Comparative Stability Studies of Poly(2-methyl-2-oxazoline) and Poly(ethylene glycol) Brush Coatings
© The Author(s) 2012
Received: 19 September 2011
Accepted: 31 October 2011
Published: 9 February 2012
Non-fouling surfaces that resist non-specific adsorption of proteins, bacteria, and higher organisms are of particular interest in diverse applications ranging from marine coatings to diagnostic devices and biomedical implants. Poly(ethylene glycol) (PEG) is the most frequently used polymer to impart surfaces with such non-fouling properties. Nevertheless, limitations in PEG stability have stimulated research on alternative polymers that are potentially more stable than PEG. Among them, we previously investigated poly(2-methyl-2-oxazoline) (PMOXA), a peptidomimetic polymer, and found that PMOXA shows excellent anti-fouling properties. Here, we compare the stability of films self-assembled from graft copolymers exposing a dense brush layer of PEG and PMOXA side chains, respectively, in physiological and oxidative media. Before media exposure both film types prevented the adsorption of full serum proteins to below the detection limit of optical waveguide in situ measurements. Before and after media exposure for up to 2 weeks, the total film thickness, chemical composition, and total adsorbed mass of the films were quantified using variable angle spectroscopic ellipsometry (VASE), X-ray photoelectron spectroscopy (XPS), and optical waveguide lightmode spectroscopy (OWLS), respectively. We found (i) that PMOXA graft copolymer films were significantly more stable than PEG graft copolymer films and kept their protein-repellent properties under all investigated conditions and (ii) that film degradation was due to side chain degradation rather than due to copolymer desorption.
Densely-grafted polymer films (brush regime) on metal oxide surfaces are frequently applied to convey biopassive properties, i.e. to reduce protein adsorption , bacteria adhesion , and cell-surface interactions . Poly(ethylene glycol) (PEG)  represents the gold standard in this respect [2, 5–15], but other polymers such as polyacrylamide (PAAM) , poly(N-vinyl pyrrolidone) (PVP) , and peptidomimetic polymers (PMP1 ) as well as poly(2-methyl-2-oxazoline) (PMOXA) [19–21] have been shown to be similarly effective.
The preference for PEG-based non-fouling materials is due to its biocompatibility, non-immunogenic and non-cytotoxic properties . Moreover, PEG has been approved by the US food and drug administration (FDA) for application in pharmaceutical and coating technologies [22–24]. Nevertheless, several limitations of PEG-based technology have been reported. One of them is loss of biopassive function in case of long-term application [3, 25–27]. McGary  reported PEG degradation upon aging in aqueous solutions. It was believed that PEG degrades by auto-oxidation due to repetitive oxygen units in its structure. Moreover, it has been reported that PEG degrades autocatalytically in bulk (solid state)  as well as in surface-bound state [30–33] and in dilute solutions . A discussion of complications associated with the non-biodegradability and tendency to autoxidation in the presence of oxygen can also be found in a recent review by Knop et al. .
Zoulalian et al.  reported instability of PEG-based films on titanium oxide (TiO2) and niobium oxide (Nb2O5) upon exposure to different aqueous media, with and without exposure to light. The authors proposed that photocatalytic activity of certain substrates (such as TiO2, but not Nb2O5) is one of the factors affecting PEG (and possibly other polymer) degradation. Moreover, the authors suggested that 4-(2-hydroxyethyl) piperazine-1-ethanesulfonic acid (HEPES) contributes to degradation of PEG. Wet oxidation of PEG with various molecular weights between 2 and 20 kDa under high pressure and at elevated temperature was investigated by Imamura et al. . The authors suggested that PEG susceptibility to degradation correlated to the ease of intramolecular hydrogen abstraction in the propagation step of oxidation.
Polypeptides or peptidomimetic polymers are potential alternatives to PEG for applications where non-fouling properties are required. For example, Chilkoti et al.  reported the application of artificial thermoresponsive elastin-like polypeptides in drug delivery to combat solid tumours in mice. It was found that this peptidomimetic polymer enhanced drug accumulation by ~5-fold at the tumour site level if compared to that delivered using PEG . This finding provides evidence that the peptidomimetic polymer created a stable, stealth drug carrier upon exposure to serum protein and blood cells contained in blood plasma of the mice. Also, Messersmith  performed a long-term study in cell culture medium using a mono(ethylene glycol)-functionalized peptidomimetic polymer as surface coating on titania surfaces, anchored through an oligo(DOPA-lysine) binding moiety. The authors reported resistance to cell adhesion for more than 5 months, which is much longer than what can be achieved with PEG-based systems. However, it remains unclear to what extent the ethylene glycol functionalization is essential to long-term performance of this particular peptidomimetic polymer.
Another promising polymer with a peptidomimetic structure is poly(2-methyl-2-oxazoline) (PMOXA) [19–21, 39]. PMOXA has been reported to have favourable properties for a number of biological and medical applications, such as stealth liposomes for cell targeting and drug delivery [40–45]. Protein adsorption  and bacteria adhesion  on PMOXA films have already been compared to those on PEG films, where PMOXA has shown comparable non-fouling properties as PEG if organized as a brush with optimum density. However, no systematic comparative stability study between PMOXA and PEG has so far been published to the best of our knowledge.
In this study, the stability of PMOXA films was compared to that of PEG films with Nb2O5 as substrate. The chemical structure of PMOXA suggests that this polymer is less prone to oxidation compared to PEG since N-vicinal C–H bonds are less polarized than O-vicinal C–H bonds. Thus degradation initiated through hydrogen abstraction in the PMOXA chains is presumably less likely. To perform a direct comparison, the PMOXA-based surface coating was designed in analogy to PEG-based coating in previously reported quantitative study , i.e. PLL-X graft copolymers (X = PMOXA or PEG). These graft copolymers present a polycationic poly(l-lysine) backbone that allows spontaneous assembly and anchoring of the grafted copolymer to negatively charged surfaces. Nb2O5 is a particularly suitable substrate considering its high negative surface charge density .
A solution of 10 mM H2O2. H2O2 was used to represent oxidative substances secreted by cells such as macrophages or bacteria.
A solution of 160 mM ion concentration that mimics the ionic strength of body fluid. The solution contained 10 mM HEPES + 150 mM NaCl. The 150 mM NaCl presents a solution of salt that is isotonic with the body fluids, while the 10 mM HEPES acts as a buffering agent to maintain the physiological pH (~7.4).
A solution of 10 mM H2O2 + 10 mM HEPES + 150 mM NaCl. This solution mimics the physiological solution (10 mM HEPES + 150 mM NaCl), in the presence of an oxidative substance (10 mM H2O2).
The film stability was investigated using three different surface characterization techniques, i.e. variable angle spectroscopic ellipsometry (VASE), X-ray photoelectron spectroscopy (XPS) and optical waveguide lightmode spectroscopy (OWLS). VASE allows for a fast and sensitive determination of total film thickness. XPS provides chemical information of the studied surfaces. OWLS was used to monitor in situ the degradation kinetics in real time. The non-fouling properties of the copolymer films before and after stability test were evaluated from protein (human serum) resistance test.
2.1 Water and Chemical Products
2.1.1 PLL, PLL-PEG, and PLL-PMOXA
PLL was purchased from Sigma-Aldrich. PLL-PEG polymers were purchased from Surface Solutions AG (Zurich, Switzerland). PLL-PMOXA polymers were synthesized and characterized using matrix-assisted laser desorption/ionization-time of flight (MALDI-ToF) and proton nuclear magnetic resonance spectroscopy (1H NMR) following the previously published protocol [21, 39].
All stability test solutions were prepared by using ultrapure water. Ultrapure water was produced in a Milli-Q system Gradient A 10 from Millipore (Zug, Switzerland). The system is equipped with Elix3 that removes 95–99% of inorganic ions, 99% of dissolved organic compounds, bacteria, and particulates.
2.1.3 H2O2, NaCl, and HEPES
Hydrogen peroxide 30% H2O2 (perhydrol*) pro analysis (p.a) was purchased from Merck KGaA (Damstadt, Germany). It contains impurities such as ≤40 ppm free acid as H2SO4, ≤50 ppm nonvolatile matter, and diverse metal impurities such as Al, Ca, Na, etc., in which each of them present in a concentration of <0.5 ppm. NaCl was purchased from Sigma-Aldrich. HEPES (4-(2-hydroxyethyl)-1-piperazine ethane sulfonic acid) and other chemicals used for the preparation of physiological solution were purchased from Fluka (Buchs, Switzerland).
2.1.4 Human Serum Proteins
Human serum proteins (Precinorm U) were purchased from Roche Diagnostics GmbH (Mannheim, Germany). The lyophilized powder was dissolved in ultrapure water as recommended by the supplier. The solution contains human serum components, along with enzymes and other additives, with a total concentration of approximately 70 mg/ml, which is about equal to that found in clinically normal human serum. The solution was divided into 0.5 ml aliquots in eppendorfs and stored in a −20°C freezer. The serum solution was defrosted at room temperature prior to use.
2.2 Stability Test Solutions
All stability test solutions were freshly prepared before the stability test.
2.2.1 A Solution of 10 mM H2O2
10 mM H2O2 solution was prepared by dilution of the commercially available hydrogen peroxide 30% H2O2 (perhydrol*) pro analysis (p.a) from Merck KGaA in ultrapure water. The pH of the solution was approximately 7 as measured with pH paper.
2.2.2 A Solution of 160 mM Ion Concentration (10 mM HEPES + 150 mM NaCl)
10 mM HEPES solution with additional 150 mM NaCl was prepared by dissolving the commercially available HEPES from Fluka and NaCl from Sigma-Aldrich in ultrapure water. The pH of the solution was then measured using pH-meter and adjusted to pH 7.4 by addition of 6 M NaOH. This solution is also called HEPES2 buffer.
2.2.3 A Physiological Solution in the Presence of Oxidative Substance (10 mM H2O2 + 10 mM HEPES + 150 mM NaCl)
The solution was prepared by dilution of the commercially available hydrogen peroxide 30% H2O2 (perhydrol*) pro analysis (p.a) from Merck KGaA in the solution of 10 mM HEPES + 150 mM NaCl described above.
2.3 Substrate and Surface Modification Protocol
The silicon wafers for VASE and XPS investigations were purchased from Si-Mat Silicon Materials (Landsberg, Germany). The silicon wafers were sputter-coated with a 21-nm-thick Nb2O5 layer (reactive magnetron sputtering, Paul Scherrer Institute, Villigen, Switzerland). The waveguides (OW2400) for OWLS experiments were purchased from Microvacuum Ltd. (Budapest, Hungary). Each waveguide consists of a 0.5-mm-thick AF45 glass substrate and a 200-nm-thick Si0.75Ti0.25O2 waveguiding layer on the surface. An additional 6-nm-thick Nb2O5 layer was sputter-coated on the top of the waveguiding layer.
2.3.2 Surface Modification
Prior to the copolymer adsorption, Nb2O5 substrates were ultrasonicated for 2 × 10 min in toluene (Fluka) followed by 2 × 10 min of ultrasonication in 2-propanol (Fluka) and blow-drying under a stream of nitrogen. Nb2O5 substrates were subsequently cleaned by oxygen plasma treatment for 2 min (Plasma cleaner/sterilizer PDC-32G, Harrick scientific products Inc.).
Polymer films were prepared by dip and rinse protocol . Briefly, for ex situ experiments using VASE and XPS techniques, the polymers were dissolved at 0.1 mg/ml concentration in a filtered (0.22 μm) HEPES buffer solution containing 10 mM HEPES supplemented with 150 mM NaCl and adjusted to pH 7.4 (HEPES2). Then, 50 μl of copolymer solutions were placed onto the freshly pre-cleaned substrates completely covering their surfaces. Polymer adsorption was allowed to proceed for around 2 h, followed by extensive washing with ultrapure water and blow-drying under a stream of nitrogen. The surface modification protocol for in situ experiments using OWLS techniques is described below.
2.4 Stability Test Protocol
Stability of the (ex situ) prepared films was tested by immersion in different stability test solutions at room temperature in γ-sterilized polystyrene cell-plate boxes (TPP, test plates, 92024). To protect the samples from daylight, the cell-plate boxes were wrapped with a piece of aluminum foil. The immersion time was varied between 5 h and 2 weeks. Sterile condition during stability experiments was assured by filtering all solutions before use with a 0.22 μm filter. Moreover, the preparation of copolymer films and the stability tests were performed in a sterile flow box.
At the end of stability test, the samples were rinsed with ultrapure water and dried under nitrogen stream prior to their characterizations of thickness (VASE) and surface composition (XPS) for the detection of polymer degradation. The copolymer films were subsequently re-hydrated by placing ultrapure water on top of the samples, fully covering the surface, for 30 min. The non-fouling properties of the copolymer films after stability tests were then evaluated by exposing the films to human serum solution for 15 min, followed by rinsing with ultrapure water, and drying under nitrogen stream. The change of film thickness after exposure to serum proteins was measured by VASE. The stability test protocol for in situ experiments using OWLS technique is described below.
2.5 Characterization Techniques
The dry copolymer film thickness was measured in air by variable angle spectroscopic ellipsometry (VASE) using the M-2000F variable angle spectroscopic ellipsometer (J. A. Woollam Co., Inc.). The measurements were performed at 65°, 70° and 75° relative to the surface normal, under ambient conditions and in the spectral range of 370–1,000 nm. Ellipsometry data were fitted with a multilayer model using the custom analysis software (WVASE 32) and a Cauchy model (A n = 1.45, B n = 0.01, C n = 0) [49, 50] to obtain the dry thickness of the adsorbed polymer layers.
A Sigma2 instrument (Thermo Fisher Scientific, Loughborough, Great Britain) was utilized for routine XPS experiments in this study. The Sigma2 is equipped with a UHV chamber (pressure <10−6 Pa during measurements) and an Al-Kα non-monochromated X-ray source (300 W, hν = 1486.6 eV) illuminating the sample at an angle of 54° to the surface normal. A hemispherical analyzer is mounted at 0° with respect to the surface normal, thus operating at the magic source-analyzer angle, which eliminates the need for angular-distribution correction.
The spot size of the analyzed area (large-area mode) was 400 μm, and the results therefore represented a laterally averaged chemical composition. Standard measurements comprised averages over nine (for C, and N) or three (for Nb and O) scans for each element with pass energies of 25 eV, as well as survey scans with pass energy of 50 eV. The dwell time was left at 100 ms at all times, resulting in 3–5 min measurement time per spot for each element, accumulating to about 30 min for a complete elemental scan on each measuring position.
OWLS measurements were performed on an OWLS 110 with BioSense 2.2 software from Microvacuum Ltd. (Budapest, Hungary). OWLS allows for quantitative in situ monitoring of copolymer and serum adsorption in a flow-through cell (typical cell dimension: 8 × 2 × 1 mm3 or 16 μl-volume). OWLS experiments were performed following previously described procedure [19, 20], i.e. by sequential injections of solutions.
Incubation of the Nb2O5-coated waveguides in copolymer solutions for 30–120 min resulted in the formation of a saturated layer. Subsequently, a stability test solution was injected followed by further 5 h incubation. Incubation of the (degraded) films in full human serum for 15 min followed by rinsing allowed determination of the adsorbed protein mass. The experiments were repeated three times for each copolymer.
3 Results and Discussion
3.1 Polymer Films
PLL, PLL-PEG and PLL-PMOXA polymers were immobilized onto Nb2O5 surfaces via electrostatic interaction between the negatively charged substrate and positively charged free lysine residues of the PLL-backbone [19, 20]. Both copolymer systems have identical PLL backbones (20 kDa as HBr salts, with approximately 96 lysine repeating units).
Side chain molecular weights were chosen for the purpose of comparing PMOXA and PEG polymer having approximately the same monomer repeating units (polymerization degree). PMOXA 4 kDa with approximately 44 MOXA-mers per PMOXA chain is comparable to PEG 2 kDa with approximately 50 EG-mers per PEG chain. Similarly, PMOXA 8 kDa (approximately 90 MOXA-mers per PMOXA chain) is comparable to PEG 5 kDa (approximately 113 EG-mers per PEG chain).
The copolymers used in this study were PLL-PMOXA4 (α = 0.33), PLL-PMOXA8 (α = 0.25), PLL-PEG2 (α = 0.31) and PLL-PEG5 (α = 0.28), with the following notation: PLL-PEG5 (α = 0.28) refers to PEG with molecular weight of 5 kDa, grafted to PLL with molecular weight of 20 kDa (as HBr salt), and a PEG grafting density of 0.28 PEG/lysine and correspondingly for the other copolymers.
3.2 Characterization of the Initial Polymer Films
Characteristics of the four copolymers and the films after surface assembly: molecular weights, film thicknesses, adsorbed masses, surface densities of molecules, side chains, and monomers
PLL-PMOXA4 (α = 0.33)
PLL-PMOXA8 (α = 0.25)
PLL-PEG2 (α = 0.31)
PLL-PEG5 (α = 0.28)
Molecular weight of polymer (kDa)
Polymer thickness (nm)
2.07 ± 0.17
2.36 ± 0.04
1.30 ± 0.03
2.47 ± 0.10
Polymer mass on the surface (ng/cm2)
210 ± 10
240 ± 28
152 ± 16
245 ± 75
Molecular surface density (×10−3 nm−2),
8.7 ± 0.9
7.1 ± 0.8
12.7 ± 1.3
10.2 ± 3.2
Side chain surface density (nm−2)
0.28 ± 0.01
0.17 ± 0.02
0.39 ± 0.05
0.27 ± 0.09
Monomer surface density (nm−2)
14 ± 1
16 ± 2
18 ± 2
31 ± 10
Serum thickness on initial polymer layer (nm)
0.03 ± 0.03
0.02 ± 0.03
0.02 ± 0.01
0.01 ± 0.01
6.4 ± 0.3
Serum mass on initial polymer layer (ng/cm2)
1 ± 2
0 ± 1
2 ± 3
0 ± 1
223 ± 22
The adsorbed polymer mass for PLL-PEG2 is somewhat lower due to the lower side-chain molecular weight. The maximum in adsorbed copolymer mass for each system is typically reached at a medium grafting density where an almost defect-free monolayer is formed on the surface. This maximum corresponds to a minimum in protein adsorption and is a result of the interplay of increasing side chain density and accordingly adsorbed mass at low to medium grafting densities and an increasingly weak binding to the surface due to increasing steric repulsion of the bottle brushes from the surface and decreasing backbone charge density with increasing grafting density at high grafting densities. These effects have been discussed in detail in earlier studies [9, 19].
Before exposure to stability test solution, all PLL-PMOXA and PLL-PEG copolymers resisted serum (Table 1, entries 7 and 8). Serum protein adsorption on the unprotected Nb2O5 and PLL interfaces was found to be in a good agreement with previously reported results .
XPS data for bare Nb2O5, and Nb2O5 coated with PLL-PEG2 (α = 0.31), PLL-PEG5 (α = 0.28), PLL-PMOXA4 (α = 0.33), and PLL-PMOXA8 (α = 0.28): element orbital, the assignment of components with their binding state, binding energy, and theoretical and experimental composition (in atomic percentages)
Binding energy (eV)
68.6 ± 1.1
Nb ox 5/2
32.2 ± 0.1
Nb ox 3/2
PLL-PEG2 (α = 0.31)
8.3 ± 0.5
C–C–O and C–C–N
51.9 ± 0.9
2.8 ± 0.4
9.5 ± 0.4 a
24.1 ± 0.8
2.5 ± 0.1
0.8 ± 0.1
PLL-PEG5 (α = 0.28)
5.4 ± 0.5
C–C–O and C–C–N
55.8 ± 1.3
2.1 ± 0.2
6.3 ± 1.3 a
28.5 ± 0.7
1.6 ± 0.1
0.2 ± 0.2 b
PLL-PMOXA4 (α = 0.33)
H3C–C=O and C–C–C
14.7 ± 0.3
36.1 ± 0.3
13.4 ± 0.3
19.1 ± 0.4
15.5 ± 0.2
1.0 ± 0.2
PLL-PMOXA8 (α = 0.25)
H3C–C=O and C–C–C
16.2 ± 0.4
34.3 ± 1.2
13.3 ± 0.4
19.3 ± 1.5
15.8 ± 0.4
Characteristics of the four copolymers and the films after surface assembly: C/Nb and N/Nb ratios before stability tests, evaluated from XPS analysis
PLL-PMOXA4 (α = 0.33)
PLL-PMOXA8 (α = 0.25)
PLL-PEG2 (α = 0.31)
PLL-PEG5 (α = 0.28)
1.87 ± 0.06
2.26 ± 0.05
1.32 ± 0.07
2.22 ± 0.08
0.22 ± 0.01
0.49 ± 0.03
0.60 ± 0.01
0.07 ± 0.00
0.06 ± 0.01
3.3 Stability Study
3.3.1 Stability upon Exposure to 10 mM H2O2
For both types of copolymers, a correlation between degradation of copolymer films and their biopassive function was observed, i.e. serum thickness increased with time (or as the copolymer thickness decreased) (Fig. 2b, c). A high adsorbed serum thickness (~3.5 nm) on PLL-PEG2 film was detected when the remaining thickness of the polymeric layer was about 60% (compare Fig. 2a, b, at 168 h). At the same copolymer remaining thickness (~60%), however, PLL-PEG5 still showed a low adsorbed serum thickness of ~0.2 nm (Fig. 2c). The degradation kinetics of PLL-PEG2 and PLL-PEG5 as judged from the relative thickness reduction (Fig. 2a) were similar; however the PLL-PEG5 film had a significantly higher initial EG monomer surface density (Table 1, entry 6). Therefore, for the same remaining thickness of 60%, significantly less serum proteins were adsorbed on the PLL-PEG5 compared to the PLL-PEG2 film. For PLL-PMOXA4 and PLL-PMOXA8, no significant difference in film degradation rate was observed (Fig. 2a), and only slight difference was observed in serum adsorption where PLL-PMOXA4 showed slightly higher values (Fig. 2c). This phenomenon is probably a consequence of the initial monomer surface density of PLL-PMOXA4 being slightly lower than that of PLL-PMOXA8 (Table 1, entry 6).
3.3.2 Stability upon Exposure to 160 mM Ion Concentration (10 mM HEPES + 150 mM NaCl, pH = 7.4)
A solution of 160 mM ion concentration containing 10 mM HEPES + 150 mM NaCl was used to mimic the physiological solution. Copolymer films were exposed to this solution for the same duration as applied in the experiments with 10 mM H2O2 solution.
The same experimental conditions resulted in 10–20 and 40–50% C/Nb loss for PLL-PEG films after 1 day and 1 week, respectively (Fig. 5b). Only 5–10 and 15–25% N/Nb reduction (N is characteristic of PLL but not PEG) for the same duration indicates that the film degradation was not simply due to copolymer detachment, but that PEG chains degraded preferentially to PLL. In a control experiment, PLL film showed only ~5% decrease of N/Nb over 1 week, in the same stability test solution (data not shown).
In Fig. 6, both PLL-PEG2 and PLL-PEG5 showed ~20% decrease in the PEG/PLL ratio after 1 day of exposure to 10 mM HEPES + 150 mM NaCl, indicating PEG degradation preferentially to PLL. The PEG/PLL ratio further decreased over 1 week of stability test, resulting in a total of ~40% of PEG loss, relative to the initial value. This is in a close agreement with previously published result of Zoulalian et al.  where ~30% of PEG loss was observed upon stability test of PLL-PEG2- modified Nb2O5 surfaces in the same solution for 1 week.
3.3.3 Stability upon Exposure to Physiological Solution in the Presence of Oxidative Substance (10 mM H2O2 + 10 mM HEPES + 150 mM NaCl, pH = 7.4)
In addition to the stability tests in 10 mM H2O2 and physiological solution (10 mM + 150 mM NaCl), a stability study of PLL-PEG and PLL-PMOXA films in the mixture of 10 mM H2O2 and 10 mM HEPES + 150 mM NaCl was performed.
While PLL-PMOXA films remained more stable (only 5–10% decrease), 40–45% decrease in thickness was observed for PLL-PEG films in the mixture of 10 mM H2O2 + 10 mM HEPES + 150 mM NaCl (Fig. 7). This value shows that the degradation of PLL-PEG films proceeded faster in this solution when compared to either H2O2 or 10 mM HEPES + 150 mM NaCl solution.
3.3.4 Quantitative Correlation VASE, XPS, and OWLS Results, as well as between Copolymer Film Thickness and Serum Thickness and Mass
3.3.5 Hypotheses on the Degradation Mechanism of PLL-PEG and PLL-PMOXA Films
As shown and discussed above, incubation of copolymer films in the different stability test solutions lead to different levels of film degradation. Here we propose hypotheses on the degradation mechanisms.
10 mM H2O2 solution was used to simulate in vivo conditions [upon phagocytosis, macrophages produce superoxide (O2−) and hydrogen peroxide (H2O2)]. These reactive oxygen species can produce highly toxic radicals such as hydroxyl radicals (HO*) which may directly promote the oxidative polymer degradation at the surface .
Degradation of copolymer films in 10 mM HEPES + 150 mM NaCl solution is less intuitive. HEPES has been reported to strongly take part in radical reactions [53–55]. Most importantly, under the influence of light, HEPES-containing cell culture solutions have been found to form hydrogen peroxide in a light-dose dependent manner. Furthermore, the piperazine ring as well as the alcohol function of HEPES can act as a hydroxyl radical scavenger and HEPES can form nitroxide-type radicals at the piperazine function. These findings suggest that HEPES might promote the oxidative degradation of polymers in biological media not protected from oxygen and light. In this study, all samples were protected from light by covering the containers with aluminum foil during the stability test period. However, solutions were not degassed and it is likely that reactive oxygen species were formed during the preparation of ultrapure water which involves UV light exposure to degrade organic contaminants. These reactive oxygen species might subsequently have reacted with HEPES. This could also explain the observed synergistic action of HEPES and hydrogen peroxide containing solutions on polymer degradation.
An alternative explanation for film instability would be the detachment of entire graft copolymer molecules from the Nb2O5 substrate. We have ruled out this mechanism by XPS measurements showing that the PEG signal decreases faster than the PLL signal during exposure to the stability test solutions. This shows that the graft copolymer side chains degrade while the backbone polymer, PLL, remains electrostatically bound to the substrate.
4 Summary and Conclusions
We found that the degradation rates differed significantly between PLL-PEG and PLL-PMOXA films, with the latter showing better stability in all model environments tested in this work. The higher stability of PLL-PMOXA films compared to PLL-PEG films was proven by surface analysis performed using three complementary techniques (VASE, XPS, and OWLS). Despite the similar architecture of the PMOXA- and PEG-copolymers used in terms of grafting density, initial adsorbed mass (thickness), molecular-, side chain-, and monomer-surface density, as well as initial C/Nb ratio, films prepared from PLL-PMOXA always showed considerably higher stability when compared to PLL-PEG.
Although the film degradation mechanisms could not be fully deciphered, the XPS analysis supports the hypothesis that the stability of PLL-PMOXA and PLL-PEG films is primarily limited by the side chain (PMOXA or PEG) degradation. Further investigations are needed to gain a deeper understanding on the exact degradation mechanism as well as on the degradation products of PLL-PMOXA and PLL-PEG films.
In conclusion, this study suggests PMOXA coatings as a potent alternative to PEG coatings due to their higher stability in physiological and oxidative environments and related prolonged resistance to protein fouling. The findings of this work are believed to also be relevant in the context of application of polymeric, anti-fouling surfaces in vivo. Degradation of polymers that result in the production of reactive species such as radicals, hydrogen peroxide or hydrogen superoxide are likely to contribute to an inflammatory and foreign body response by the host resulting in encapsulation of the biomaterial or medical device by the formation of an avascular capsule. It would be interesting in our view to investigate whether PMOXA-modified biomaterial surfaces would result in a more natural healing and integration when implanted in (soft) tissue in comparison to PEG. Complementary investigations are underway.
We acknowledge financial support from the Swiss National Science Foundation (SNSF, Project 200021-116163) and Materials for the Life Sciences Unit of the Swiss Competence Center for Materials Research and Technology of the ETH Domain.
- Kenausis GL, Vörös J, Elbert DL, Huang N, Hofer R, Ruiz-Taylor L, Textor M, Hubbell JA, Spencer ND (2000) J Phys Chem B 104:3298View ArticleGoogle Scholar
- Maddikeri RR, Tosatti S, Schuler M, Chessari S, Textor M, Richards RG, Harris LG (2008) J Biomed Mater Res Part A 84A:425View ArticleGoogle Scholar
- Lussi JW, Falconnet D, Hubbell JA, Textor M, Csucs G (2006) Biomaterials 27:2534View ArticleGoogle Scholar
- Roosjen A, Norde W, van der Mei HC, Busscher HJ (2006) Program Colloid Polym Sci 132:138View ArticleGoogle Scholar
- Malmsten M, Emoto K, Van Alstine JM (1998) J Colloid Interface Sci 202:507Google Scholar
- McPherson T, Kidane A, Szleifer I, Park K (1998) Langmuir 14:176View ArticleGoogle Scholar
- Sofia SJ, Premnath V, Merrill EW (1998) Macromolecules 31:5059View ArticleGoogle Scholar
- Dalsin JL, Lin L, Tosatti S, Vörös J, Textor M, Messersmith PB (2005) Langmuir 21:640View ArticleGoogle Scholar
- Pasche S, DePaul SM, Vörös J, Spencer ND, Textor M (2003) Langmuir 19:9216View ArticleGoogle Scholar
- Feller LM, Cerritelli S, Textor M, Hubbell JA, Tosatti SGP (2005) Macromolecules 38:10503View ArticleGoogle Scholar
- Roosjen A, Kaper HJ, van der Mei HC, Norde W, Busscher HJ (2003) Microbiology 149:3239View ArticleGoogle Scholar
- Koh WG, Revzin A, Simonian A, Reeves T, Pishko M (2003) Biomed Microdevices 5:11View ArticleGoogle Scholar
- Park JH, Bae YH (2003) J Appl Polym Sci 89:1505View ArticleGoogle Scholar
- Harris LG, Tosatti S, Wieland M, Textor M, Richards RG (2004) Biomaterials 25:4135View ArticleGoogle Scholar
- Marie R, Beech JP, Vörös J, Tegenfeldt JO, Hook F (2006) Langmuir 22:10103View ArticleGoogle Scholar
- Cringus-Fundeanu I, Luijten J, van der Mei HC, Busscher HJ, Schouten AJ (2007) Langmuir 23:5120View ArticleGoogle Scholar
- Abd El-Mohdy H, Ghanem S (2009) J Polym Res 16:1View ArticleGoogle Scholar
- Statz AR, Meagher RJ, Barron AE, Messersmith PB (2005) J Am Chem Soc 127:7972View ArticleGoogle Scholar
- Konradi R, Pidhatika B, Mühlebach A, Textor M (2008) Langmuir 24:613View ArticleGoogle Scholar
- Pidhatika B, Möller J, Vogel V, Konradi R (2008) CHIMIA Int J Chem 62:264View ArticleGoogle Scholar
- Pidhatika B, Möller J, Benetti EM, Konradi R, Rakhmatuliina E, Muehlebach A, Zimmermann R, Werner C, Vogel V, Textor M (2010) Biomaterials 31:9462View ArticleGoogle Scholar
- Veronese FM, Pasut G (2005) Drug Discov Today 10:1451View ArticleGoogle Scholar
- Pasut G, Veronese FM (2009) Adv Drug Deliv Rev 61:1177View ArticleGoogle Scholar
- Matthews SJ, McCoy C (2004) Clin Ther 26:991View ArticleGoogle Scholar
- Donbrow M (1987) In: Schick MJ (ed) Nonionic surfactants: physical chemistry, vol. 23. Marcel Dekker, New YorkGoogle Scholar
- Shen M, Martinson L, Wagner MS, Castner DG, Ratner BD, Horbett TA (2002) J Biomater Sci Polym Ed 13:367View ArticleGoogle Scholar
- Roosjen A, Vries Jd, van der Mei HC, Norde W, Busscher HJ (2005) J Biomed Mater Res B Appl Biomater 73B:347View ArticleGoogle Scholar
- McGary-Jr CW (1960) J Polym Sci Part A Polym Chem 46:51Google Scholar
- Han S, Kim C, Kwon D (1995) Polym Degrad Stab 47:203View ArticleGoogle Scholar
- Branch DW, Wheeler BC, Brewer GJ, Leckband DE (2001) Biomaterials 22:1035View ArticleGoogle Scholar
- Sharma S, Johnson RW, Desai TA (2003) Langmuir 20:348View ArticleGoogle Scholar
- Zhang F, Kang ET, Neoh KG, Wang P, Tan KL (2001) J Biomed Mater Res 56:324View ArticleGoogle Scholar
- Bindra DS, Williams TD, Stella VJ (1994) Pharm Res 11:1060View ArticleGoogle Scholar
- Knop K, Hoogenboom R, Fischer D, Schubert US (2010) Angew Chem Int Ed 49:6288View ArticleGoogle Scholar
- Zoulalian V, Zürcher S, Tosatti S, Textor M, Monge S, Robin JJ (2009) Langmuir 26:74View ArticleGoogle Scholar
- Imamura S-i, Tonomura Y, Kawabata N, Kitao T (1981) Bull Chem Soc Jpn 54:1548View ArticleGoogle Scholar
- Chilkoti A, Dreher MR, Meyer DE, Raucher D (2002) Adv Drug Deliv Rev 54:613View ArticleGoogle Scholar
- Yamaoka T, Tabata Y, Ikada Y (1994) J Pharm Sci 83:601View ArticleGoogle Scholar
- von Erlach T, Zwicker S, Pidhatika B, Konradi R, Textor M, Hall H, Lühmann T (2011) Biomaterials 32:5291View ArticleGoogle Scholar
- Woodle MC, Engbers CM, Zalipsky S (1994) Bioconjugate Chem 5:493View ArticleGoogle Scholar
- Hoogenboom R (2009) Angew Chem Int Ed 48:7978View ArticleGoogle Scholar
- Zalipsky S, Hansen CB, Oaks JM, Allen TM (1996) J Pharm Sci 85:133View ArticleGoogle Scholar
- Cheon Lee S, Kim C, Chan Kwon I, Chung H, Young Jeong S (2003) J Control Release 89:437View ArticleGoogle Scholar
- Broz P, Benito SM, Saw C, Burger P, Heider H, Pfisterer M, Marsch S, Meier W, Hunziker P (2005) J Control Release 102:475View ArticleGoogle Scholar
- Ranquin A, Versees W, Meier W, Steyaert J, Van Gelder P (2005) Nano Lett 5:2220View ArticleGoogle Scholar
- Zaikov GE (1985) Polym Rev (Philadelphia, PA, US) 25: 551Google Scholar
- Williams DF (1982) J Mater Sci 17:1233View ArticleGoogle Scholar
- Ali SAM, Doherty PJ, Williams DF (1994) J Appl Polym Sci 51:1389View ArticleGoogle Scholar
- Statz AR, Kuang J, Ren C, Barron AE, Szleifer I, Messersmith PB (2009) Biointerphases 4: FA22Google Scholar
- Tompkins HG, McGahan WA (1999) Spectroscopic ellipsometry and reflectometry: a user’s guide. Wiley, New YorkGoogle Scholar
- Moulder JF, Chastain J (1992) Handbook of X-ray photoelectron spectroscopy: a reference book of standard spectra for identification and interpretation of XPS data. Physical Electronics Division, Perkin-Elmer Corp., Eden PrairieGoogle Scholar
- Vörös J, Ramsden JJ, Csúcs G, Szendro I, De Paul SM, Textor M, Spencer ND (2002) Biomaterials 23:3699View ArticleGoogle Scholar
- Zigler J, Lepe-Zuniga J, Vistica B, Gery I (1985) In Vitro Cell Develop Biol Plant 21:282View ArticleGoogle Scholar
- Lepe-Zuniga JL, Zigler JS Jr, Gery I (1987) J Immunol Methods 103:145View ArticleGoogle Scholar
- Grady JK, Chasteen ND, Harris DC (1988) Anal Biochem 173:111View ArticleGoogle Scholar
This article is published under license to BioMed Central Ltd. Open Access This article is distributed under the terms of the Creative Commons Attribution License which permits any use, distribution and reproduction in any medium, provided the original author(s) and source are credited.