Single Lipid Bilayer Deposition on Polymer Surfaces Using Bicelles

Subscriber access provided by UNIV OF FINDLAY

Article

Single Lipid Bilayer Deposition on Polymer Surfaces using Bicelles Qasim Saleem, Zhenfu Zhang, Amy Petretic, Claudiu C Gradinaru, and Peter M. Macdonald Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.5b00042 • Publication Date (Web): 09 Feb 2015 Downloaded from http://pubs.acs.org on February 18, 2015

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Biomacromolecules is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Single Lipid Bilayer Deposition on Polymer Surfaces using Bicelles Qasim Saleem,1,2 (#) Zhenfu Zhang,2,3 (#) Amy Petretic,2 Claudiu C. Gradinaru2,3 (*) and Peter M. Macdonald 1,2 (*) Department of Chemistry1 and Department of Physics,3 University of Toronto and Department of Chemical and Physical Sciences,2 University of Toronto Mississauga 3359 Mississauga Road North, Mississauga, Ontario, Canada L5L 1C6 #

These authors contributed equally to this work.

*

Authors to whom correspondence should be addressed:

[email protected] [email protected]

ABSTRACT A lipid bilayer was deposited on a 3 µm diameter polystyrene (PS) bead via hydrophobic anchoring of bicelles containing oxyamine-bearing cholesteric moieties reacting with the aldehyde functionalized bead surface. Discoidal bicelles were formed by mixing dimyristoylphosphatidylcholine

(DMPC),

dihexanoylphosphatidylcholine

(DHPC),

dimyristoyltrimethylammonium propane (DMTAP), and the oxyamine terminated cholesterol derivative cholest-5-en-3β-oxy-oct-3,6-oxa-an-8-oxyamine (CHOLOA) in the molar ratio DMPC/DHCP/DMTAP/CHOLOA (1/0.5/0.01/0.05) in water. Upon exposure to aldehyde-bearing PS beads, a stable single lipid bilayer coating rapidly formed at the bead surface. Fluorescence recovery after photobleaching demonstrated that the deposited lipids fused into an 1 ACS Paragon Plus Environment

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 32

encapsulating lipid bilayer. Electrospray ionization mass spectrometry showed that the short chain lipid DHPC was entirely absent from the PS adherent lipid coating.

Fluorescence

quenching measurements proved that the coating was a single lipid bilayer. The bicelle coating method is thus simple and robust, can be modified to include membrane-associated species, and can be adapted to coat any number of different surfaces.

KEYWORDS Bicelles, lipid coating, oxime bioconjugation, polystyrene beads, FRAP.

INTRODUCTION Supported bilayer membranes (SBM) were introduced by McConnell and co-workers three decades ago,1,2 and are used now in an ever-widening range of applications.3–5 SBM consist of a phospholipid bilayer deposited onto a planar solid substrate such as glass or mica which provides enhanced mechanical stability while retaining the essential fluid properties of natural membranes. A polymer “cushion” layer may be placed between the substrate and the inner leaflet of the supported bilayer to provide an aqueous space between the two.6,7 Recently, interest has grown in lipid membranes supported on colloidal particles,8 since these have many potential applications as membrane models, in biomolecule screening, as drug delivery reservoirs, and as therapeutic vectors. Recently, lipid bilayers supported on silica beads have even proved useful in the isolation of functional presynaptic complexes.9,10 Lipid bilayer deposition onto colloidal particles typically involves adhesion of lipid bilayer vesicles onto the surface followed by fusion of individual vesicles. Adhesion is encouraged via

2 ACS Paragon Plus Environment

Page 3 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

electrostatic attraction,11-13 or bioconjugation,14–16 or hydrophobic attraction between vesicle and surface.17–20 Fusion into a planar bilayer is induced using some trigger such as Ca2+ addition,21–23 freeze-thaw,20,24 or dehydration-rehydration cycling.25 However, such fusion methods often leave intact liposomes adsorbed alongside or atop patches of fused lipid bilayer.26,27

A further problem is that the method of inducing fusion may compromise

membrane protein native structure and function. Most recently, a new method of lipid bilayer deposition at solid surfaces has been reported involving the use of bicelles.28–30 Bicelles, or bilayered micelles, are biomimetic model membranes consisting of mixtures of long-chain and short-chain amphiphiles. The long-chain amphiphiles assemble into a planar lipid bilayer stabilized at its edges by the short-chain amphiphiles. The planar bilayer region of a bicelle provides an excellent approximation of the natural membrane environment, so that bicelles have become increasingly popular in membrane protein structural studies using a variety of techniques, including NMR,31,32 EPR,33– 36

and X-ray diffraction.37 Bicelles are also used as membrane protein38 and pharmaceutical

delivery vehicles.39

Relative to liposomes, bicelles enjoy two important advantages as

precursors for lipid bilayer coatings. First, bicelle fabrication is simple and carried out under mild conditions conducive to retention of native membrane proteins structure and function. Second, bicelles in the form of small planar discs can adapt readily to rough or irregular surfaces. Here, we demonstrate that bicelles, when deposited at the surface of a polystyrene (PS) bead (3 µm diameter) and retained there by hydrophobic anchors, spontaneously fuse into a continuous, unilamellar lipid bilayer completely coating and encapsulating the PS bead. The


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 32

bicelles were composed of mixtures of the zwitterionic long-chain phospholipid 1,2-dimyristoylsn-glycero-3-phosphocholine (DMPC), the zwitterionic short-chain phospholipid 1,2-dihexanoylsn-glycero-3-phosphocholine (DHPC), the cationic long-chain amphiphile 1,2-dimyristoyl-3trimethylammonium-propane (DMTAP), and the oxyamine-bearing cholesterol derivative cholest-5-en-3β-oxy-oct-3,6-oxa-an-8-oxyamine

(CHOLOA).

The

structures

of

all

four

amphiphiles are shown in Fig. 1. The molar ratio of long-chain to short-chain species, q = (DMPC+DMTAP+CHOLOA)/DHPC ≈ 2.1 was chosen to provide small discoidal bicelles. The oxyamine moiety of CHOLOA is intended to react with aldehyde groups displayed at the surface of the PS bead and thus form a covalent oxime linkage to the cholesterol ring intercalated within the bicelle bilayer, thereby hydrophobically anchoring the bicelle to the bead surface. Using a combination of fluorescence recovery after photobleaching (FRAP), fluorescence quenching and mass spectrometry, we show that simply mixing such bicelles with PS beads at a temperature above the gel-to-liquid-crystalline phase transition (TM) of DMPC sufficed to produce fusion of individual hydrophobically-anchored bicellar discs into a continuous, unilamellar lipid bilayer completely encapsulating the PS bead. Figure 1

EXPERIMENTAL Materials DMPC, DMTAP, DHPC, and NBD-PE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamineN-(7-nitro-2-1,3-benzoxadiazol-4-yl)) were purchased from Avanti Polar Lipids (Alabaster, AL). RhB-PE

(1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine

rhodamine

B

sulfonyl)) was purchased from Invitrogen (Carlsbad, CA). PS latex beads, surfactant free-


Page 5 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

ultraclean grade, 3 μm mean diameter, specific surface area 1.8x104 cm2/g, with a high density of surface grafted aldehyde (surface density 1/50 Å2) and sulfate charge groups (surface density 1/190 Å2), were purchased from Life Technologies (Burlington, ON) and used as received. All other reagents were purchased from Sigma Aldrich (Oakville, ON). CHOLOA synthesis Scheme 1 shows the synthetic route to cholest-5-en-3β-oxy-oct-3,6-oxa-an8-oxyamine (CHOLOA), commencing from cholest-5-en-3β-oxy-oct-3,6-oxa-an-8-ol (1), which was prepared as described by Davis and Szoka.40

The azidodicarboxylate-activated

intermediate (2) was prepared by first dissolving 1.3 g of (1), 1.4 g of triphenylphosphine (2 eq.) and 0.8 g of N-hydroxyphthalimide (2 eq.) in 45 ml of anhydrous THF. Upon full dissolution of the reactants under N2 a dilute solution of diisopropyl azodicarboxylate (1 ml in 8 ml of anhydrous THF) was added in drop wise fashion and allowed to react for 16 hours. After quenching by the addition of 20 ml of ethanol, the reaction mixture was concentrated by rotary evaporation, 70 ml of hexanes was added, the resulting precipitate of triphenylphosphine oxide (TPPO) was removed by filtration and the solution was concentrated again. The residue was treated multiple times with ethyl acetate (ETOAc) and hexanes to eliminate any traces of TPPO. The product was applied to a silica gel column using dichloromethane (DCM), and eluted with a 10-50% ETOAc:hexanes gradient. After concentration via rotary evaporation, the product was eluted a second time with a 25-50% ETOAc:hexanes gradient to yield 1.2 g (72% yield) of a gummy paste. 1H NMR (400 MHz, CDCl3): 7.84 (dd, J = 5.4, 3.1 Hz, 2H), 7.75 (dd, J = 5.4, 3.0 Hz, 2H), 5.38 – 5.28 (m, 1H), 4.38 (dd, J = 5.6, 3.4 Hz, 2H), 3.91 – 3.84 (m, 2H), 3.67 (dd, J = 5.8, 3.7 Hz, 2H), 3.61 – 3.51 (m, 6H), 3.23 – 3.06 (m, 1H), 2.39 – 2.13 (m, 2H), 2.05 – 1.75 (m, 5H), 1.61 – 1.24 (m, 15H), 1.19 – 0.83 (m, 22H), 0.67 (s, 3H). TLC (1:1 EtOAc:Hexanes) Rf = 0.4 (developed


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 32

with I2 or visualized on Fluorescent TLC plates). HRMS (ESI+): (M + CH3OH adduct + Na+) calculated, 718.47; found, 718.44 Scheme 1 CHOLOA (3) was prepared by reacting 1.2 g of (2) with 0.4 ml (5 eq.) of hydrazine monohydrate co-dissolved in 20 ml of dry DCM.

Although the expected phthalhydrazide

precipitate formed immediately, the reaction was allowed to proceed for 18 hours, after which the solution was filtered and diluted to 50 ml using chloroform. The organic phase was washed 5x with water, followed by a single wash with brine, and was subsequently dried using MgSO4. After filtration and evaporation of the solvent, 0.7 g (77% yield) of a gum was obtained.

1

H

NMR (400 MHz, CDCl3): 5.52 (s, 2H), 5.38 – 5.27 (m, 1H), 3.92 – 3.80 (m, 2H), 3.72 – 3.61 (m, 9H), 3.26 – 3.11 (m, 1H), 2.42 – 2.15 (m, 2H), 2.05 – 1.75 (m, 6H), 1.61 – 1.24 (m, 13H), 1.19 – 0.83 (m, 22H), 0.67 (s, 3H). HRMS (ESI+): (M + Na+) calculated, 556.43; found, 556.42. Preparation of Discoidal Bicelles

Bicelles were prepared with a molar composition

DMPC/DMTAP/CHOLOA/DHPC = 1.0/0.01/0.05/0.5, producing a long-chain/short chain molar ratio q = (DMPC+DMTAP+CHOLOA)/DHPC ≈ 2.1, which is expected to yield discoidal bicelles with diameters in the region of 200 Å.41 Appropriate quantities of the individual amphiphiles were co-dissolved in chloroform and the solvent was removed under a stream of nitrogen gas. For FRAP measurements 0.05 mol% RhB-PE was included, while for fluorescence quenching experiments 0.10 mol% NBD-PE was incorporated. The resulting lipid film was placed in a desiccator under vacuum overnight to remove final traces of solvent. The lipid film was hydrated with 1 mL milliQ water to produce a lipid concentration of CL = 1.31% w/v, and the resulting lipid suspension was subjected to four freeze-thaw cycles wherein the hydrated lipids


Page 7 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

were frozen in liquid N2, thawed in a 40°C water bath, and vortexed. The bicelle preparation was then stored at 4°C until use. Preparation of Unilamellar Liposomes Unilamellar lipid vesicles of molar composition DMPC/DMTAP/CHOLOA = 1.0/0.01/0.05 (note the absence of DHPC) were prepared as described above for bicelles, except that the multilamellar vesicles formed upon hydrating the lipid film were extruded eleven times through a 50 nm pore-size polycarbonate membrane at 37°C to produce unilamellar vesicles. Coating of Polystyrene Beads with Bicelles or Vesicles

Typically 1.0 mg of PS beads were

added to 2.0 mg of lipid, either bicelles or liposomes, and diluted to 2 mL with milliQ water. This represents an approximately 200-fold excess over the amount of lipid estimated to be required to coat the PS beads with a single lipid bilayer. Details of this calculation are provided in the Supplementary Information (SI). The PS bead+lipid mixture was gently swirled for 4 hours at 30°C. Excess lipid was removed by low-speed centrifugation (3 min, 6000 rpm) to pellet the lipid coated beads, which were then resuspended in 1 mL of water. This washing procedure was repeated 3-5 times. Fluorescence Imaging and FRAP Measurements

Wide-field

fluorescence

images

were

obtained using a custom-built Total internal reflection fluorescence (TIRF) microscope.42 Confocal fluorescence imaging and FRAP measurements were carried out using a custom-built confocal fluorescence microscope.20,43 Lipid coated PS beads, labelled with 0.05 mol% RhB-PE, were incubated on a plasma-cleaned coverslip for 10 minutes at 20°C. Confocal images of the sample were obtained at 20°C using laser excitation at 532 nm with an intensity of 1 W/cm2. For FRAP experiments, the laser intensity was set to 3 kW/cm2 for 0.5 seconds in order to


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

photobleach the lipids within a diffraction-limited spot on the surface of the PS beads. The photobleached area constituted around 4% of the total surface area of a lipid coated PS bead. Subsequent images were obtained by re-scanning the entire bead with a laser intensity of 1 W/cm2. To obtain high-resolution fluorescence recovery curves, the photobleached spot was monitored immediately after the photobleaching using a laser intensity of a 1 W/cm2 and 10 ms binning. The operation sequence was realized by an automatic filter wheel (Pacific Scientific, Model No. 5240) controlled by a custom-written Labview program. In order to reduce the impact of additional photobleaching, a time lapse acquisition scheme was employed, with 1 second sampled every 5 seconds at time delays longer than 1 minute after the photobleaching event. More details about the time-lapse data acquisition scheme are provided in the SI. FRAP measurements were repeated on several individual beads and produced essentially identical recovery curves, as shown in Fig. S2. To extract diffusion coefficients from measured FRAP curves, a Monte Carlo simulation of the random Brownian motion of fluorophores on the surface of a sphere was used as previously described,20 and as detailed further in the SI. Briefly, the simulation uses as input parameters two lateral diffusion coefficients (“fast” and “slow”) and their relative populations, and so generates a FRAP curve with a specified time resolution and duration. The input diffusion coefficients and relative populations were varied iteratively to obtain the best match to the experimental FRAP curves. Details regarding the parameter search procedure are given in the SI.


Page 8 of 32

Page 9 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

For comparative purposes with the three-dimensional geometry of the sphere, an analytical expression was derived describing two-dimensional diffusion on a circular disc into a circular photobleached spot. Details of the derivation are provided in the SI. Fluorescence Quenching Assay using Sodium Dithionite

A

sodium

dithionite

NBD-PE

quenching assay was performed on lipid-coated beads, all incorporating 0.10 mol% NBD-PE. A 1 M sodium dithionite, 1 M Tris solution was prepared immediately prior to use. Using a QuantaMaster PTI spectrofluorimeter (Photon Technology International, Lawrenceville, NJ) equipped with a Quantum Northwest TC 125 temperature controller (Liberty Lake, WA), the excitation was set to 470 nm and the emission was monitored at 532 nm using a time based scan via the FelixGX software. An aliquot of sample was diluted to 2 ml using MilliQ water in a 4 mL quartz cuvette and allowed to thermally equilibrate to a set temperature for at least 10 minutes. The emission was then monitored for several minutes at the rate of 1 point/sec. After obtaining a stable fluorescence reading for 1 minute, 20 μL of the sodium dithionite solution was added. The fluorescence emission intensity was monitored until a new baseline was achieved, following which 20 μL of 5 wt% Triton X-100 solution (Triton) was added to disrupt the lipid bilayer and quench the fluorescence entirely. Electrospray Ionization Mass Spectrometry (ESI-MS)

The lipid composition of the PS bead

bicelle lipid coating was analyzed via ESI-MS, and compared to whole bicelles. Bicelle lipid coated PS beads were stripped of bound lipids by immersion in hexane/isopropanol (2/1), the naked beads were removed by centrifugation, and the supernatant was diluted with methanol prior to injection of an aliquot into the mass spectrometer. Whole bicelles were treated identically. Using a syringe pump, samples were introduced into a Micromass ZQ single


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 32

quadruple electrospray ionization mass spectrometer (Waters Corporation, MA) operating in the positive ion mode at a flow rate of 30 μL/min with a capillary charge of +3.1 kV, cone voltage of 20 V and a source temperature of 100 °C. The samples were scanned over the range of 400–700 m/z with a scan acquired every second for 1 min.

RESULTS AND DISCUSSION Chemoselective Bicelle Binding to Polystyrene Beads

With the objective of depositing a

lipid bilayer onto the surface of a spherical PS bead, we assembled bicelles containing an oxyamine-derivatized cholesterol moiety, CHOLOA, targeted to react with aldehyde groups present at the PS bead surface, as outlined in Fig. 2. The oxime linkages formed by reaction between the oxyamine and the aldehyde functional groups are an example of chemoselective ligation, referring to the formation of bonds exclusively between particular pairs of functional reactants.44 The oxime linkage, in particular, is chemically stable and forms rapidly under mild conditions appropriate to biologically-relevant applications.45

The cholesterol moiety of

CHOLOA intercalates between the phospholipids of the bicellar bilayer, ensuring that the oxyamine group remains anchored to the bicelle surface. The ethylene oxide spacer ensures that the oxyamine functional group is minimally sterically restricted and able to encounter and react with the PS bead surface aldehydes. The presence of the cationic amphiphile DMTAP within the bicelles encourages their close apposition with the anionic PS surface via Coulombic attraction to the sulfate groups present there. However, electrostatic attraction alone is not sufficient to ensure a robust lipid coating. But once the oxime link forms, the cholesterol


Page 11 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

moiety anchored in the bicelle’s hydrophobic interior ensures that the bicelles remain attached at the PS bead surface. Figure 2 Fig. 3A shows a representative wide-field fluorescence image of PS beads coated with bicelles containing 0.05 mol% RhB-PE. The cross-section of fluorescence intensity through one such bicelle-coated PS bead is shown in Fig. 3B. The bead diameter is around 3 µm, as expected. It is evident that extensive lipid binding has been achieved, even after numerous washing cycles to remove unbound excess bicelles, and that lipid has deposited exclusively at the surface of the PS beads. The near homogenous density of fluorescence intensity suggests that defects larger than the diffraction limit (184 nm) do not exist in the deposited layer and that the surface is uniformly coated. However, the resolution limits in such images do not permit one to draw conclusions regarding the thickness of the lipid coating, i.e., single or multiple bilayer, or whether individual bicelles have fused into a continuous bilayer and formed a complete permeability barrier. Nor can it be assumed that the composition of bound amphiphiles is the same as that of the added bicelles. Figure 3 DHPC is Absent from the PS Bead Lipid Coating

To examine qualitatively the composition of

the lipids coating the PS beads, ESI-MS analysis was undertaken. ESI-MS is rapid, sensitive, and readily discriminates species having different molecular weights, such as DHPC and DMPC,46, 47 but rather similar 31P and 1H NMR chemical shifts. Result obtained with control bicelles are shown in Fig. 4A, versus lipids stripped from the PS bead coating in Fig. 4B. Since the ESI-MS instrumental response increases with decreasing phospholipid acyl chain length,46 DHPC is


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 32

detected with far greater sensitivity than either DMPC, or DMTAP, or CHOLOA. (Control measurements on DMPC/DHPC mixtures of known composition confirm this to be the case, data not shown). In Fig. 4A, specifically, the ESI mass spectrum of bicelles of molar composition DMPC/DMTAP/CHOLOA/DHPC = 1.0/0.01/0.05/0.5, q ≈ 2.1, shows DHPC peaks at 454.26 m/z [MDHPC + H+] and 476.23 m/z [MDHPC + Na+] that appear with far greater intensity than those due to DMPC (678.46 m/z [MDMPC + H+], 700.43 m/z [MDMPC + Na+]) or CHOLOA (MCHOLOA+CH3OH+Na+=588.29 m/z). DMTAP (554.49 m/z), being present at only 1% relative to

DMPC, is barely detected. In the ESI mass spectrum of the lipids bound to, and then stripped from, the PS bead surface, shown in Fig. 4B, peaks corresponding to DHPC are conspicuously absent, and only DMPC and DMTAP are evident. Thus, DHPC appears to have been removed preferentially during the various washing stages to which the PS bead lipid coatings were subjected during fabrication. Morigaki et al.30 in their studies of POPC/DHPC bicelles coating glass surfaces also suggested that DHPC was absent from the deposited bilayer. It is noteworthy that no CHOLOA peak is evident in the ESI mass spectrum of lipids stripped from the PS bead lipid coating (Fig. 4B), implying that all CHOLOA present in the bound lipid fraction had reacted with surface aldehydes and become covalently attached through the oxime linkage. The PS bead surface density of aldehydes is ~50 Å2/CHO. The bicelle surface density of CHOLOA, assuming 60 Å2 per DMPC and given 5 mol% CHOLOA relative to DMPC, is roughly 1200 Å2/CHOLOA. Thus, it is reasonable that all CHOLOA within a bicellar disc bound at the PS bead surface had become covalently attached and, hence, could not be removed merely by stripping with organic solvent.


Page 13 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

The DMTAP peak in Fig. 4B is notably intense, particularly so in comparison to Fig. 4A, implying that DMTAP is enriched at the PS bead surface relative to the 1 mol% DMTAP present in the initial bicelle formulation. The PS bead surface density of sulfates is ~190 Å2/SO4 which is roughly a factor of 30 greater anionic charge density than the bicelle cationic surface density due to 1 mol% DMTAP. Bicelles are highly fluctional, in that there is a constant dynamic exchange of lipids between individual discoidal fragments, as established from fluorescence studies.48 Thus it is entirely plausible, given the higher surface charge density of the PS beads versus the bicellar discs plus the fluctional nature of bicelles, that DMTAP becomes enriched at the PS bead surface via exchange under the motive force of Coulombic attraction. Because ESIMS as shown in Figs. 4A and 4B is not strictly quantitative without extensive calibration efforts, further direct quantitative measurements will be necessary to address this possibility. Figure 4 Bicelles Bound at the PS Bead Surface Fuse into a Continuous Lipid Bilayer

To address the

question of whether the bicelles bound at the PS bead surface fuse into a continuous lipid bilayer, FRAP measurements were performed at room temperature (20°C). Fig. 5 shows the results obtained for the case of bicelles allowed to bind PS beads at 30°C, i.e., above the TM of DMPC. Prior to photobleaching (Fig. 5A), the RhB-PE fluorescence is a continuous ring arising from lipid bound at the PS bead surface. Immediately after photobleaching a diffraction-limited spot, the ring of fluorescence is discontinuous (Fig. 5B), but it reacquires the continuous shape on a time scale of 10-20 minutes (Fig. 5C). This can only occur if individual bicelles have fused and formed a continuous lipid bilayer (or bilayers) on the surface of the PS bead. A movie showing the time course of the recovery of the continuity of the ring of fluorescence was made


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 32

by stringing together a sequential series of confocal images of a photobleached PS bead and is provided in the SI. Figure 5 To quantify the lateral diffusion of lipids bound to the PS bead surface, FRAP curves were measured and analyzed using Monte Carlo simulations to extract lateral diffusion coefficients. Fig. 6 shows the experimental FRAP curve obtained at 20°C from a single PS bead bicelle lipid coating prepared at 30°C, along with the corresponding best-fit Monte Carlo simulation. Essentially identical recoveries were obtained when different beads were samples, as detailed in Fig. S2. Lateral diffusion coefficients and relative populations obtained from the Monte Carlo simulations are listed in Table 1. The simulation indicates the presence of two populations, virtually equal in size, but differing by several orders of magnitude in their lateral diffusivity. Simulations assuming a single population of diffusing lipids always produced inferior fits to the experimental recovery curves. Comparisons of several single population versus twopopulation simulations are provided in Fig. S3. Figure 6 The lateral diffusion coefficient of the “fast” fraction (D = 1x10-14 m2s-1) is in accord with values for DMPC lipid bilayers at temperatures below TM = 24°C.49

As for the “slow”

population, supported bilayer membranes generally exhibit two roughly equal lipid populations with differing lateral diffusion properties and this is usually attributed to friction between the inner bilayer leaflet and the underlying support that is absent from the outer leaflet.50,51 Even if the bilayer is separated from the support by spacer groups intended to alleviate such friction,


Page 15 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

the anchors are immobile and can introduce a “picket fence”-type frictional barrier within the inner leaflet.19,20 The mobile fraction of lipids in the supported bilayer m (Table 1) is extracted by applying Eqs. (1) and (2) to the raw FRAP data: 1 − 1 − + 1 − = 0,

(1)

1 − 1 − + 1 − + 1 − = +∞,

(2)

where 0 ≤ ≤ 1 is the degree of photobleaching in the photobleached area, and is the fraction of total fluorophores photobleached. F(0) and F(+∞) are the intensities measured immediately after the photobleaching event (average of first 10 data points) and at the end of the recovery (average of last 30 data points), respectively, both normalized relative to the prephotobleaching intensity −∞. For bicelle-coated PS beads, the mobile lipid fraction is on the order of 70% (Table1). This must be considered a lower limit, since an inspection of Fig. 6 shows that the fluorescence continues to recovery slowly out to long times. In order to examine the possibility that the biphasic FRAP curves were merely the result of a single population diffusing across the three-dimensional geometry of the PS beads, as opposed to two distinct populations, a two-dimensional system was modelled, consisting of a disk of finite radius with a small photobleached spot. The derivation of the analytical formula describing FRAP curves in this instance is provided in the SI (Eq. S3). Fig. 6 shows the fluorescence recovery in the two-dimensional disk case using the same diffusion coefficients and populations obtained from the Monte Carlo simulation for the three-dimensional sphere. The comparison shows that while FRAP in the “fast” phase is similar in the two cases, recovery


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 32

in the “slow” phase is enhanced in the three-dimensional case. This is likely due to the more complex geometry of the photobleached region in the three-dimensional case,20 relative to the strictly circular photobleached spot assumed for the two-dimensional case. Nevertheless, this comparison makes clear that the dimensionality of the PS bead is not the origin of the biphasic FRAP curves, but rather, that there are indeed two different diffusing populations. Permeability of PS Bead Lipid Bilayers

To examine the integrity of the lipid bilayer coating,

NBD-PE fluorescence quenching by sodium dithionite was examined. Ideally, dithionite ions added externally will not permeate to the interior compartment of an intact unilamellar spherical lipid bilayer, and the NBD-PE fluorescence would be reduced to 50% of the initial value. When PS bead lipid coatings formed from bicelles were interrogated in this fashion, as shown in Fig. 7, NBD-PE fluorescence was reduced to roughly 30%, indicating significant permeation of dithionite to the interior side of the lipid bilayer coating. Figure 7 Increased permeability is a general concern with SBM and perturbations by the underlying supporting surface are often implicated as the cause. For instance, Nollert et al.52 found that an intact POPC lipid bilayer deposited on a hard glass surface (planar or spherical) was freely permeable to dithionite ions, an effect attributed to surface roughness. Similarly, Ng et al.18 found that coating soft hydrophobically-modified dimethacrylamide beads with eggPC resulted in a continuous bilayer that was almost completely permeable to cobalt ions, in contrast to the impermeability of egg PC liposomes.

It was proposed that the stress

experienced by the bilayer when in close proximity to the dynamic, possibly highly corrugated, surface of the bead was the origin of the increased permeability.


Page 17 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Comparison with Liposome-Based PS Bead Coatings

Because

liposomes

are

most

commonly used to deposit lipid bilayer coatings on solid supports, comparative studies were undertaken for the case of PS bead lipid coatings formed with CHOLOA-containing liposomes. Fluorescence images were largely indistinguishable from those obtained with bicelle-based coatings (not shown). At the level of resolution achievable with the technique used it could not be determined from such images whether individual intact liposomes were bound at the PS bead surface, or if fusion into regions of planar bilayer had occurred, or some mixture of the two. However, FRAP recovery curves, as shown in Fig. 6, were characterized by a large immobile fraction and much slower apparent lateral diffusion coefficients than the corresponding bicelle-based coatings, as detailed in Table 1.

Furthermore, fluorescence

quenching measurements, shown in Fig. 7, revealed fluorescence reduced to about 60% of the initial value, in line with reductions measured on the initial liposomes. Given these FRAP and fluorescence quenching results, it would seem that, in the absence of any external trigger, fusion of the liposomes bound at the PS bead surface was incomplete. This might be the result of electrostatic repulsion between adjacent cationic liposomes. Mechanism of Single Bilayer Formation from Bicelles Bound at the PS Bead Surface The mechanism by which a single bilayer coating forms at the PS bead surface upon deposition of bicelles, as deduced from the experiments described here, is summarized schematically in Fig. 2. The initial interaction is electrostatic attraction from a distance between cationic bicelles and the anionic PS bead surface. Upon close approach, the oxyamine groups displayed at the bicelle surface are able to chemoselectively react with aldehyde groups displayed at the PS bead surface, forming oxime linkages which anchor the bicelles via the hydrophobically-


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 32

intercalated cholesterol ring. The neutral, relatively water-soluble short-chain species DHPC is preferentially removed during the washing steps to remove excess lipid. Elimination of DHPC forces fusion of bicelles into larger, unilamellar structures, as the self-assemblies seek to minimize edge regions formerly occupied by DHPC. Provided a sufficient density of surfacebound bicelles had been attained initially, fusion occurs and begets a continuous unilamellar lipid bilayer completely encasing, and closely conforming to, the PS bead surface.

CONCLUSIONS A novel method has been described for the deposition of a single lipid bilayer onto a hard polymer bead starting from discoidal bicelles and using chemoselective chemistry to hydrophobically anchor the lipid assemblies. This method of lipid bilayer deposition is of general relevance for coating both hard and soft matter systems, extending the established use of bicelles in coating silicon28,29 and lipidic30 surfaces. Since discoidal bicelles can be made with a variety of saturated and unsaturated phospholipids, various lipid bilayer coating compositions can be achieved. Relative to conditions required for liposome fusion into a continuous lipid bilayer, bicelle fusion is induced simply and spontaneously under mild conditions. And because bicelles can be composed to optimize membrane protein native structure and function, this approach should prove advantageous in depositing membrane proteins at such surfaces for analytical, diagnostic or therapeutic applications.

SUPPORTING INFORMATION AVAILABLE


Page 19 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Calculation of polystyrene bead – bicelle surface ratios, comparison of continuous and timelapse FRAP data acquisition modes, the reproducibility of the lipid bilayer coating on PS beads, Monte-Carlo FRAP simulation, 2D vs. 3D FRAP recovery analysis, 1H NMR spectra of compounds and a gif movie showing the time course of the recovery of fluorescence recovery. This material is available free of charge via the Internet at http://pubs.acs.org.

ACKNOWLEDGEMENTS Financial support from the Natural Science and Engineering Research Council (NSERC) of Canada is acknowledged (P.M.M and C.C.G). Q.S. was supported by an Ontario Graduate Studies (OGS) Scholarship and Z.Z. was supported by a Canada Institutes of Health Research (CIHR) Training Grant. We would like to thank University of Toronto Profs. Ulrich Krull, David McMillen and Heiko Heerklotz for kind access to instruments.

REFERENCES (1)

Brian, A. A.; McConnell, H. M. Proc. Natl. Acad. Sci. 1984, 81, 6159–6163.

(2)

Tamm, L.; McConnell, H. Biophys. J. 1985, 47, 105–113.

(3)

Sackmann, E. Science 1996, 271, 43–48.

(4)

Tanaka, M.; Sackmann, E. Nature 2005, 437, 656–663.

(5)

Castellana, E. T.; Cremer, P. S. Surf. Sci. Rep. 2007, 38, 429–444.

(6)

Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667–

1671. (7)

Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400–1414.


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(8)

Troutier, A.; Ladavière, C. Adv. Colloid Interface Sci. 2007, 133, 1–21.

(9)

Gopalakrishnan, G.; Thostrup, P.; Rouiller, I.; Lucido, A. L.; Belkaïd, W.; Colman, D. R.;

Lennox, R. B. ACS Chem. Neurosci. 2009, 1, 86–94. (10) Lucido, A. L.; Sanchez, F. S.; Thostrup, P.; Kwiatkowski, A. V.; Leal-Ortiz, S.; Gopalakrishnan, G.; Liazoghli, D.; Belkaid, W.; Lennox, R. B.; Grutter, P.; Garner, C. C.; Colman, D. R. J. Neurosci. 2009, 29, 12449–12466. (11) Katagiri, K.; Caruso, F. Adv. Mater. 2005, 17, 738–743. (12) De Geest, B. G.; Stubbe, B. G.; Jonas, A. M.; Van Thienen, T.; Hinrichs, W. L. J.; Demeester, J.; De Smedt, S. C. Biomacromolecules 2006, 7, 373–379. (13) Helwa, Y.; Dave, N.; Liu, J. Soft Matter 2013, 9, 6151–6158. (14) MacKinnon, N.; Guérin, G.; Liu, B.; Gradinaru, C. C.; Macdonald, P. M. Langmuir 2009, 25, 9413–9423. (15) El-Hamed, F.; Dave, N.; Liu, J. Nanotechnology 2011, 22. (16) Gopalakrishnan, G.; Rouiller, I.; Colman, D. R.; Lennox, R. B. Langmuir 2009, 25, 5455– 5458. (17) Jin, T.; Pennefather, P.; Lee, P. I. FEBS Lett. 1996, 397, 70–74. (18) Ng, C. C.; Cheng, Y.; Pennefather, P. S. Biophys. J. 2004, 87, 323–331. (19) MacKinnon, N.; Guérin, G.; Liu, B.; Gradinaru, C. C.; Rubinstein, J. L.; Macdonald, P. M. Langmuir 2010, 26, 1081–1089. (20) Saleem, Q.; Liu, B.; Gradinaru, C. C.; Macdonald, P. M. Biomacromolecules 2011, 12, 2364–2374.


Page 20 of 32

Page 21 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(21) Wilschut, J.; Duzgunes, N.; Hoekstra, D.; Papahadjopoulos, D. Biochemistry 1985, 24, 8– 14. (22) Csúcs, G.; Ramsden, J. J. Biochim. Biophys. Acta, Biomembr. 1998, 1369, 61–70. (23) Seantier, B.; Kasemo, B. Langmuir 2009, 25, 5767–5772. (24) Sugihara, K.; Jang, B.; Schneider, M.; Vörös, J.; Zambelli, T. Soft Matter 2012, 8, 5525– 5531. (25) Kaufmann, M.; Jia, Y.; Werner, C.; Pompe, T. Langmuir 2011, 27, 513–516. (26) Majewski, J.; Wong, J. Y.; Park, C. K.; Seitz, M.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1998, 75, 2363–2367. (27) Reviakine, I.; Brisson, A. Langmuir 2000, 16, 1806–1815. (28) Zeineldin, R.; Last, J. A.; Slade, A. L.; Ista, L. K.; Bisong, P.; O’Brien, M. J.; Brueck, S. R. J.; Sasaki, D. Y.; Lopez, G. P. Langmuir 2006, 22, 8163–8168. (29) Tabaei, S. R.; Jönsson, P.; Brändén, M.; Höök, F. J. Struct. Biol. 2009, 168, 200–206. (30) Morigaki, K.; Kimura, S.; Okada, K.; Kawasaki, T.; Kawasaki, K. Langmuir 2012, 28, 9649– 9655. (31) Sanders, C. R.; Hare, B. J.; Howard, K. P.; Prestegard, J. H. Prog. Nucl. Magn. Reson. Spectrosc. 1994, 26, Part 5, 421–444. (32) Marcotte, I.; Auger, M. Concepts Magn. Reson., Part A 2005, 24A, 17–37. (33) Cardon, T. B.; Tiburu, E. K.; Lorigan, G. A. J. Magn. Reson. 2003, 161, 77–90. (34) Dave, P. C.; Nusair, N. A.; Inbaraj, J. J.; Lorigan, G. A. Biochim. Biophys. Acta, Biomembr. 2005, 1714, 141–151. (35) Nusair, N. A.; Lorigan, G. A. Chem. Phys. Lipids 2005, 133, 151–164.


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(36) Cardon, T. B.; Dave, P. C.; Lorigan, G. A. Langmuir 2005, 21, 4291–4298. (37) Ujwal, R.; Bowie, J. U. Methods 2011, 55, 337–341. (38) Kang, C.; Vanoye, C. G.; Welch, R. C.; Van Horn, W. D.; Sanders, C. R. Biochemistry 2010, 49, 653–655. (39) Barbosa-Barros, L.; Rodríguez, G.; Barba, C.; Cócera, M.; Rubio, L.; Estelrich, J.; LópezIglesias, C.; de la Maza, A.; López, O. Small 2012, 8, 807–818. (40) Davis, S. C.; Szoka, F. C. Bioconjugate Chem. 1998, 9, 783–792. (41) Lucyanna, B.-B.; Gelen, R.; Merce, C.; Laia, R.; Carmen, L.-I.; Alfons, D. la M.; Olga, L. Pharmaceutics 2011, 3, 636–664. (42) Saleem, Q.; Zhang, Z.; Gradinaru, C. C.; Macdonald, P. M. Langmuir 2013, 29, 14603– 14612. (43) Liu, B.; Mazouchi, A.; Gradinaru, C. C. J. Phys. Chem. B 2010, 114, 15191–15198. (44) Tiefenbrunn, T. K.; Dawson, P. E. Biopolymers 2010, 94, 95–106. (45) Jencks, W. P. J. Am. Chem. Soc. 1959, 81, 475–481. (46) Koivusalo, M.; Haimi, P.; Heikinheimo, L.; Kostiainen, R.; Somerharju, P. J. Lipid Res. 2001, 42, 663–672. (47)

Han, X.; Gross, R.W. Mass Spectr. Reviews 2005, 24, 367-412.

(48) Rowe, B. A.; Neal, S. L. Langmuir 2003, 19, 2039–2048. (49) Fahey, P. F.; Webb, W. W. Biochemistry 1978, 17, 3046–3053. (50) Hetzer, M.; Heinz, S.; Grage, S.; Bayerl, T. M. Langmuir 1998, 14, 982–984. (51) Scomparin, C.; Lecuyer, S.; Ferreira, M.; Charitat, T.; Tinland, B. Eur. Phys. J. E 2009, 28, 211–220.


Page 22 of 32

Page 23 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(52) Nollert, P.; Kiefer, H.; Jahnig, F. Biophys. J. 1995, 69, 1447–1455.

Table 1.

Results of Monte Carlo Simulations of FRAP Curves of Lipids Bound to PS Beads D1 (*10-14m2/s)

D2 (*10-16m2/s)

P(D1):P(D2)

Mobile Fraction (%)

Bicelles

1.0

4.6

50:50

68 ± 3

Liposomes

0.40

1.4

40:60

46 ± 7

Lipid Morphology

FIGURE CAPTIONS Figure 1. Structures of DMPC, DHPC, DMTAP and CHOLOA. Figure 2. Schematic of bicelles binding to PS bead and their fusion to form a unilamellar lipid bilayer. Discoidal bicelles are composed of mixtures of DMPC (orange head groups) and DHPC (green head groups) with a small percentage of CHOLOA (red oblong) and DMTAP (not indicated). Initial attraction between bicelles (cationic) and PS beads (anionic) is electrostatic. Oxyamine groups of CHOLOA, displayed at the bicelle surface, react with aldehyde groups displayed at the PS bead surface to form oxime linkages, thereby anchoring the bicelles to the PS beads. DHPC is preferentially removed by washing, which forces fusion of adjacent bicelles into a continuous unilamellar lipid bilayer covering the PS bead. Figure 3. (a) Wide field fluorescence image of bicelle-coated PS beads containing 0.05 mol% RhB-PE and obtained at 20°C. The scale bar represents 5 μm. (b) Cross-section of fluorescence intensity across an individual bicelle-coated PS bead.


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure

4.

ESI

mass

spectrum

of

(a)

bicelles

Page 24 of 32

having

molar

composition

DMPC/DMTAP/CHOLOA/DHPC = 1.0/0.01/0.05/0.5, q=2.1, (b) Lipids stripped from PS beads after binding via oxyamine-aldehyde chemoselective formation of oxime linkages. Figure 5. FRAP images (measured at 20 °C) of 0.05 mol% RhB-PE-labelled PS bead bicelle lipid coatings formed at 30⁰C: prior to photobleaching (a), immediately after photobleaching (b), and 20 minutes subsequent to photobleaching (c). Figure 6. FRAP recovery curves (black), Monte Carlo simulations (red) and 2D analytical plot (blue) for 0.05 mol% RhB-PE-labelled PS bead bicelle lipid coatings. Simulations assumed two different lateral diffusion coefficients and population weightings. Results of best-fits for the lateral diffusion coefficients and populations are listed in Table 1. Parameters used in 2D analytical plot are identical to those used in the simulation. Also shown are FRAP recovery curves for PS bead lipid coatings formed from liposomes of molar composition DMPC/DMTAP/CHOLOA = 1.0/0.01/0.05. Figure 7. Sodium dithionite induced quenching of NBD-PE fluorescence from PS bead lipid coatings

formed

using

bicelles

(black

curve)

of

molar

composition

DMPC/DMTAP/CHOLOA/DHPC = 1.0/0.01/0.05/0.5, q≈2.1 and liposomes (gray curve) formed of molar composition DMPC/DMTAP/CHOLOA = 1.0/0.01/0.05. Measurements were performed at 15°C. Dithionite was added at time point “D” and the detergent Triton X100 was added at time point “T”.


Page 25 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 1


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Scheme 1 Reaction scheme outlining synthesis of the cholesterol bearing oxyamine linker.


Page 26 of 32

Page 27 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 2


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 3


Page 28 of 32

Page 29 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 4


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 5 30 ACS Paragon Plus Environment

Page 30 of 32

Page 31 of 32

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 6


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 7

TOC Figure


Page 32 of 32

Single Lipid Bilayer Deposition on Polymer Surfaces Using Bicelles

Recommend Documents