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Lipid layers on nanoscale surface topography: stability and effect on protein adsorption Irma Liascukiene, Karim El Kirat, Mathieu Beauvais, Svajus Joseph Asadauskas, Jean-Francois Lambert, and Jessem Landoulsi Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b00431 • Publication Date (Web): 05 Apr 2017 Downloaded from http://pubs.acs.org on April 7, 2017

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Lipid layers on nanoscale surface topography: stability and effect on protein adsorption Irma Liascukiene a,b,c,†, Karim El Kirat d, Mathieu Beauvais e, Svajus J. Asadauskas c, JeanFrançois Lambert a,b, Jessem Landoulsi a,b,*

a

Sorbonne Universités, UPMC Univ Paris 06, Laboratoire de Réactivité de Surface, F-75005, Paris, France b

c

CNRS, UMR 7197, Laboratoire de Réactivité de Surface, F-75005, Paris, France

State Research Institute Center for Physical Sciences and Technology, Saulėtekio al. 3, Vilnius, LT-10257, Lithuania

d

Laboratoire de Biomécanique &Bioingénierie, CNRS, UMR 7338, Université de Technologie de Compiègne, BP 20529, F-60205 Compiègne Cedex, France e

Alcatel-Lucent Bell Labs France, 7 route de Villejust, 91620 Nozay, France

* Corresponding author: [email protected] Tel. +33 1 44 27 52 91 Fax. +33 1 44 27 60 33

† Present address : LadHyX, Ecole Polytechnique, 91128 Palaiseau Cedex, France

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Abstract We report the coating of a surface with random nanoscale topography with a lipid film formed by an anchoring stearic acid (SA) monolayer and phospholipid (DPPC) layers. For this purpose, different procedures were used for phospholipid coating, including adsorption from solution, drop-deposition and spin-coating. Our results reveal that the morphology of the obtained lipid films is strongly influenced by the topography of the underlying substrate, but also impacted by other factors, including the coating procedure and surface wettability (modulated by the presence of SA). These coated surfaces showed a remarkable antifouling behaviour towards proteins with different yields of repellency (Yrp), depending on the amount/organization of DPPC on the nanostructured substrate. The interaction between proteins and phospholipids involves a partial detachement of the film. The use of characterization techniques with different charcateristics (accuracy, selectivity, analysis depth) did not reveal any obvious vertical heterogenity of the probed interface, indicating that the lipid film acts as a nonfouling coating on the whole surface, including the outermost part (nanoprotrusions) and in depth (valleys).


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1. Introduction The adsorption of biomolecules on planar surfaces with nanoscale topographies is a phenomenon with a common occurrence in diverse biomedical applications. The surfaces of biomaterials, typically metals, oxides and bioceramics, exhibit indeed a nanoscale roughness which is originally present (native), or deliberately obtained to achieve and/or improve specific properties. Extensive research has been conducted to design nanostructured functional biointerfaces with the aim to tailor the organization of biomolecules and their bioactivity.

1-4

Widespread strategies used to control rough surface topographies at the

nanoscale consist of physical processes based on lithography and physical vapor deposition techniques,

5-7

8

and self-assembly of

9

It appears that nanoscale roughness influences the adsorption behaviour of

7, 10-14

while microscale roughness impacts rheological properties of the cells and

nano-objects. proteins,

chemical processes, including etching, oxidation, etc,

their ability to adapt their contact area to the topographies.

15-20

In contact with biological

fluids, the adsorption of proteins is ubiquitous and strongly affects the function of biomaterials because cells actually interact with the protein layer rather than the raw surface of biomaterials.

21

On the one hand, some specific adsorbed proteins play a pivotal role to

mediate cell adhesion and other biological functions (molecular recognition, signalling, etc). 22

On the other hand, a non-specific adsorption of proteins may induce various biological

processes (blood coagulation, platelet activation and adhesion, leukocyte adhesion, etc), resulting in serious clinical issues. 21, 23 The scenario of protein adsorption is also determining for the colonization of biomaterials by microorganisms; the latter interact with proteins and other macromolecules previously adsorbed on the surface, leading to the formation of a biofilm. 24 The design of nonfouling surfaces with remarkable performances may be achieved if both surface chemistry and roughness are finely modulated. Most frequently, the surface of 3 ACS Paragon Plus Environment

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biomaterials is chemically modified with the aim to modify the physicochemical properties (hydrophobicity, electrical charge, solvation) which control the interactions with biomolecules and/or to retain biochemical entities. The functionalization of planar surfaces with nanoscale topographies presents two main challenges. First, the chemical schemes which motivated the procedures of surface modification are usually evaluated on flat rather than rough substrates with nanoscale topography. This question is crucial as many chemical schemes that work well on model flat surfaces are difficult to apply on nanostructured surfaces. Second, the characterization of such rough surfaces is a challenge, owing to the chemical heterogeneities (along and perpendicular to the surface plan) which may be induced by the nanoscale topography. To reduce nonspecific adsorption of proteins, zwitterionic function groups, have been used as an alternative to poly-ethylene glycol (PEG), which is the most studied synthetic nonfouling system

25

but presents important disadvantages related to the dependence of its conformation

on the temperature and its autoxidation.

26-27

The use of zwitterions to reduce nonspecific

protein adsorption was originally inspired by biological processes that occur on the external surface of the cell membrane of mammalian cells, the extracellular side of which is rich in phospholipids, especially phosphatidylcholine, while only few zwitterions are present on the cytoplasmic side.

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Different strategies have been developed to allow the grafting of

phospholipid on inorganic substrates, including the conjugation of phosphorylcholine with anchoring group (e.g. siloxanes polymers,

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29

or thiols,

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the synthesis of phosphorylcholine-based

etc. The use of natural phospholipids to design nonfouling surface is still scarce

however, probably due to the sensitivity of these molecules at interfaces towards physicochemical factors. In the present study, we report the chemical modification of a superficially nanostructured aluminum substrate with dipalmitoylphosphatidylcholine (DPPC) bearing zwitterion head 4 ACS Paragon Plus Environment

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group, with the aim of scrutinizing the impact of this coating on the adsorption behavior of human serum albumin (HSA). The idea consists in studying coating procedures that do not require synthetic anchors and/or activators to create covalent links,

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but can function using

only natural lipids. For this purpose, the surface is first modified with an anchoring stearic acid (SA) layer, chosen on the basis of our previous work,

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then coated with DPPC thus

yielding a layered “hybrid” lipid film. The organization of the lipid layers, their stability upon conditioning in buffer and the adsorption behaviour of HSA are probed by a variety of techniques including infrared-based techniques sensitive to thin films, X-ray photoelectron spectroscopy (XPS), atomic force microscopy (AFM) and water contact angle measurements. Particular attention is dedicated to the possibilities to explore vertical heterogeneities of the nanostructured surfaces to assess whether adhesive/repellent properties are comparable in the outermost part and in depth. 2. Experimental section 2.1. Surface modification procedures Polycrystalline aluminum wafers (specimens of ~1 cm2 surface area cut from a 2-mm thick plate, Al 99.99%, GoodFellow, France) were used in this study. The sample was polished with a series of SiC papers of decreasing grit size (1200-4000) followed by 1 µm diamond suspensions (Struers, France). The samples were then rinsed in binary mixtures of ultrapure water/ethanol (50%/50%, v/v) in a sonicating bath (70 W, 40 kHz, Branson, USA) for 20 min and dried under nitrogen gas flow. The obtained samples exhibit a native oxide layer, presumably Al2O3, and are called “AlOx-flat” (Table 1). The latter samples were subjected to a hydrothermal treatment yielding a superficial hydroxylation of the surface (samples called “AlOx-rough"), following the procedure described elsewhere. 34 Both AlOx-flat and AlOx-rough samples were coated with lipids as summarized in Table 1. Adsorption of stearic acid (SA, ≥ 98.5%) on the aluminum substrate was carried out following 5 ACS Paragon Plus Environment

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the procedure described previously.

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Briefly, samples were placed in glass Petri dishes

containing the solution of dissolved SA in heptane (HPLC grade, ≥ 99%) at a concentration of 10 mmol/l, and incubated for 1 h under a gentle stirring. The samples were then immersed in a solution of pure heptane for 5 min for rinsing, dried under nitrogen gas flow and used immediately. A phospholipid, dipalmitoylphosphatidylcholine (DPPC, ≥ 99%) was adsorbed on either bare aluminum substrates (AlOx-flat; AlOx-rough) or the same after modification with stearic acid (SA-AlOx-flat; SA-AlOx-rough). For this purpose, DPPC was dissolved in chloroform (HPLC grade, ≥ 99%) and solutions with concentrations varying from 0.0007 to 10 mg/mL were used. Three adsorption procedures were carried out as follows: (i) “Incubation”: samples were placed in glass Petri dishes containing the solution of dissolved DPPC in chloroform and incubated for 1 h under a gentle stirring at room temperature. The samples were then immersed in a solution of pure chloroform for 5 min for rinsing and dried under nitrogen gas flow. This procedure is similar to the one used for stearic acid coating and to the "selective adsorption" procedure in supported catalysts preparation. Because of the application of a rinsing (or washing) step, only DPPC molecules irreversibly adsorbed on the surface will be retained. (ii) “Drop-deposition”: 20 µL of DPPC solution was deposited on the aluminum substrate and left to stand for 2 h in a 1.5 dm3 closed recipient under vacuum at room temperature. This procedure does not involve a washing step and is similar to the "impregnation" procedures in supported catalysts preparation. All the DPPC molecules initially introduced in the deposited drop will be retained on the surface: those that did not initially establish irreversible interactions with surface sites may be structured during the drying step by weaker interactions.


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(iii) “Spin-coating”: a volume of 40 µL of DPPC solution was applied on the alumium substrate, leading to full coverage with liquid. The spin-coater (Laurell technologies corporation, WS-650Mz-23NPP) was accelerated immediately to 3000 rpm for 40 s. The samples were then placed for 2 h in the closed recipient under vacuum at room temperature. This procedure is intermediate between "incubation" and "drop-deposition": no rinsing step is applied, but part of the solution is physically eliminated during the operation of the spincoater. One may surmise that the surface will retain irreversibly adsorbed DPPC molecules and perhaps a limited number of weakly adsorbed ones. All chemical products were purchased from Sigma-Aldrich (Saint-Quentin Fallavier, France) and used without further purification. 2.2. Conditioning in buffers The stability of adsorbed SA film was checked in different standard buffers at room temperature for 2 and 24 h of incubation. The following buffers were used: phosphate buffer saline (PBS, pH ~ 7.4), Tris-HCl (pH ~ 7.4 and ~ 9.4) and carbonate buffer (pH ~ 9.4). For DPPC layers, the conditioning test was performed in Tris-HCl buffer solution (pH ~ 7.4) during 2 h at room temperature. 2.3. Protein adsorption tests Solutions of human serum albumin (HSA, ≥ 99%) were prepared by dissolving the protein powder in Tris-HCl buffer solution (pH~ 7.4) at concentrations of 10, 50, 100, 200, 500 and 1000 µg/mL. The aluminum substrates, either coated with lipids or not, were incubated in protein solutions at room temperature for 2 h. The samples were rinsed in two baths of the buffer used for protein solution and then in one bath of ultrapure water (2 min each) and dried with a nitrogen gas flow. 2.4. Infrared analyses Polarization-modulation infrared reflection absorption spectroscopy (PM-IRRAS) 7 ACS Paragon Plus Environment

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PM-IRRAS is a non-destructive physico-chemical analysis, which is convenient for the characterization of floating monolayers

35

or thin films deposited on metal substrates.

36

This

method utilizes the differences in reflectivity of interfaces for p-polarized (perpendicular to the plane of the surface) and s-polarized (parallel to the plane of the surface) light. A differential reflectance spectrum is computed as follows: ∆R R p − Rs = R 0 R p + Rs

(Eq. 1)

where R0 is the reflectivity without adsorbate, Rp is the intensity of the p-polarized component of the radiation, and Rs is the intensity of the s-polarized component of the radiation. Because of this polarization modulation, the interfering atmospheric absorptions, mostly caused by water vapour and CO2, are eliminated. The film of lipids on the Al substrate interacts with the p-polarized fraction of light, but not with the s-polarized one, resulting in molecular information on the chemical nature of the analyzed film. On flat surfaces, it is possible to deduce the orientation of the molecules with respect to the surface plane. On the contrary, for AlOx-rough, information on the orientation cannot be derived, because of surface roughness, and the infrared spectrum is a statistical average over the different orientations of the molecules with respect to the underlying metallic surface plane.

37

For this reason, the

quantification of adsorbed molecules by IRRAS is extremely complicated. But the areas of specific bands can still be used for a rough, semi-quantitative estimation of the amount of corresponding molecules. In this work, PM-IRRAS spectra were recorded on a commercial Thermo-Scientific (France) Nexus spectrometer. The external beam was focused on the sample with a mirror, at an optimal incident angle of 80°. A ZnSe grid polarizer and a ZnSe photoelastic modulator, modulating the incident beam between p- and s-polarizations (HINDS Instruments, PEM 90, modulation frequency = 37 kHz), were placed prior to the sample. The light reflected at the 8 ACS Paragon Plus Environment

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sample was then focused onto a nitrogen-cooled MCT detector. All the spectra presented were obtained from the sum of 128 scans; the band positions are estimated to have a 8 cm−1 resolution. Grazing-angle attenuated total reflection (GA-ATR) GA-ATR analyses were performed using the VariGATR (Harrick Scientific, Pleasantville, NY) equipped with a horizontal reflection ATR accessory including a germanium crystal. The spectrometer is equipped with a nitrogen-cooled mercury-cadmium-telluride (MCT)wideband detector. The angle of incidence is fixed at 65°. The aluminum sample (AlOx-flat) was placed face-down onthe Ge crystal, and a force was applied via a pressure tip. The background was recorded in ambient conditions without any substrate pressed against the crystal. IR spectra were recorded in a wavelength range from 600 to 4000 cm-1. For each spectrum, 128 scans were collected with a nominal resolution of 8 cm-1. IR analyses were performed on, at least, three independent sets of samples to check the reproducibility. The reported values of the area of a specific band were the average of the results obtained from different spectra. 2.5. X-ray photoelectron spectroscopy (XPS) XPS analyses were performed using a ESCA+ spectrometer (Omicron NanoTechnology), equipped with a monochromatized aluminum X-ray source (powered at 20 mA and 14 kV) and a MCD 128 channeltrons detector. Charge stabilization was insured using the CN10 device with an emission current of 5.0µA and a beam energy of 1eV. Analyses were performed in the sweeping mode; the resulting analyzed area was 1 mm in diameter. A pass energy of 20 eV was used for narrow scans. Under these conditions, the full width at half maximum (FWHM) of the Ag 3d5/2 peak of a clean silver reference sample was about 0.6 eV. The pressure in the analysis chamber was around 10-10 torr. The photoelectron collection angle, θ, between the normal to the sample surface and the analyzer axis was 45°.The 9 ACS Paragon Plus Environment

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following sequence of spectra was recorded: survey spectrum, C 1s, O 1s, N 1s, Al 2p, Cl 2p, P 2p, S 2p and C 1s again to check for charge stability as a function of time and the absence of sample degradation. The binding energy scale was set by fixing the C 1s component due to carbon only bound to carbon and hydrogen at 284.8 eV. The data treatment was performed with the CasaXPS software (Casa Software Ltd., UK). The peaks were decomposed using a linear baseline, and a component shape defined by the product of a Gaussian and a Lorentzian function, in the 70:30 ratio, respectively. Molar fractions were calculated using peak areas normalised on the basis of acquisition parameters and sensitivity factors provided by the manufacturer. Two independent sets of samples were analyzed by XPS to check the reproducibility. 2.6. Atomic force microscopy (AFM) AFM images were recorded using a commercial AFM (NanoScope VIII MultiMode AFM, Bruker Nano Inc- Nano Surfaces Division, Santa Barbara, CA.). The substrates were fixed on a steel sample puck using a small piece of adhesive tape. Images were recorded in air at room temperature (~22°C). AFM experiments were performed using peak force tapping (PFT) mode, as detailed elsewhere. 38 In this mode, the z-piezo is modulated far below the cantilever resonance frequency (2 kHz), with an amplitude around 120 nm. Oxide-sharpened microfabricated Si3N4 cantilevers were used (Bruker Nano Inc- Nano Surfaces Division, Santa Barbara, CA.). The spring constants of the cantilevers were measured using the thermal noise method, yielding values of 0.5 ( ±0.05) N/m and the curvature radius of silicon nitride tips was about 3 nm (manufacturer specification). The root-mean square roughness (Rrms) was calculated on 1x1µm2 height images. For each sample, the reported value is the average of the results obtained with, at least, three different images. 2.7. Water contact angle (WCA)


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Static water contact angles were measured at room temperature using the sessile drop method and image analysis of the drop profile. The apparatus (Krüss, Germany) is equipped with CCD camera and an image analysis processor. The ultrapure water droplet volume had a 1 µL volume and the contact angle was measured about 3s after the drop was deposited on the sample surface. For each sample, the reported value is the average of the results obtained with 3 droplets.

Table 1. Designation of samples used in this study. Details about procedures (×) and characteristics of the obtained surfaces (θw : water contact angle; Rrms : root-mean-square roughness; SD : standard deviation obtained on three independent measurements). hydroxylation AlOx-flat SA-AlOx-flat AlOx-rough SA-AlOx-rough

× ×

Surface modification SA coating DPPC coating × × × × × ×

Surface properties θw ° (±SD) Rrms nm (±SD) 24 (3.0) 4.1 (0.38) 126 (2.9) 3.2 (0.45) 15 (1.2) 19.3 (0.16) 130 (4.3) 20.7 (2.96)

3. Results and discussion 3.1. Self-assembly and organization of lipids on aluminum substrates The nanostructured aluminum oxide layer was obtained by a straightforward and reproducible hydrothermal treatment yielding superficial chemical and structural transformations, as reported elsewhere.

38

The treatment leads to the conversion of a native Al2O3 oxide (sample

called "AlOx-flat") to an oxy-hydroxide, presumably AlOOH (sample called "AlOx-rough"). Figure 1 shows the morphology of flat (AlOx-flat, Figures 1A, E) and nanostructured (AlOxrough, Figures 1B, F) aluminum surfaces probed by AFM, revealing that the nanostructured surface exhibits randomly distributed cylindrical nanostructures (nanoprotrusions) and nanoporous domains (valleys). On the AlOx-flat surface, the presence of stripes and some defects are due to the polishing procedure. These surfaces (AlOx-flat) were only used in the present study as a control to assess and discriminate the effect of the nanoscale topography. When aluminum substrates were coated with SA, the surfaces became hydrophobic while no 11 ACS Paragon Plus Environment

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noticeable morphological changes were observed (Table 1 and Figures 1C, D and G, H). The apparent water contact angle (θw) reached values around 130° and was almost the same on AlOx-flat and AlOx-rough (Table 1), suggesting a complete surface coverage with SA molecules. We may assume that SA molecules interact with the inorganic surface through a coordinative-bonded carboxylate species on the AlOx surface but do not assemble into multilayers, as the presence of free carboxylic acid groups, i.e. unbound to the surface, was not detected in the PM-IRRAS spectra (Figure S1, ESI), in agreement with previous findings. 34

Figure 1. Representative AFM (A-D) height and (E-H) deflection images recorded on (A, E) AlOx-flat, (B, F) AlOx-rough, (C, G) SA-AlOx-flat and (D, H) SA-AlOx-rough samples (for height images z-scale = 140 nm; bars = 500 nm).

DPPC adsorption was performed on AlOx-rough sample using three coating procedures, as described above (see section 2.1.): (i) incubation, (ii) drop-deposition and (iii) spin-coating. The evolution of θw as a function of the concentration of DPPC solution is given in Figure 2 for the three procedures. When DPPC adsorption was performed by incubation on the hydrophilic AlOx, θw reached the values of 75 and 50° in the solvent (without DPPC) and at the lowest concentration of DPPC, respectively (open symbols, Figure 2A). θw decreased then 12 ACS Paragon Plus Environment

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to stabilize at values around 30°. On the hydrophobic SA-AlOx-rough, θw decreased with the increase of DPPC concentration (close symbols, Figure 2A). By using drop-deposition method, the presence of DPPC on the AlOx-rough surface did not induce significant changes of θw, regardless of DPPC concentration (open symbols, Figure 2B), keeping the surface hydrophilic. By contrast, on the SA-modified surface, θw decreased markedly after the adsorption of DPPC at 1 mg/mL or higher, to reach the same level of wettability as for AlOxrough. Finally, when the adsorption was performed using spin-coating, a slight progressive increase of θw was observed upon DPPC adsorption on AlOx-rough, and an opposite trend upon DPPC adsorption on SA-AlOOH (Figure 2C). This evolution only started at a concentration of 1 mg/mL and levelled off for concentrations higher than 5 mg/mL. The changes in surface wettability are certainly correlated with the presence of DPPC on the aluminum surfaces. The adsorption of DPPC was also monitored by PM-IRRAS analysis. The IR vibrational features and band assignments are detailed in ESI (see Figure S2 and Table S1). The integrated intensity of the C=O stretching band at 1741 cm-1 due to the ester group (Aν(C=O)), i.e. the area of the band, was used to provide a rough estimate of the adsorbed amount of DPPC. Aν(C=O) increased when increasing the concentration of DPPC in the solution for all three deposition methods. When the incubation procedure was used (Figure 2D), the adsorbed amount levelled off for rather low DPPC concentrations, consistent with a Langmuir-like site adsorption. In this case, the saturation DPPC coverage (all strong adsorption sites occupied) could not be exceeded. Saturation coverages on bare and SA-coated nanostructured surfaces are similar and quite low. For the drop deposition procedure (Figure 2E), it is known that all DPPC introduced remains on the surface, and therefore plotting Aν(C=O) as a function of concentration must give a straight line going through the origin; furthermore, all data points must fall on the same line, 13 ACS Paragon Plus Environment

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irrespective of the underlying surface. This is indeed what is observed. The data of Figure 2E could be used for absolute quantification of the adsorbed amounts in the other two procedures, but on a rough surface these values could not be translated to surface densities in mol/nm2, and therefore the calculation was not carried out. Finally, for the spin-coating procedure (Figure 2F), the deposited quantities increased with DPPC concentrations in the solution, but not linearly. This is probably because the volume of DPPC solution deposited on the sample is higher in the case of the drop deposition procedure compared to spin-coating. Here too, the presence of an underlying hydrophobic SA layer does not seem to affect the adsorbed amount of DPPC. 150

2.5

incubation

A

125

D

2.0

(C=O)

75

Aν

o

θw ( )

100

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CDPPC (mg/ml)

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(C=O)

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75

Aν

)

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drop-deposition

B

100

o

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θw (

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CDPPC (mg/ml)

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CDPPC (mg/ml)

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spin-coating

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100 (C=O)

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)

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CDPPC (mg/ml)

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θw (

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50

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CDPPC (mg/ml)

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CDPPC (mg/ml)


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Figure 2. Evolution of the water contact angle (θw, A-C) and the intensity of the IR band for the ν(C=O) ester group of DPPC at 1741 cm-1(Aν(C=O), D-F) as a function of the concentration of DPPC solution used for the coating. Data obtained on AlOx-rough (open symbols) and SA-AlOx-rough (closed symbols) surfaces by using incubation (A, D), drop-deposition (B, E) or spin-coating (C, F).

The composition of the lipid layers may be also conveniently probed by means of XPS analyses. This technique has the advantage of being sensitive to weak variations in the surface composition and selective towards bio-organic/inorganic interfaces.

39-41

A detailed

description of spectral data, peak decomposition and assignment are presented in ESI (Figures S3). The amount of adsorbed DPPC was evaluated by means of the molar concentration of the contribution at 402.3 eV in the N 1s peak (denoted as NDPPC) which is an exclusive marker of this phospholipid for the samples studied here (Figure S4A, ESI). Results showed that no clear distinction between AlOx-rough (open symbols) and SA-AlOx-rough (closed symbols) can be made on NDPPC values, regardless of the deposition method used. The differences observed on the mole fraction of NDPPC between SA-AlOx-rough and AlOx-rough (Table 2) are only due to random sampling variability, as confirmed by data given in Figure S4A (ESI). This indicates that the underlying SA layer has a little influence on the adsorbed amount of DPPC. Based on the evolution of NDPPC (Table 2), the adsorbed amount of DPPC followed this trend: incubation < spin-coating < drop-deposition, in agreement with PM-IRRAS data (Figure 2D-F). The elemental composition given by XPS also revealed the presence of Cl, originating from chloroform, when DPPC was adsorbed by incubation in contrast to the other methods (Table 2). The trend was more pronounced on AlOx-rough compared to SA-AlOx-rough. This result supports the attribution of the IR band at 1546 cm-1 to chloroform 42 , suggesting that solvent molecules remain entrapped on the surface after drying, particularly on AlOx-rough. The presence of chloroform explains the unexpected hydrophobic character observed on AlOx15 ACS Paragon Plus Environment

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rough surface after an incubation in solvent or in low concentration DPPC solutions (Figure 2A, open symbols). Solvent entrapment was a serious issue, as attempts to remove it in vacuum were not completely successful, probably due to a strong interaction between chloroform and aluminum substrate. Indeed, CHCl3 can adsorb on alumina from the gas phase and interact with either amphoteric or basic sites.

43

It can even slowly react at room

temperature with atmospheric O2 to give formate ions, releasing chlorine.

44

Because of this

contamination with solvent, and of the limited amounts of DPPC retained (see above), the deposition of DPPC using incubation is not a relevant way to coat the samples studied here with DPPC, and it will not be further considered in the following. Taken together, the evolutions of θw (Figures 2B, C) and Aν(C=O) (Figure 2E, F) provide relevant information regarding the organization of the DPPC layers at the molecular level. Owing to their amphiphilic properties, DPPC molecules have the aptitude to change the surface wettability, depending on their amount/organization in the adsorbed phase in combination with the properties of the underlying substrate. According to Jurak and Chibowski, the surface free energy decreased appreciably after the deposition of a DPPC bilayer on hydrophilic surfaces (glass or mica), while it remained almost unchanged on more hydrophobic surface (PMMA). In this study, the authors stated that the surface free energy is determined by the orientation of adsorbed DPPC molecules and the homogeneity of the layer. 45

On AlOx-rough surface, the formation of a DPPC monolayer is expected to lead to a more

hydrophobic surface, as DPPC molecules should interact with the surface via their polar heads and expose their hydrophobic chains. The evolution of θw (Figure 2B, open symbols) showed however that the surface remains hydrophilic after the deposition of DPPC, suggesting the formation of, at least, one bilayer. On the hydrophobic SA-AlOx-rough, the abrupt decrease of θw (Figure 2B, closed symbols) may be consistent with the formation of a DPPC monolayer in which molecules interact with the underlying SA layer via their acyl chains and expose 16 ACS Paragon Plus Environment

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their hydrophilic head group. However, on the top of this monolayer the presence of one or several bilayers is very likely, because an increase of the DPPC concentration from 1 to 5 mg/mL leads to a significant increase of the adsorbed amount (increase of Aν(C=O)) while θw was kept unchanged. A similar trend was observed when spin-coating was used, but the surface is less hydrophilic, suggesting that the DPPC film exhibited more discontinuity (Figure 2C). The formation of multilayered DPPC film using drop-deposition or spin-coating, as suggested by results given in Figure 2, is supported by previous observations made on flat substrates.

46

While the first monolayer of DPPC is strongly adsorbed on the surface, the

following layers are formed through weak bonding with the first one.

Table 2. Surface concentration (mole percentage computed over all elements except hydrogen) of elements determined by XPS and apparent water contact angle, θw, recorded on AlOx-rough and SA-AlOx-rough samples after adsorption of DPPC. Note that N402.3 is denoted NDPPC in the text. XPS Al 2p

P 2p

Cl 2p

C 1s

O 1s

θw(°) (±SD) N399.8

N 1s N402.3

Ntot

0.39 0.30 0.26

b.d.l 0.12 0.47

0.39 0.42 0.73

50.43 (5.89) 39.80 (2.83) 39.54 (3.66)

0.17 0.18 0.19

0.11 0.25 0.34

0.28 0.43 0.53

116.30 (7.23) 110.63 (5.00) 40.56 (5.20)

incubation AlOx-rough 0.0007mg/mL 0.007mg/mL 0.07mg/mL SA-AlOx-rough 0.0007mg/mL 0.007mg/mL 0.07mg/mL AlOx-rough 1mg/mL SA-AlOx-rough 1mg/mL

27.62 21.31 23.34

b.d.l 0.30 0.72

0.37 0.33 0.53

26.75 25.31 23.15

0.23 0.43 0.67

0.07 0.09 0.00

0.92

1.88

0.00

83.00

12.77

0.06

1.37

1.43

16.73 (7.79)

4.55

1.57

0.00

76.65 0.00 spin-coating

1.09

1.09

16.15

28.7 (13.87)

1.14

0.00

60.98

25.33

0.10

0.83

0.93

43.4 (0.88)

0.78

0.00

59.21

26.75

0.11

0.68

0.79

80.0 (3.34)

AlOx-rough 5mg/mL 11.62 SA-AlOx-rough 5mg/mL 12.48 b.d.l : below detection limit

21.62 37.13 35.20

50.01 40.52 39.49

31.32 41.37 34.47 39.27 38.40 37.25 drop-deposition

The morphology of DPPC films was probed by AFM, recorded in the dried state, i.e. after the solvent was completely evaporated. Results showed that when DPPC was adsorbed by dropdeposition on the flat substrate, the film exhibited a lamellar structure (Figure 3A). The Young modulus map given in Figure 3B, obtained using the quantitative nanomechanical 17 ACS Paragon Plus Environment

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38

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, which reflects the elasticity of the layer in a semi-quantitative

way, showed almost no contrast except at the boundary lines of the DPPC layers. This suggests that the substrate is completely covered by lipids and that the AFM tip did not enter into a direct contact with the underlying inorganic substrate. Typical line scans revealed an interlayer spacing of ca. 5 nm, as shown in Figure 3A (line scan 1) and confirmed by several scan lines performed on different locations of the AFM images. The values determined with several line scans, performed in different zones and different images, revealed an average height of 5.4 ± 0.7 nm (Figure 3C). This is close to the repeat distance of DPPC multi-bilayers observed elsewhere

47

and corresponds to the thickness of one DPPC bilayer.

48

By contrast,

when spin-coating was used, the lamellar structure was less obvious and the surface was dominating by fairly circular droplets (Figure S5, ESI).

Figure 3. Representative AFM height images (A, D) and Young modulus map (B, E) recorded on SA-AlOx-flat samples after DPPC adsorption prior to (A, B) and after (D, E) conditioning in Tris-HCl buffer for 2 h. Scan


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lines taken at the location indicated in (A, D) by straight line and numbered. (C, F) Histograms of domain heights corresponding to sample (A) and (D), respectively (for height images z-scale = 25 nm, bars = 200 nm).

The dewetting patterns formed after DPPC deposition, as shown by AFM, result from the rupture of the outermost layers upon solvent evaporation, with possible formation of holes, islets, etc. These processes seem to be strongly influenced by the method of deposition (dropdeposition vs spin-coating), the wettability of the underlined substrate (presence or not of SA layer) and its roughness (flat vs rough). AFM images clearly showed a well-defined lamellar structure of DPPC film on the hydrophobic flat substrate (SA-AlOx-flat) only when DPPC was drop-deposited. On the remaining cases, the lamellar structure was not obvious, but some disordered domains with smaller areas were still visible (Figure S5, ESI). These results suggest that the drop-deposition allows a better spreading of phospholipids on SA-AlOx-flat. By contrast, the formation of the lamellar structure is hindered when phospholipids directly interact with the aluminum substrate, or when the dewetting process is accelerated, i.e. via spin-coating.


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Figure 4. Representative AFM height images recorded in the dried state on SA-AlOx-rough samples after DPPC adsorption prior to (A) and after (B) conditioning in Tris-HCl buffer for 2 h. (C, D) Comparison between AFM height images recorded on SA-AlOx-rough sample modified with DPPC in (C) the hydrated state (buffer) and in (D) the dried state. Scan lines were taken at the location indicated by dashed lines (for height images z-scale = 140 nm, bars = 200 nm).

On the nanostructured substrates, AFM images did not show any obvious lamellar structure (Figure 4A). Moreover, the nanoporous domains, observed on bare AlOx-rough, remained visible after DPPC deposition, as also evidenced by scan lines (Figure 4A and S6, ESI). Even though the lipid domain sizes formed on the flat surfaces may significantly exceed those of nanoporous areas, the presence of lamellar structures on top of the nanostructured surfaces has never been observed by AFM. While the amount of adsorbed DPPC is not appreciably affected by the presence of a hydrophobic SA layer, it appears that the organization of the DPPC film is different in terms of number of layers (at least two and three DPPC layers on SA-coated and non-coated AlOxrough substrates, respectively), layer discontinuity and probably molecule orientations. It is noteworthy that the formation of mixed SA/DPPC bilayers was not observed. This formation would involve a mechanism of exchange leading to the detachment of self-assembled SA molecules from the aluminum surface, and PM-IRRAS data did not reveal any significant decrease of the amount of self-assembled SA (data not shown), nor the presence of free carboxylic acid groups. It may thus be concluded that the main role of the anchoring SA layer consists in improving the spreading of phospholipids leading to their organization in more continuous and less disordered layers. Clearly, the question deserves further investigation, and requires determining the number of layers, the orientation of DPPC molecules, etc. This may be achieved by means of neutron reflectivity, for instance, even the surface roughness remains a physical constraint.

3.2. Stability of lipid layers in buffers 20 ACS Paragon Plus Environment

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The stability of DPPC layers upon conditioning in aqueous solution at room temperature was examined by incubating the samples in a Tris-HCl buffer (pH ~ 7.4) for 2h. These conditions were shown to keep the underlying self-assembled SA layer intact (Figure S7, ESI). Insights regarding the stability of DPPC layers in Tris-HCl buffer were provided by IR data. Figure 5 presents the evolution of Aν(C=O) measured after conditioning in buffer as compared to Aν(C=O) measured prior to conditioning. This ratio reflects the fraction of DPPC that remains adsorbed after conditioning. This ratio remained close to 1 on the nanostructured surfaces, AlOx-rough and SA-AlOx-rough (Figure 5A), indicating that the adsorbed amount

A

Aν(C=O) [Hyd] / Aν(C=O)

of DPPC did not change significantly upon conditioning in Tris-HCl buffer. 1.2 1.0 0.8 0.6 0.4 0.2 0.0

AlOx-rough

B

Aν(C=O) [Hyd] / Aν(C=O)

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SA-AlOx-rough

1.2

drop-deposition spin-coating

1.0 0.8 0.6 0.4 0.2 0.0

AlOx-flat

Figure 5. Evolution of the ratio of Aν(C=O) [Hyd], to

SA-AlOx-flat

Aν(C=O) measured after conditioning in Tris-HCl buffer (pH~7.4), denoted

Aν(C=O) measured prior to conditioning. IR data were recorded on (A) flat (AlOx-flat, SA-

AlOx-flat) and (B) rough (AlOx-rough, SA-AlOx-rough) substrates, as indicated.

The situation was different on flat substrates (Figure 5B), especially when the surface was modified with SA and when DPPC was adsorbed by spin-coating: about the half of the amount of adsorbed DPPC detached from the surface upon conditioning in buffer. These findings suggest that the stability of DPPC layers is influenced by the nanoscale topography


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when the surface is hydrophobic, i.e. coated with SA, and when DPPC was adsorbed by spincoating. AFM images support this trend. On SA-AlOx-flat, conditioning in buffer caused the partial loss of the lamellar structure, leading to a dominant presence of fairly regular droplets (Figure 3D). However, the Young modulus map suggests that the surface remains completely covered with lipid layers (Figure 3E), as observed prior to the conditioning test (Figure 3B). Scan lines revealed that the average heights of these droplets did not change appreciably compared to the height of the lamellar structures, i.e. around 5 nm which corresponds approximately to the thickness of one bilayer (Figure 3D, line 2 and Figure 3F). The conditioning in buffer induced only a little change on the morphology of the lipids film on the nanostructured substrates, particularly on SA-AlOx-rough (Figure 4A and B), but also on the remaining samples (Figure S6, ESI). This supports the stability of the DPPC film as revealed by IR data. Besides, to examine the possible effect of drying on the reorganization of phospholipids, AFM imaging was carried out in the hydrated state, i.e. in buffer solution, then after drying. Results showed that the morphology of the film was only slightly modified, probably due to the sensitivity of the layer in the liquid phase (Figure 4C), and the nanostructured feature of the underlying substrate remained visible. After drying, the film morphology was kept almost unchanged (Figure 4D), suggesting that drying did not induce drastic re-organization of phospholipids on the nanostructured substrates. The above findings indicate that the nanoscale topography enhances the stability of DPPC layers upon conditioning in buffer. This effect probably stems from the fact that DPPC film did not organize into lamellar structure and conforms to the nanostructured surface topography, particularly in nanoporous domains (valleys).

3.3. Adsorption behaviour of proteins


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Protein adsorption tests were performed in Tris-HCl buffer using HSA following the procedure described above. The samples were characterized by means of IR analysis, which allowed to specifically identify proteins through their amide bands and provided a rough estimate of their amount (Figure S8, ESI). Results indicated that the area of amide bands,

Aamide, was higher on AlOx-rough compared to SA-AlOx-rough (Figure S8A, B). On the former surface, electrostatic interaction is probably the predominant driving force for protein adsorption. Indeed, the theoretical isoelectric point (iep) of HSA, computed on the basis of the amino acid composition of the protein, is 5.7,

49

while experimental values around 5.0 have

been reported for serum albumin. 50 The point of zero charge of pseudo-boehmite, presumably present on AlOx-rough surfaces, has been reported to be around 9.

51

Accordingly, in the

studied conditions (Tris-HCl buffer, pH = 7.4), the surface charge is positive while the protein charge is negative and electrostatic interactions are favorable to the adsorption of proteins on AlOx-rough surface. By contrast, the area of amide bands was significantly lower on DPPCmodified surfaces, suggesting a lower adsorbed amount of proteins (Figure S8C, ESI). The meaning of these results appears more clearly if it is summarized in terms of a yield of protein repellency ( Y rpIR ), which can be defined as follows:

YrpIR (%) =

Aamide(bare)− Aamide(DPPC) Aamide(bare)

(Eq. 2)

where Aamide(bare) and Aamide(DPPC) are the areas of amide bands observed on the surface of interest, respectively without DPPC (i.e. AlOx or SA-AlOx) and after DPPC deposition. It can be seen that YrpIR is higher when DPPC is deposited by drop-deposition on the nanostructured surfaces and may reach about 80%, while it does not exceed about 60% when it is deposited by spin-coating (Figure 6A).


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drop-deposition spin-coating 100

Yrp(IR) (%)

A

80 60 40 20 0

AlOx-rough

SA-AlOx-rough

AlOx-rough

SA-AlOx-rough

AlOx-flat

SA-AlOx-flat

100

Yrp(XPS) (%)

B

80 60 40 20 0

C

100

Yrp(XPS) (%)

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

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80 60 40 20 0

Figure 6. Yield of repellency (%) computed from (A) IR (YrpIR) or (B, C) XPS (YrpXPS) data recorded on (A, B) rough (AlOx-rough, SA-AlOx-rough) and (C) flat (AlOx-flat, SA-AlOx-flat) substrates, as indicated.

XPS analyses may also provide an estimate of the adsorbed amount of proteins. For this purpose, spectral data were explored on the basis of methodological developments reported recently to characterize bio-organic/inorganic interfaces, in particular to extract exclusive markers of proteins.

40-41

A detailed description of the methodology applied here is given in

ESI. Figure S4B presents the variation of the molar concentration of C287.8, which is assigned to carbon doubly bonded with oxygen, as a function of N399.8, revealing a 1:1 relationship which suggests that the N 1s contribution at 399.8 eV is mainly due to peptidic links in proteins. This trend demonstrates the accuracy of XPS data and the validity of the attribution of chemical functions. As regards, the N 1s peak at 401.0 eV, which like the previous one is only observed after HSA adsorption, it may be due to the amine groups in lysine and arginine residues of HSA (the protein contains 59 Lys, ~10%, and 24 Arg, 4%) which are partially 24 ACS Paragon Plus Environment

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protonated in the studied conditions (Tris-HCl buffer, pH~7.4). Accordingly, the total nitrogen due to proteins (NProtein) may be computed as follows:

Nprotein = N399.8 + N401.0

(Eq. 3)

or using an equivalent equation:

Nprotein = Ntot -NDPPC

(Eq. 4)

where NDPPC = N402.3 corresponds to the mole concentration of nitrogen typical of DPPC molecules. Accordingly, a yield of protein repellency can be computed from XPS data as follows:

YrpXPS (%) =

N protein (bare) − N Protein (DPPC)

(Eq. 5)

N Protein (bare)

where NProtein(bare) and NProtein(DPPC) are the molar concentrations of protein-marker nitrogen (computed using Eq. 3 or Eq. 4) observed on the surface of interest, respectively without DPPC (i.e. AlOx or SA-AlOx) and after DPPC deposition. Results showed that the yield of repellency computed from XPS data, YrpXPS, followed almost the same trend as observed for the one computed from IR data, YrpIR on the nanostructured surfaces, taking into account the random sampling variability (Figure 6B). This trend was, however, not observed on flat substrates (Figure 6C). Results showed that YrpXPS values ranged between 92 and 98%, revealing the remarkable performances of these coated flat surfaces to repel proteins, regardless of the presence of an underlined SA layer and the procedure used for DPPC coating. McClellan and Franses have shown that the presence of a DPPC layer on solid surfaces leads to a decrease of the adsorbed amount of BSA by about 58%.

52

Vermette et al.

reported an elegant way to tune the adsorption of plasmatic proteins by exposing either the acyl or the zwitterionic group of DPPC to the solution.

53

In the latter case, the authors

obtained a yield of repellency equal to 67% for HSA and 76% for fibrinogen. Phosphatidylcholine-based polymers were also used instead of DPPC to repel BSA, such as in 25 ACS Paragon Plus Environment

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the studies of Malmsten et al. (Yrp ~ 42%),

54

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Hoshi et al. (Yrp ranges from ~60 to ~80%) 55-56

and Tauk et al. (Yrp ranges from ~85 to ~99%).57 The variations in these studies are generally related to the nature of the polymer and the procedure used for its immobilization on a solid surface. In the present study, the techniques used to estimate the amount of adsorbed proteins, IR and XPS, provide data which are different in terms of selectivity, precision and accuracy. The method adopted here to compare different situations (adhesive or repellent) in a series of samples was to work with normalized data; this does not provide absolute protein concentrations but allows situations to be compared with each other. In fact, while both techniques probe an area of several hundred micrometres square on the surface plan, the main difference between them concerns their respective analysis depths. XPS probes a thickness up to about 7 nm (see ESI for more details), PM-IRRAS may probe a depth of more than 1 µm. On flat (unstructured) surfaces (AlOx-flat), the organic-containing layer is only a few nanometres thick, and both techniques would have access to all adsorbed molecules. Nanostructured surfaces (AlOx-rough), in contrast, consist of stacked cylindrical nano-objects in a structure that can be several tens of nm thick. Accordingly, it is reasonable to assume that on AlOx-rough, XPS probes the outermost part of the surface and PM-IRRAS the whole nanostructured surface. As the evolution of YrpIR and YrpXPS, computed on the nanostructured surfaces, followed almost the same trend (Figure 6A and B) it may be concluded that protein repellency occurs on the whole surface in a comparable way. In other words, the data in Figure 6 do not reveal any obvious vertical heterogeneity, suggesting that the repellency of proteins occurs in a similar way on the outermost part (nanoprotrusions) as within nanoporous domains (valleys).


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The dependence between the amount/organization of DPPC layers and the adsorption behaviour of protein was examined by means of correlations involving the amount of adsorbed proteins to the respect of (i) the amount of adsorbed DPPC (estimated by IR, Aν(C=O), or by XPS, NDPPC) and (ii) surface roughness, Rrms (estimated by AFM). A clear trend is that Nprotein decreased when the amount of deposited DPPC increased irrespective of whether the AlOx-rough surface is modified with SA or not (Figure S9A, ESI). Similar tendency was observed with IR data (Figure S9B). Another trend is also revealed when considering the surface roughness Rrms (Figure S9C and D, ESI), showing that HSA adsorption is more favourable when the surface roughness increases. Thus, it may be concluded that each parameter - chemistry and roughness - influences the amount of adsorbed proteins independently. However, these two parameters are not independent, as the change of roughness is essentially due to the adsorption/organization of DPPC on the nanostructured support. The significant variation of Rrms from about 15 to about 22 nm due to the organization of DPPC is influenced by the topography of the underlying aluminum substrate, as on flat supports the deposition of DPPC in similar conditions induces only Rrms variations in the range of 1-2 nm. 46 These results support the idea that chemistry and roughness cannot be used as linearly independent predictors for protein affinity for the studied surfaces. Furthermore, the dependence of the amount of adsorbed proteins on both the surface chemistry (amount of DPPC) and roughness (Rrms) reflects a dependence with the organization of DPPC layers, including number of layers, layer discontinuity, molecules alignment and orientation, etc, which impact the accessibility of the zwitterionic moieties towards proteins. The mechanism of protein repellency by zwitterionic functional groups has been studied by numerous authors, but interfacial processes remain incompletely understood. Schlenoff has recently reviewed the main processes by which proteins may be repelled from zwitterion27 ACS Paragon Plus Environment

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modified surfaces. 58 This author points out the importance of the amount and organization of interfacial water molecules, located around zwitterions, which cause a lowering of interfacial energy. Other mechanisms, e.g. associated with steric effects and ion release processes, are also listed and may contribute to the mechanism of repellency. Deciphering the mechanism of interaction of HSA with DPPC-modified surfaces requires coping with the complexity of the organic adlayer, which includes lipids (fatty acids and/or phospholipids), proteins and, unavoidably, adventitious organic contamination. The possibility of gathering more information regarding the composition of the adlayer is examined using XPS data and correlation between consistent spectral data, as reported elsewhere.

41

Figures 7A and B present the plot of NDPPC and Nprotein, respectively, vs. total

carbon (Ctot), showing the impact of each compound, i.e. phospholipids and proteins, on the nature of the adlayer. The adsorption of DPPC leads to an increase of the total carbon, as expected. More interestingly, the regression line computed by considering all DPPC-modified samples has a slope of 0.027 which is close to the N/C ratio expected for DPPC, 0.025 according to the structural formula, C40H80NO8P, (Figure 7A). By contrast, the enrichment of the adlayer with proteins is accompanied with a decrease of the total carbon (Figure 7B). The regression line has a slope of 0.28 which is consistent with the N/C ratio of HSA (0.27) computed on the basis of amino acid amount and protein composition. However, it is not known whether the decrease of Ctot is due to a screening effect of the protein adlayer (presumably on top of the DPPC layer), to a partial desorption of phospholipids, or to a combination of both phenomena.


Page 29 of 35

1.5

12

A 1.0

0.5

0.0 0

20

B

10

Nprotein (%)

NDPPC (%)

40

60

80

100

8 6 4 2 0 0

20

Ctot (%)

40

60

80

100

Ctot (%)

Figure 7. Relationship between molar concentrations, as indicated, measured by XPS recorded on AlOx-rough (open symbols) and SA-AlOx-rough (closed symbols or semi-open) after modification with DPPC, using dropdeposition (, green) or spin-coating (, yellow), and subsequent adsorption of HSA (). Dashed lines correspond to the regression lines: (A) NDPPC = 0.027060 × Ctot - 0.945123 (adjusted R2 = 0.90, correlation coefficient between NDPPC and Ctot is equal to 0.95 and p-value is less than 0.001 (0.00000001182)); (B) Nprotein = -0.27714 × Ctot+ 23.17760 (adjusted R2 = 0.94, correlation coefficient between Nprotein and Ctot is equal to 0.97 and p-value is less than 0.001 (0.000000107)).

The discrimination between these interfacial phenomena can be made by means of IR data, as in PM-IRRAS analyses, all the organic adlayer is probed without a significant screening effect. 1.0

Aν(C=O) [Prot] / Aν(C=O)

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

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drop-deposition spin-coating

0.8

0.6

0.4

0.2

0.0

AlOx-rough

SA-AlOx-rough

Figure 8. Evolution of the ratio of Aν(C=O) measured after conditioning in protein solution (HSA in Tris-HCl buffer, pH~7.4), denoted Aν(C=O) [Prot], by Aν(C=O) measured prior to conditioning. IR data were recorded on AlOx-rough and SA-AlOx-rough substrates, as indicated.


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Figure 8 presents the evolution of Aν(C=O) measured after protein adsorption tests as compared to Aν(C=O) measured before. These data relate the stability of the DPPC layers in protein solution. Results revealed that this ratio was significantly lower compared to that observed in buffer solution without proteins (see Figure 5B), independently on the coating procedure and the presence of SA layer. These trends suggest a partial desorption of phospholipids during the protein adsorption test. Phospholipids desorption may occur through adsorption exchange or via the formation of a soluble DPPC-HSA complex, in view of the high affinity of these entities. This question could be addressed by monitoring HSA adsorption using real time techniques, but the surface roughness would certainly hamper data analysis and interpretation. Another alternative would be to perform successive adsorption cycles, or to use proteins with different affinity toward DPPC.

4. Conclusion The formation of lipid layers, made with fatty acids and/or phospholipids, on surfaces with random nanoscale topography was investigated by means of a variety of techniques, including AFM, XPS, IR and water contact angle measurements. It was found that the organization of phospholipids is strongly influenced by the topography of the underlying substrate, and other factors, including the coating procedure and surface wettability. On flat surfaces, the films exhibit multilamellar structures, which are not completely preserved upon conditioning in buffer solution, while on nanostructured surfaces, DPPC film conforms to the nanoscale topography but is appreciably more stable. Upon conditioning in protein solution, all the surfaces coated with DPPC showed an antifouling behaviour with different yields of repellency (Yrp). The latter parameter seems to be strongly influenced by the amount/organization of DPPC on the nanostructured substrate. Moreover, the mechanism of protein interaction with the coating led to a partial desorption of phospholipids. By exploring XPS and IR data to probe the vertical heterogeneity of the 30 ACS Paragon Plus Environment

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nanostructured surface, it was shown the amount of adsorbed proteins did not differ significantly on the outermost part and within nanoporous domains. The bio-functionalization of the nanostructured substrate reported in this study can be extended to other inorganic materials with stochastic topographies or other coating procedures, e.g. from aqueous solution, and is particularly important in applications such as drug loading, anti-biofilm surfaces and cell culture.

Acknowledgements The authors thank Christophe Calers (LRS, UPMC, France) for his assistance in XPS analyses, Michel Genet (IMCN, UCL, Belgium) and Neal Fairly (Casa Software Ltd., UK) for fruitful discussion. I.L. acknowledges financial support of the French Embassy in Lithuania (Egide # 719306H and Campus France No. 785192H grants) and Research Council of Lithuania (SMM-01-V-V-02-003).


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