable ∆f (solid line, ) and ∆D (dashed line, )


values after the SLB formation with S. Typhimurium LPS (S-LPS) as function of LPS concentration. For the formation of SLBs with S-LPS TBS with 2 mM Ca2+ was used for all concentrations (5 wt%, 10 wt%, 20 wt%). Higher variations in the dissipation shift in QCM-D measurements of S-LPS (e.g. for 10 wt%) are suspected to be caused by variations in the glycan 11


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structure of the S-LPS or the occurrence of more a few defects in the membrane. The overall integrity of the membrane is however not compromised as demonstrated by additional FRAP measurements as presented below, suggesting that the major variation is due to the inherent sample variation in terms of glycan structure for the S-LPS..For the formation of SLBs with SLPS TBS with 2 mM Ca2+ was used for all concentrations (5 wt%, 10 wt%, 20 wt%).

In Table 1 the QCM-D values of the E. coli and S. Typhimurium. LPS-SLB measurements are listed. The adsorbed mass is roughly proportional to ∆f and is increasing for SLBs with incorporated LPS. The frequency shifts with the lowest values are found for POPC with ∆f = 25.7 ± 0.6 Hz and the highest for the wild type with ∆f = -31.2 ± 1.8 Hz. Additionally, FRAP measurements were performed to determine the mobility of POPC in the LPS-SLBs by using the labeled NBD-PC lipid. The diffusion coefficients and recovered fractions are also presented in Table 1, which all demonstrate very high quality SLB formation.

Table 1: QCM-D and FRAP values obtained for E. coli and S. Typhimurium LPS-SLBs. The frequency ∆f and dissipation shifts ∆D were obtained from QCM-D. The adsorbed mass m was calculated from the frequency shift ∆f via the Sauerbrey relation (k = -18.06 ng/cm2). FRAP measurements yield the diffusion coefficients D and the recovered fraction. ∆f

∆D

m

D

recovered fraction

Species

wt%

(Hz)

(10-6)

(ng/cm2)

(µm2/s)

(%)

POPC

100

-25.7 ± 0.6

0.2 ± 0.1

464 ± 11

1.2 ± 0.2

93 ± 3

LPS1

10

-28.1 ± 2.3

0.4 ± 0.1

507 ± 41

1.6 ± 0.1

97 ± 2

EY2

10

-28.5 ± 1.6

0.4 ± 0.1

515 ± 29

1.7 ± 0.1

96 ± 3

Rd

10

-28.5 ± 0.4

0.3 ± 0.1

515 ± 7

0.7 ± 0.1

94 ± 2

Htype II

10

-27.0 ± 0.7

0.3 ± 0.1

488 ± 13

2.0 ± 0.2

95 ± 3

Htype II

50

-30.1 ± 2.5

0.5 ± 0.2

544 ± 45

1.1 ± 0.1

99 ± 1

wild type

10

-31.2 ± 1.8

0.7 ± 0.2

563 ± 32

1.5 ± 0.1

98 ± 1

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S-LPS

5

-26.3 ± 0.9

0.4 ± 0.1

475 ± 16

1.6 ± 0.1

98 ± 2

S-LPS

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-29.5 ± 0.3

0.9 ± 0.5

533 ± 5

1.6 ± 0.2

99 ± 1

S-LPS

20

-28.4 ± 0.6

0.7 ± 0.1

513 ± 11

1.5 ± 0.2

98 ± 2

Figure 2: QCM-D data of LPS-SLB formation. ∆f is displayed as solid line and filled symbols and ∆D with dashed lines and blank symbols. (a) LPS-SLB formation of 50 wt% Htype II performed in buffers with different additives with TBS (, ), TBS + 2 mM Ca2+ (, ) and TBS + 20 mM urea (, ). (b) LPS-SLB formation with S-LPS with and without 2 mM Ca2+. 10 wt% S-LPS with (, ) and without 2 mM Ca2+ (, ) and 20 wt% S-LPS with (, ) and without Ca2+ (, ).

The SLB formation of bacteria mimics can be improved with buffer additives. The formation of 50 wt% Htype II proceeds in TBS, but is slightly accelerated with 20 mM urea and significantly accelerated with 2 mM Ca2+ as shown from QCM-D results in Figure 2Figure 2


(a). For the formation of 20 wt% S-LPS containing SLBs the addition of 2 mM Ca2+ is essential in the inspected time frame (4 h) and accelerates the formation for lower concentrations as shown in Figure 2Figure 2 (b).


COMPRESSION OF SUPPORTED LIPOPOLYSACCHARIDE BILAYERS Figure 3Figure 3 (a) shows averaged force-distance curves of 10 wt% EY2 (), 10 wt% wild type (), 10 wt% S-LPS () and 20 wt% S-LPS () with an additional averaged forcedistance curve of POPC () as comparison. The SLB rupture forces which specify the break13



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through of the AFM tip through the membrane are lowest for 10 wt% EY2 and increase for 10 wt% wild type, 10 wt% S-LPS and 20 wt% S-LPS. Figure 3Figure 3 (b) shows a magnification of the compression before the rupture of POPC, 10 wt% S-LPS and 20 wt% S-LPS of the averaged curves. POPC and 10 wt% S-SLP show similar and 20 wt% a longer interaction length.

Figure 3: (a): Force-distance measurements of LPS-SLBs with 10 wt% EY2 (), 10 wt% wild type (), 10 wt% S-LPS (), 20 wt% S-LPS () and POPC (). The force-distance curves are averaged and include 21, 67, 55, 52 and 206 single force curves. (b): Magnification of the compression before the lipid bilayer rupture in logarithmic scale for POPC, 10 wt% S-LPS and 20 wt% S-LPS.

LECTIN BINDING ON BACTERIA DISPLAYING THE HTYPE II EPITOPE FRAP measurements show that bound RSL on 50 wt% Htype II has similar lateral mobility as the lipids in the SLB. The diffusion coefficient of the FITC-RSL was calculated to be 0.6 ± 0.3 µm2/s with a recovered fraction of 83 ± 9 %. A FRAP series is shown in Figure S4 in the Supplementing information. The specificity of the RSL binding was confirmed by measurements on a pure POPC and on a 10 wt% EY2 SLB. No binding of the RSL-FITC on POPC SLBs was observed as expected due to the absence of appropriate binding molecules and the non-fouling properties of a completely covering POPC SLB38. EY2-LPS only differs from Htype II-LPS by a N14



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acetylglucosamine, a glucose and a fucose sugar. Figure 4Figure 4 shows a sketch of the


performed measurements with 50 wt% Htype II versus 10 wt% EY2 with biotinylated-RSL and subsequently added FluoProbes647-labeled streptavidin. The samples were rinsed to remove labeled, unbound streptavidin from the solution. This procedure will also remove reversibly bound RSL-streptavidin complexes. The measurements were performed by observing two channels (λ = 488 nm and λ = 633 nm). One channel (λ = 488 nm) was used as reference to detect the reflection of the laser beam at the glass surface to estimate the focal plane in the case of missing fluorescence and the second channel was used at the emission of the fluorophore where λ=633 nm excited the FluoProbes647. Figure 4Figure 4 (a) and (c) shows the intensity distribution for Htype II and


EY2 in a z-stack measurement as a y-section (upper graphic) and as an averaged fluorescence intensity (lower graphic: x,y layer averaged to give one intensity value per z-value). The graph of the average intensity vs. z-distance of the microscopy stack is given in Figure 4Figure 4 (b) and (d) for Htype II and EY2 respectively indicating the averaged intensity per frame.

Figure 4: (a) The biotinylated lectins bind to the Htype II and the subsequent FluoProbes647 labeled streptavidin binds to the biotin as indicated in the sketch. The fluorescence channel gives a signal from the FluoProbes647 and

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the reflection channel from the glass surface. The y section (upper graphic) and the x,y averaged section (lower graphic) show two distinct layers representing fluorescence and reflection signal. (b) Intensity vs. z-distance of the lectin-binding control measurement of the reflection (dashed, ) and fluorescence (line, ) channel for 50 wt% Htype II showing two distinct peaks (reflection and fluorescence peak). (c) For a 10 wt% EY2 SLB no fluorescence is expected due to the absence of specific binding. The y section (upper graphic) and x,y averaged section (lower graphic) show one distinct peak representing the reflection signal. (d) Intensity vs. z-distance of the lectin-binding control measurement of the reflection (dashed, ) and fluorescence (line, ) channel for 10 wt% EY2 showing only one distinct peak (reflection). Note that the slight offset between the two channels is an artifact induced by the optics of the used microscope.

DISCUSSION SUPPORTED BILAYER FORMATION WITH E. COLI LPS The QCM-D curves in Figure 1Figure 1 (a) with 10 wt% E. coli LPS show formation kinetics


comparable with the pure POPC vesicles. Only 10 wt% of the wild type LPS features an extended formation time. Figure 1Figure 1 (b) shows the largest shifts in ∆f and ∆D for wild


type LPS, which is expected due to the longest sugar chains attached to the lipid A + inner core. While the LPS-SLBs clearly have higher |∆f| than pure POPC, the differences between the Rd mutant, LPS1 and EY2 are not significant. However, an overall trend to increasing |∆f| compared to pure POPC is visible. The dissipation shifts also adopt increasing values scaling with approximate molecular weight of the LPS, with the highest value for the wild type LPS of ∆D = (0.7 ± 0.2) 10-6. Figure 1Figure 1 (c) shows the trend of increasing |∆f| and |∆D| for increasing concentrations of Htype II LPS. Higher Htype II concentrations resulted in slower SLB formation and vesicles with 100 wt% Htype II LPS did not form a SLB in TBS with 2 mM Ca2+ (see Supplementing information Figure S5). This trend indicates a shielding or repulsive effect of the dense LPS layer. This agrees with our previously published observation that vesicles containing high 16



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concentrations of lipids with sterically repulsive polymer headgroups prevent liposome adsorption and SLB formation25b. The addition of molecules to the buffer solution (Ca2+, urea, EDTA) has been shown to be beneficial for SLB formation of negatively charged lipids36, 39, E. coli mixtures20 and PEGlipids25a. The improvement of the SLB formation is due to a change of the electrostatic properties (Ca2+) or the density of hydroxyl groups/pH (urea, EDTA) at the substrate surface. Figure 2Figure 2 (a) demonstrates improved SLB formation kinetics of 50 wt% Htype II with the additive urea (20 mM) compared to pure TBS, which is further improved with 2 mM Ca2+. For the SLB formation of 20 wt% S-LPS the addition of 2 mM Ca2+ is necessary as without Ca2+ no formation is observable in the inspected time frame. It has been found that Ca2+ increases the electron density near the inner core of the LPS40 and that Ca2+ results in a collapse of the O-antigen chains towards the core saccharide40a. Theoretic models further suggest that Ca2+ binds to the KDO groups in the inner core and stabilize the lipid bilayer41. The lipid A part of the LPS contains two negatively charged phosphate groups (PO4-) and depending on the structure of the inner core additional negative phosphate groups are added. Htype II has, e.g., a third phosphate group. A Ca2+-induced collapse of the saccharide unit could therefore also be responsible for improved SLB formation in addition to charge bridging. The molecular weight of the used E. coli LPS (≈ 2000 - 2700 Da) is 2.5 - 3.5 times higher than the one of POPC (760 Da). Due to the presence of six acyl chains, LPS has a larger surface area per lipid molecule than POPC, but the LPS acyl chains are in saturated form and also allow denser packing than the acyl chains of POPC42. Snyder et al.42 estimated the area per hydrocarbon chain in rough E. coli chemotypes to be 21 Å2 – 26 Å2 for the gel and liquidcrystalline phases respectively, resulting in a total area of 126 Å2 – 156 Å2. With an area of 60 Å2 for phospholipids43 this corresponds to a factor of 2 - 2.6 between the area per lipid 17



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molecule of LPS and POPC. For LPS with three times the molecular weight of POPC and an area per lipid molecule of 2 - 2.6 times that of POPC, a mass increase from 464 ng/cm2 (POPC) to 475 – 500 ng/cm2 (10 wt%) or 500 – 600 ng/cm2 (50 wt%) is predicted by a straightforward calculation. This estimation however ignores that LPS is expected to have coupled water which would result in further increase of the adsorbed mass, as previously shown for SLBs of PEGlipids25b. However, the estimates above based purely on the increase in lipid molecular weight agree reasonably well with the experimentally measured values. This indicates that (i) the coupled water is lower for LPS, (ii) the concentration of LPS in the SLB is lower than assumed or (iii) a combination of both. The slightly increased dissipation values for increasing weight percentages can also have two different origins: (a) ∆D increases due to the contribution of coupled water from the sugar chains since QCM-D measures the total mass* coupled to the oscillation of the quartz sensor; (b) ∆D increases due to an increasing number of intact vesicles on the surface for an increasing LPS concentration. Intact vesicles increase the dissipation as well as the absolute frequency shift due to a higher amount of coupled water while SLBs only contribute to the absolute frequency shift due to the rigidity and the lower amount of coupled water, as for instance observed for egg-PC vesicle adsorption on TiO244. Similarly, the dissipation values found for PEG-SLBs were shown to increase due to coupled water for increasing PEG-lipid concentrations resulting in more water coupled on the surface by extended PEG chains25. Intact vesicles on the surface, however, also yield a pronounced larger frequency shift and decrease in the recovered fraction in FRAP measurements. As indicated in Table 1 the frequency shifts changed only marginally with increased LPS concentration and the recovered fractions are

*

The total mass is the mass of the biomolecules plus the mass of the bound or dynamically coupled water. 18


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almost 100%. We therefore conclude that the slightly higher dissipation values predominantly originate from additional water coupled by the saccharide headgroups. The FRAP measurements using NBD-PC† as label show no clear trend within the different LPS species and the diffusion coefficients are all in the same order of magnitude. Only the Rd mutant shows a significantly lower diffusion coefficient. The Rd mutant possesses 6 acyl chains compared to only 2 for the other LPS species which have been de-O-acylated. Such a three-fold increase of the lipid anchor size could be responsible for the decreased lipid mobility21, 45. Figure 1Figure 1 (c) shows the final frequency and dissipation values from QCM-D


measurements from SLBs formed with 5, 10 and 20 wt% of S-LPS with TBS and 2 mM Ca2+. The formation kinetics shows a trend to longer formation time for higher S-LPS concentration, similar to the observations for PEG-SLBs25b. The formation of 20 wt% S-LPS only proceeds with added Ca2+. In comparison to the significant decrease in ∆f and increase in ∆D for increasing PEG-lipids in PEG-SLBs25 the increase in S-LPS concentration only results in a slight decrease of ∆f and increase in ∆D as shown in Figure 1Figure 1 (c). A significant additional mass contribution from the O-antigens and coupled water is expected for the S-LPS species compared to the E. coli species. The measured shifts in frequency and dissipation for SLPS are however small, suggesting that the incorporated oligosaccharides are on average smaller than 2000 and 5000 Da PEG chains to which the results can be compared25b, or that significantly less water is coupled per molecule of similar molecular weight. It has to be taken into account that LPS concentrations are in weight percentages and that the area per molecule is larger for LPS than for POPC. As seen in the SDS-PAGE results, the LPS from S. Typhimurium has O-antigens ranging up to a molecular weight of 100 – 150 kDa. We consider two extreme cases where all LPS that are † Note that the mobility of NBD-PC was measured, which corresponds to the mobility of the main lipid component and not to the one of LPS, which remains unknown through lack of direct labeling, but can be indirectly estimated to be similar from the lectin binding measurements.

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included into the SLB are (i) from the lower range of molecular weights, i.e. 10 kDa corresponding to a radius of gyration Rg of 2.5 nm and a concentration cLPS of 1,5 mol% or (ii) from the higher range of molecular weights, i.e. 100 kDa corresponding to Rg of 7.9 nm and cLPS of 0.15 mol% (see the Supporting information for calculation) These estimates suggest that in case (i) with 10 kDa the situation would be comparable with a 22 mol% DOPE-PEG(2) (mushroom) with an Rg of 3.8 nm or with a 1 mol% DOPE-PEG(5) (mushroom) with Rg of 6.5 nm. The respective QCM-D values of PEG-SLBs and LPS-SLBs are listed in Table 2Table 2. They indicate that the absolute frequency |∆f| and dissipation shifts


|∆D| of 21 mol% PEG-DOPE(2) and 1 mol% DOPE-PEG(5) are much higher than the ones from S-LPS.

For the case (ii) with 100 kDa the situation would be comparable with a 0.25 mol% DOPEPEG(5) (mushroom) with Rg of 6.5 nm. The QCM-D values in Table 2Table 2 show similar absolute frequency shifts |∆f| for 0.25 mol% DOPE-PEG(5) and S-LPS, but higher absolute dissipation shifts |∆D| for 0.25 mol% DOPE-PEG(5) than for S-LPS. The comparison with PEG-SLBs shows that the QCM-D response from S-LPS SLBs is lower than expected.

Table 2: Comparison of QCM-D results of DOPE-PEG(2) and DOPE-PEG(5) SLBs with results of S-LPS SLBs.

Species

Concentration

∆f

∆D

(Hz)

(10-6)

DOPE-PEG(2)

21 mol%

-34.31.8 ± 1.81

2.01.8 ± 0.12

DOPE-PEG(5)

0.25 mol%

-28.6 ± 0.6

1.2 ± 0.1

DOPE-PEG(5)

1 mol%

-35.4 ± 0.5

3.0 ± 0.1

S-LPS

20 wt%

-28.4 ± 0.6

0.7 ± 0.1

COMPRESSION OF S. TYPHIMURIUM LPS 20



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One of the conclusions from the QCM-D measurements, namely that SLBs with S-LPS have a low saccharide layer thickness, should be observable by force-distance measurements that directly measure the thickness of the SLBs. S-LPS SLBs and E. coli LPS-SLBs were compared to POPC-SLBs in Figure 3Figure 3 (a).


The onset of the force-distance curves of 10 wt% EY2 and wild type SLBs at approximately 5 nm did not show significant deviations from the pure POPC SLB. EY2 and wild type add 2 and 5 additional monosaccharides, respectively, to the inner core where the size of a single glucose molecule was estimated as 8.6 Å47. The S-LPS however possesses repeats of the O-antigen, which are expected to be much larger than the EY2 and wild type sugar chains. The onset of the repulsive interaction in the force-distance curve of 20 wt% S-LPS SLBs, however, is starting approximately 1 nm earlier than the onset for pure POPC, 10 wt% EY2, wild type and also 10 wt% S-LPS SLBs. This increase of the onset distance of the S-LPS is smaller than expected. Tong and McIntosh23 reported a thickness of 100% Rd and Ra LPS-SLB formed on positively charged polyethylenimine (PEI) of 7 and 9 nm respectively. Compared to this study, in our case the LPS are expected to have even longer sugar chains since the S-LPS have an O-antigen repeat (lipid A + inner core + outer core + (n × O-antigen) whereas the Ra (lipid A + inner core + outer core) and Rd (lipid A + inner core) do not have O-antigens. On the other hand, the SLPS SLB is less densely packed with 20 wt% S-LPS of heterogeneously distributed O-antigen lengths and 80 wt% POPC than the LPS-SLB assembled by Tong and McIntosh23. The small effect of the S-LPS seen in Figure 3Figure 3 agrees with the only slight change in ∆f and ∆D in


the QCM-D data for increasing S-LPS concentration. In Figure 5Figure 5 (a) we compare the force-distance curve of 20 wt% with a corresponding


PEG-SLB, 2 mol% DOPE-PEG(2). Figure 5Figure 5 (a) shows averaged force-distance curves


of POPC (), of 20 wt% S-LPS () and 2 mol% DOPE-PEG(2) () SLBs. S-LPS shows a

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slightly increased interaction distance than POPC, but compared to 2 mol% DOPE-PEG(2) this increase in distance is low.

Figure 5: (a) Comparison of LPS- with PEG- and POPC SLBs with averaged force-distance curves of POPC (, 165 curves), 20 wt% S-LPS (, 52 curves) and 2 mol% DOPE-PEG(2) (, 206 curves). (b): Schematic representation of force-distance curves in the case of PEG-SLBs (upper) and LPS-SLBs (lower sketch).

The force-distance curves of S-LPS SLBs and 2 mol% DOPE-PEG(2) SLBs show different shapes. For DOPE-PEG(2) the compression of the PEG-chains, as discussed by Kaufmann et al.25, results in a clear increase in the force up to a specific force Fcompression where the forcedistance curve adapts the shape of a pure POPC force-distance curve (see Figure 5 (b)). This behavior was assigned to the compression of the polymer tethered to the SLB and the jump to the POPC force-distance characteristic due to the escape of the PEG-lipids from below the AFM tip as they are squeezed out from the interface by the applied pressure. PEG is well soluble in water and adapts a highly hydrated state which makes the PEG-chain extend from the surface. The force-distance curve for S-LPS however does not display the same characteristics. A distinct force Fcompression cannot be assigned in the force-distance curve of S-LPS; rather the whole curve resembles the force-distance curve of POPC with a 1-nm increased interaction distance (see Figure 5 (b)). This difference suggests that the layer formed by the LPS is stiffer 22


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than the one formed by the PEG-lipopolymers. The presence of Ca2+ in the buffer and a collapse of the O-antigens of LPS as shown by Schneck et al.40a could lead to a stiffer saccharide layer compared to the highly hydrated PEG-lipopolymers. This would cause a shorter interaction length and force-distance curves that do not have the polymer-like characteristic compression seen for DOPE-PEG(2). 10 wt% S-LPS SLBs formed in TBS‡ and in TBS + 2 mM Ca2+, however, indicated no differences in the frequency and dissipation shifts. This makes it unlikely that Ca2+ alone is responsible for the findings of a low thickness and a low hydration of the LPS chains. SLB formation for 20 wt% S-LPS did not proceed in TBS without Ca2+, which makes it difficult to completely rule out the effect of Ca2+ which could also still be present in trace, but significant, amounts. Further measurements are necessary to explore the specific effect of Ca2+ on the formation and structure of S-LPS SLBs. Another effect contributing to the interaction length in the AFM data is the real concentration of S-LPS in the membrane and the length of the saccharide chain. Due to its native origin iIt is likelylikely that the purity of theconcentration of S-LPS powder is lower than the one from synthetic ally synthesized lipopolymers. It can for example contain large glycopolymers or glycopolymer aggregates. The S-LPS lipid weight originally mixed into the liposomes could therefore contain effectively less lipidLPS material and more such contaminants, which will be filtered out in the extrusion process. The re, resultingsult would be

in a lower S-LPS

concentration incorporated into the vesicles than expected from he original mass ratio. Further, and that S-LPS with shorter saccharide chains have a higher probability to be incorporated into the membrane due to steric hindrance as shown for PEG-SLBs. The concentration compared to the PEG-lipopolysaccharide is, as discussed above, also likely to be lower (in mol% notation). Note that DLS measurements of S-LPS vesicles did not indicate micelles in solution which makes it highly unlikely that S-LPS was excluded from the SLB formation process. ‡

TBS is not calcium free, but not enriched in calcium 23


Formatted: Superscript

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In conclusion, S-LPS do not show polymer-like compression behavior as observed for PEGlipids and do not result in long interaction distances as could have been expected from their high molecular weights. The incorporation of S-LPS in SLBs was proven by QCM-D measurements with the corresponding slow-down in the formation process which was improved by Ca2+ and DLS measurements confirmed incorporation of the LPS into liposomes by the absence of micelles in solution. However, the exact structure, concentration and distribution of the saccharides in the SLB remain unclear.

LECTIN BINDING ON BACTERIA MIMICS WITH HTYPE II The specific binding of RSL to the fucose sugar unit in Htype II gives direct proof that Htype II LPS-lipids are included into the SLB. Figure S4 in Supplementing information demonstrates that RSL-FITC can be photobleached on a fluorescently unlabeled SLB with 50 wt% of Htype II (50 wt% POPC). Note that RSL non-specifically bound to the substrate would not show fluorescence recovery after photobleaching since protein adsorbed directly to the substrate would not possess lateral mobility. The diffusion coefficient of the FITC-RSL of 0.6 ± 0.3 µm2/s is approximately half of the value of POPC lipids in Htype II (Table 1Table 1) and the


recovered fraction of 83 ± 9 % indicates a small fraction of immobile lectins. The FRAP measurements demonstrate binding of RSL to the Htype II SLB. The control measurements on pure POPC and 10 wt% EY2 (Figure 4Figure 4) demonstrate that this binding is also specific.


The two layers of fluorescence and reflected laser beam in Figure 4Figure 4 demonstrate the


presence of a fluorescent layer on the substrate originating from the bound RSL-streptavidin complex. The presence of only the reflection signal in Figure 4Figure 4 (b) proves the absence of a fluorescence layer and hence no bound RSL-streptavidin complex in the case of EY2 LPS. EY2 does not offer specific binding sites to the biotinylated lectins due to the absence of the fucose sugar unit. 24



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This demonstrates that in the case of EY2 no fluorescence signal was measured from the focal plane and hence lectin was not enriched as it was in the case of Htype II. The concentration of the ligand for lectin binding is important since the affinity of lectins to their saccharide ligands is low and multivalent binding is necessary48. Kostlanova et al.34 measured the affinity constant for RSL and found that the highest affinity amongst monosaccharides is for fucose with a Kd=2 × 10-6 M. They found an even higher affinity to oligosaccharides containing fucose bound to a galactose (Fuc(1→2)Gal), as present in our Htype II LPS, with a Kd= 2.5 × 10-7 M34. Further, they reported that the RSL in solution is a trimer where each monomer contains two binding sites for fucose, in total six binding sites for the trimer. Kostlanova et al.34 also showed that high molecular weight polysaccharides (> 10 kDa, xyloglycan with Fuc(1→2)Gal exposed extremities on a cellulose backbone) bound to RSL-loaded chips and remained bound even upon washing with buffer which they attributed to multivalent binding. 2-3 bound fucose units would result in a Kd of 6 × 10-14 M - 1 × 10-20 M which is comparable to biotin-avidin binding and therefore for practical purposes irreversible. The presented approach of LPS-SLBs with mobile lipids shows that for lectin binding assays the fluorescence signal can be explicitly distinguished from a control system attributing the presence of bound RSL to multivalent bindings. It was also shown, as expected, that the bound lectin overwhelmingly bound to mobile lipids and therefore remained laterally mobile on the membrane. The recovery of the fluorescence intensity in FRAP measurements and the specificity of the RSL binding to Htype II confirms that LPS-SLBs provide a means to test for specific lectin-LPS interactions. Furthermore it demonstrates the successful incorporation of LPS into a SLB via self-assembly and fusion of vesicles. Additionally, multivalent binding of RSL means a larger effective anchor in the membrane, which should lead to a lower observed diffusion coefficient. A comparison of the RSL FRAP data with that of the NBD-POPC shows that this indeed was the case. 25


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The demonstrated formation of SLBs including specific LPS opens up the possibility to use the numerous surface sensitive and highly quantitative real-time techniques established for SLB research for bacterial and/or saccharide-containing membranes respectively. In most used platforms49 the saccharide-protein interaction is measured with saccharides immobilized on the surface. The suggested platform constitutes a better model of the native membrane. Mobility is important to allow a redistribution of the LPS and make multivalent binding possible which is effectively required in biology for relevant outcomes due to the weak affinity of the proteins for their glycan ligands6b. The possibility to probe mechanisms building on multivalency, which in biology also accounts for adaptive value, dynamic binding and reversibility, is an important option that the presented systems offer.

CONCLUSIONS In this report the formation of LPS-SLBs with LPS from two different bacteria, E. coli and S. Typhimurium and the potential of these platforms for future applications were investigated. The different E. coli LPS vary in the outer core structure and have no O-antigens, whereas the S. Typhimurium LPS hold O-antigen repeats. It was shown that LPS-SLB formation with concentrations as high as 10 wt% was successful for all species. The use of additives in the buffer solution, especially Ca2+ due to the negative charges of the LPS, helped to increase the concentration of LPS that could be used. For Htype II from E. coli as much as 50 wt% could be included and 20 wt% of S-LPS from S. Typhimurium. Advantages of the presented approach compared to existing approaches23,

40

to form

supported LPS membranes are the ease of vesicle fusion for the formation of highly defect free SLBs in situ, the resulting natural fluidity, density and mechanical properties. The formation on surface sensitive sensor substrates will allow the investigation of the specificity, the binding

26


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kinetics and affinity constants that provide detailed information about e.g. multivalency, and which typically is not straightforward or even impossible to obtain with traditional techniques. Force-distance measurements showed a slight increase of the thickness of S-LPS SLBs of approximately 1 nm. The low LPS interaction observed in AFM force-distance measurements is in agreement with the low differences in the absolute frequency and dissipation shifts observed in QCM-D measurements compared to pure POPC. The collapse of the O-antigens in presence of Ca2+ could to some extent explain the low thickness found in AFM force-distance measurements, but even at low Ca2+ buffer content a close to non-existent polymer repulsion at the interface was recorded. Thus LPS-SLB formation containing LPS species with long saccharide chains (high molecular weights) raises further questions about the exact fraction of LPS which is/can be included into the SLB in relation to the LPS molecular weight, or alternatively of its conformation and extension from the membrane. Htype II contains an α-1,2-linked fucose, which is recognized by RSL. The binding of FITCRSL to LPS-SLBs with 50 wt% Htype II could be confirmed using fluorescence microscopy. It demonstrates that a weak specific affinity interaction was successfully detected where the fluidity of the membrane could play a crucial role to facilitate multivalent binding of temporally and weakly bound lectins with multiple binding sites by allowing binding to additional fucose units by lateral diffusion. It also demonstrated that a specific LPS species could not only be incorporated but remained fully accessible to binding by lectins.

1. Schindler, M.; Osborn, M. J.; Koppel, D. E., Lateral mobility in reconstituted membranes comparisons with diffusion in polymers. Nature 1980, 283 (5745), 346-350. 2. Ruiz, N.; Kahne, D.; Silhavy, T. J., Transport of lipopolysaccharide across the cell envelope: the long road of discovery. Nature Reviews Microbiology 2009, 7 (9), 677-683. 3. Nikaido, H., Prevention Of Drug Access To Bacterial Targets - Permeability Barriers And Active Efflux. Science 1994, 264 (5157), 382-388. 4. Raetz, C. R. H.; Reynolds, C. M.; Trent, M. S.; Bishop, R. E., Lipid a modification systems in gramnegative bacteria. In Annual Review of Biochemistry, 2007; Vol. 76, pp 295-329. 5. Helander, I. M.; Mamat, U.; Rietschel, E. T., Encyclopedia of Life Sciences: Lipopolysaccharides. John Wiley & Sons, Ltd.: 2001.

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6. (a) Gagneux, P.; Varki, A., Evolutionary considerations in relating oligosaccharide diversity to biological function. Glycobiology 1999, 9 (8), 747-755; (b) Paulson, J. C.; Blixt, O.; Collins, B. E., Sweet spots in functional glycomics. Nature Chemical Biology 2006, 2 (5), 238-248. 7. Morelli, L.; Poletti, L.; Lay, L., Carbohydrates and Immunology: Synthetic Oligosaccharide Antigens for Vaccine Formulation. European Journal of Organic Chemistry 2011, (29), 5723-5777. 8. Fukui, S.; Feizi, T.; Galustian, C.; Lawson, A. M.; Chai, W., Oligosaccharide microarrays for highthroughput detection and specificity assignments of carbohydrate-protein interactions. Nat Biotech 2002, 20 (10), 1011-1017. 9. Jayaraman, N., Multivalent ligand presentation as a central concept to study intricate carbohydrate-protein interactions. Chemical Society Reviews 2009, 38 (12), 3463-3483. 10. Schindler, M.; Osborn, M. J.; Koppel, D. E., Lateral diffusion of lipopolysaccharide in the outer membrane of Salmonella typhimurium. Nature 1980, 285 (5762), 261-263. 11. Edidin, M., Rotational and Translational Diffusion in Membranes. Annual Review of Biophysics and Bioengineering 1974, 3 (1), 179-201. 12. Murray, G. L.; Attridge, S. R.; Morona, R., Altering the Length of the Lipopolysaccharide O Antigen Has an Impact on the Interaction of Salmonella enterica Serovar Typhimurium with Macrophages and Complement. Journal of Bacteriology 2006, 188 (7), 2735-2739. 13. Sackmann, E., Supported membranes: scientific and practical applications. Science 1996, 271 (5245), 43-48. 14. Keller, C. A.; Glasmästar, K.; Zhdanov, V. P.; Kasemo, B., Formation of Supported Membranes from Vesicles. Physical Review Letters 2000, 84 (23), 5443-5446. 15. Engel, A.; Muller, D. J., Observing single biomolecules at work with the atomic force microscope. Nature Structural Biology 2000, 7 (9), 715-718. 16. Muller, D. J.; Engel, A., Strategies to prepare and characterize native membrane proteins and protein membranes by AFM. Current Opinion in Colloid & Interface Science 2008, 13 (5), 338-350. 17. Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E., Lipid Rafts Reconstituted in Model Membranes. Biophysical Journal 2001, 80 (3), 1417-1428. 18. Mann, D. A.; Kanai, M.; Maly, D. J.; Kiessling, L. L., Probing low affinity and multivalent interactions with surface plasmon resonance: Ligands for concanavalin A. Journal of the American Chemical Society 1998, 120 (41), 10575-10582. 19. McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A., Supported planar membranes in studies of cell-cell recognition in the immune system. Biochimica et Biophysica Acta (BBA) - Reviews on Biomembranes 1986, 864 (1), 95-106. 20. Merz, C.; Knoll, W.; Textor, M.; Reimhult, E., Formation of supported bacterial lipid membrane mimics. Biointerphases 2008, 3 (2), FA41-FA50. 21. Nomura, K.; Inaba, T.; Morigaki, K.; Brandenburg, K.; Seydel, U.; Kusumoto, S., Interaction of Lipopolysaccharide and Phospholipid in Mixed Membranes: Solid-State 31P-NMR Spectroscopic and Microscopic Investigations. Biophysical Journal 2008, 95 (3), 1226-1238. 22. (a) Nguyen, K. T.; Le Clair, S. V.; Ye, S.; Chen, Z., Molecular Interactions between Magainin 2 and Model Membranes in Situ. The Journal of Physical Chemistry B 2009, 113 (36), 12358-12363; (b) Nguyen, K. T.; Le Clair, S. V.; Ye, S.; Chen, Z., Orientation Determination of Protein Helical Secondary Structures Using Linear and Nonlinear Vibrational Spectroscopy. The Journal of Physical Chemistry B 2009, 113 (36), 12169-12180. 23. Tong, J.; McIntosh, T. J., Structure of Supported Bilayers Composed of Lipopolysaccharides and Bacterial Phospholipids: Raft Formation and Implications for Bacterial Resistance. Biophysical Journal 2004, 86 (6), 3759-3771. 24. Gutsmann, T.; Haberer, N.; Carroll, S. F.; Seydel, U.; Wiese, A., Interaction between lipopolysaccharide (LPS), LPS-binding protein (LBP), and planar membranes. Biological Chemistry 2001, 382 (3), 425-434. 25. (a) Kaufmann, S.; Borisov, O.; Textor, M.; Reimhult, E., Mechanical properties of mushroom and brush poly(ethylene glycol)-phospholipid membranes. Soft Matter 2011, 7 (19), 9267-9275; (b)

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Supported Lipopolysaccharide Bilayers - Langmuir (ACS Publications)

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