Article pubs.acs.org/Biomac
Aggregation Behavior of Amphiphilic p(HPMA)-co-p(LMA) Copolymers Studied by FCS and EPR Spectroscopy Mirjam Hemmelmann,† Dennis Kurzbach,‡ Kaloian Koynov,‡ Dariush Hinderberger,*,‡ and Rudolf Zentel*,† †
Institute of Organic Chemistry, Johannes Gutenberg University Mainz, Duesbergweg 10-14, 55099 Mainz, Germany Max Planck Institute for Polymer Research Mainz, Ackermannweg 10, 55128 Mainz, Germany
‡
S Supporting Information *
ABSTRACT: A combined study of fluorescence correlation spectroscopy and electron paramagnetic resonance spectroscopy gave a unique picture of p(HPMA)-co-p(LMA) copolymers in aqueous solutions, ranging from the size of micelles and aggregates to the composition of the interior of these self-assembled systems. P(HPMA)-co-p(LMA) copolymers have shown high potential as brain drug delivery systems, and a detailed study of their physicochemical properties can help to elucidate their mechanism of action. Applying two complementary techniques, we found that the self-assembly behavior as well as the strength of hydrophobic attraction of the amphiphilic copolymers can be tuned by the hydrophobic LMA content or the presence of hydrophobic molecules or domains. Studies on the dependence of the hydrophobic lauryl side chain content on the aggregation behavior revealed that above 5 mol % laury side-chain copolymers self-assemble into intrachain micelles and larger aggregates. Above this critical alkyl chain content, p(HPMA)-co-p(LMA) copolymers can solubilize the model drug domperidone and exhibit the tendency to interact with model cell membranes.
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INTRODUCTION In the past two decades, poly(N-(2-hydroxypropyl)-methacrylamide), p(HPMA), has been widely used for medical applications, especially for polymer−drug conjugates, a concept that was first introduced by Helmut Ringsdorf in the 1970s.1,2 The general idea is that water-soluble, nontoxic, and nonimmunogenic polymers can improve the body distribution and biocompatibility of low-molecular-weight drugs by either encapsulating or covalently binding them to a water-soluble polymer. Many of the early p(HPMA) polymers were in fact hydrophobically modified water-soluble polymers because the conjugated drug, for example, doxorubicin, was hydrophobic.2−4 A systematic study of amphiphilic copolymers of hydrophilic p(HPMA) modified with randomly distributed hydrophobic lauryl methacrylate (LMA) started recently.5 These amphiphilic copolymers self-assemble into aggregates in aqueous solution. In addition, they exhibit increased cellular uptake compared with p(HPMA) homopolymers and p(HPMA)-b-p(LMA) block copolymers.6 Studies at the air− water interface showed that they are surface-active and able to form artificial membranes.7 A recent study on the pharmacokinetics and body distribution of radioactively labeled copolymers revealed that the copolymers combine a long blood-circulation time with a low liver accumulation.8,9 Furthermore, we have shown that amphiphilic p(HPMA)-cop(LMA) copolymers are able to mediate drug delivery over © 2012 American Chemical Society
biological barriers into the brain, that is, the enhanced delivery of the Pgp substrate domperidone into the brain of mice.10 In an in vitro study using a blood−brain barrier (BBB) model of human brain microvascular endothelial cells, we also showed the enhanced transport of another Pgp substrate, rhodamine 123, using the copolymer as a transporter.11 These properties might well be related to the amphiphilic behavior of the copolymers and their aggregate formation. In general, the synthesis of such random amphiphilic polymer systems is challenging because copolymerization of two monomers of different polarities may be accompanied by microphase separation in the reaction mixture, leading to inhomogeneous copolymerization. This problem can be circumvented by RAFT-copolymerization of pentafluorophenyl methacrylate (PFPMA) and LMA, leading to well-defined, allhydrophobic reactive precursors polymers.12 Subsequent postpolymerization modification with 2-hydroxypropyl amine provides defined amphiphilic p(HPMA)-co-p(LMA) copolymers.5 Via this defined synthesis approach, the architecture as well as the amount of hydrophobic side chains can be controlled. Furthermore, additional functionalities, such as fluorescent and spin labels, can be introduced. Received: August 29, 2012 Revised: October 20, 2012 Published: November 16, 2012 4065
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General Synthetic Route of Statistic Copolymers. The synthesis of the here-described polymers was performed by analogy to previously published procedures4,5 with some alterations. Synthesis of 4-Cyano-4-((thiobenzoyl)sulfanyl)pentanoic Acid. The 4-cyano-4-((thiobenzoyl)sulfanyl)pentanoic acid (CTP) was used as CTA and synthesized according to literature.21 Synthesis of Pentafluorophenyl Methacrylate. PFPMA was prepared according to literature.12,22 RAFT Polymerization to RE-Precursor Polymers. General Synthesis of RE-Precursor Polymer of H1. In a typical reaction, 3 g (12 mmol) PFPMA and 45 mg (0.18 mmol) CTP with AIBN as an initiator (molar ratio AIBN/CTP: 1/8) were dissolved in 5 mL of dry dioxane in a Schlenk tube. After three freeze−vacuum−thaw cycles, the reaction was stirred at 65 °C overnight. p(PFPMA) was precipitated three times into hexane, isolated by centrifugation, and dried for 12 h at 40 °C under vacuum. A pink powder was obtained with a yield of 55%. 1H NMR (CDCl3): (300 MHz, CDCl3) δ: 1.25− 1.75 (br), 2.00−2.75 (br s). 19F-NMR (400 MHz, CDCl3) δ: −162.03 (br), −157 (br), −152 to −150 (br). General Synthesis of RE-Precursor Polymer of Random Copolymers C1(e)−C10(e). In a typical reaction, 3 g (12 mmol) PFPMA, 41 mg (0.16 mmol) CTP with AIBN as an initiator (molar ratio AIBN/CTP:1/8), and the respective amounts of LMA (C1: 1 mol %; C5: 5 mol %; C10: 10 mol %) were dissolved in 5 mL of dry dioxane in a Schlenk tube. After three freeze−vacuum−thaw cycles, the reaction was stirred at 65 °C overnight. p(PFPMA)-co-p(LMA) was precipitated three times in hexane, isolated by centrifugation, and dried for 12 h at 40 °C under vacuum. A pink powder was obtained with a yield of 60%. 1H NMR (300 MHz, CDCl3) δ: 0.86(br t), 1.20− 1.75 (br), 2.00−2.75 (br s). 19F-NMR (400 MHz, CDCl3) δ: −162 (br), −157 (br), −152 to −150 (br). Post-Polymerization Modification to Hydrophilic H1 and Amphiphilic C1(e)−C10(e). Post-Polymerization Modification to Homo p(HPMA) H1. We dissolved 254 mg of the precursor polymer (Mn = 24 000 g/mol) in absolute dioxane, and 55 mg (1 mmol) 2hydroxypropylamine (HPA) together with 204 mg (2 mmol) triethylamine (TEA) were added to the reaction mixture. The reaction continued overnight at 60 °C. To ensure complete removal of REgroups, an excess of again 55 mg HPA and 204 mg TEA was added to the reaction mixture. Completion of the reaction was determined using 19 F NMR. The final polymer was precipitated three times in diethyl ether, dissolved in 1 mL of DMSO, and dialyzed against deionized water. A colorless fluffy powder was obtained with a yield of 83% after lyophilization. 1H NMR (400 MHz, d.DMSO) δ: 0.90−1.40 (br), 1.6−2.20 (br), 2.75−3.10 (br), 3.50−3.80 (br), 4.60−4.80 (br). For the fluorescence labeled polymer 5 mg (0.01 mmol) of Oregon green cadaverine with 2 mg (2.0 × 10−5 mol), TEA were dissolved in dry DMSO and added to the reaction mixture in the first step. After stirring for 4 h at 60 °C, 55 mg (1 mmol) HPA and 204 mg (2 mmol) TFA were added to the reaction mixture, and the reaction was continued as described above. Purification from unbound dye was achieved using SEC (Sephadex G25) chromatography. Post-Polymerization Modification to Random p(HPMA)-cop(LMA) Copolymers C1(e)−C10(e). The reaction procedure is described for C10 as an example: 280 mg of the precursor polymer (Mn = 23 000 g/mol) was dissolved in absolute dioxane, and 48 mg (0.9 mmol) HPA together with 183 mg (1.8 mmol) TEA were added to the reaction mixture. The reaction continued overnight at 60 °C. To ensure complete removal of RE-groups, an excess of again 48 mg HPA and 183 mg TEA was added to the reaction mixture. Completion of the reaction was determined using 19F NMR. The final polymer was precipitated three times in diethyl ether, dissolved in 1 mL of DMSO, and dialyzed against deionized water. A colorless fluffy powder was obtained with a yield of 76% after lyophilization. 1H NMR (400 MHz, d.DMSO) δ: 0.85 (br t), 0.90−1.40 (br), 1.6−2.20 (br), 2.75−3.10 (br), 3.50−3.80 (br), 4.60−4.80 (br). For the fluorescence-labeled polymer 5 mg (0.01 mmol) of Oregon green cadaverine with 2 mg (2.0 × 10−5 mol), TEA was dissolved in dry DMSO and added to the reaction mixture in the first step. After stirring for 4 h at 60 °C, 48 mg (0.9 mmol) HPA and 183 mg (1.8
An important feature of p(HPMA)-co-p(LMA) copolymers with respect to their potential application as drug carrier systems is their ability to encapsulate hydrophobic drugs via hydrophobic interactions. In general, amphiphilic copolymers spontaneously self-assemble into micelle-like structures and aggregates in aqueous solutions. Whereas block copolymers are composed of well-separated hydrophilic and hydrophobic blocks, random copolymers contain many independent hydrophobic groups spread throughout the otherwise hydrophilic polymer chain. Under a physicochemical aspect, such polymers are classified as polysoaps.13,14 They are characterized by their property to form intrachain micelles.13 These intrachain micelles can further aggregate to form globules of closely packed micelles.15 The characteristics and the self-organization of polysoaps have been widely studied,13,15,16 but a full understanding is still missing. Aggregates formed by hydrophobically modified p(HPMA) copolymers might fall into this category because the formation of compound micelles has been described in former studies.5 In the present study, the self-assembly behavior and the aggregate structure of several amphiphilic p(HPMA)-cop(LMA) copolymers were investigated as a function of hydrophobic LMA content. Furthermore, the influence of encapsulating the hydrophobic model drug domperidone into the copolymers was evaluated. The characteristic aggregation behavior was analyzed on two different length scales. Fluorescence correlation spectroscopy (FCS) allows the measurement of the diffusion coefficient and size of fluorescently labeled single polymer chains as well as polymer aggregates.16 Electron paramagnetic resonance (EPR) spectroscopy elucidates the internal constitution of the polymer aggregates through a spin-probing approach, and spin-labeled polymers allow for the observation of interactions between p(HPMA)-co-p(LMA) and liposomes as model membranes.17−20 As a general trend, we found that the strength of hydrophobic attraction can be tuned by the LMA content or the presence of hydrophobic molecules or domains, leading to the formation of intrachain micelles and larger aggregates.
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MATERIALS AND METHODS
Materials. All chemicals/solvents were of reagent/analytical grade, as obtained from Sigma Aldrich and Acros Organics. Pentafluorophenol was obtained from Fluorochem (U.K.); Oregon Green cadaverine was purchased from Invitrogen. Dioxane was distilled over a sodium/potassium composition prior to use. LMA was distilled under reduced pressure to remove the stabilizer and stored at −18 °C. 2,2′-Azo-bis-(isobutyronitrile) (AIBN) was recrystallized from diethyl ether and stored at −18 °C. Characterization. 1H NMR spectra were obtained by a Bruker AC 300 spectrometer at 300 MHz. 19F-NMR analysis was carried out with a Bruker DRX-400 at 400 MHz. All measurements were carried out at room temperature, and spectroscopic data were analyzed using ACDLabs 9.0 1D NMR manager and MestReNova. The reactive ester polymers were dried overnight at 40 °C under vacuum and afterward analyzed via gel permeation chromatography (GPC). GPC was performed in tetrahydrofuran (THF) as solvent and equipped with: pump PU 1580, autosampler AS 1555, UV detector UV 1575, and RI detector RI 1530 from Jasco as well as a miniDAWN Tristar light-scattering detector from Wyatt. Columns were used from MZ Analysentechnik: MZ-Gel SDplus 106 Å 5 μm, MZ-Gel SDplus 104 Å 5 μm, and MZ-Gel SDplus 102 Å 5 μm. GPC data were evaluated using the software PSS WinGPC Unity from Polymer Standard Service Mainz. The flow rate was set to 1 mL/min with a temperature of 25 °C. 4066
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Scheme 1. Synthesis of Oregon-Green-Labeled Homopolymers, Spin-Labeled and Oregon-Green-Labeled Random Copolymers via RAFT Polymerization, and Post-Polymerization Modification
Here N is the average number of diffusing fluorescence species in the observation volume, f T and τT are the fraction and the decay time of the triplet state, τDi is the diffusion time of the ith species, f i is the fraction of component i, and S is the so-called structure parameter, S = z0/r0, where z0 and r0 represent the axial and radial dimensions of the confocal volume, respectively. Furthermore, the diffusion time, τDi, is related to the respective diffusion coefficient, Di, through Di = r02/ 4τDi.23 The experimentally obtained G(t) can be fitted, yielding the corresponding diffusion times and subsequently the diffusion coefficients of the fluorescent species. Finally, the hydrodynamic radii Rh can be calculated (assuming spherical particles) using the Stokes−Einstein relation: Rh = kBT/6πρD, where kB is the Boltzmann constant, T is the temperature, and ρ is the viscosity of the solution.19,23 CW EPR. Spectroscopy. CW EPR spectra at X-band (∼9.4 GHz) were measured on a Magnettech (Berlin, Germany) MiniScope MS200 benchtop CW EPR spectrometer with a variable-temperature cooling/heating unit (TC HO2). The sample volume was always large enough to fill the complete resonator volume in the probehead (>300 μL). Each sample was measured at 25 °C. Changes in mole fractions of species A or B after longer waiting periods were not detected. Typical experimental parameters were modulation amplitude of 0.06 mT and sweep width of 15 mT, yet note that Figures 4 and 5 do not show the complete sweep range of 15 mT. For improved representation of the spectra, mere baselines exhibiting low- and high-field parts were cut off. Sample Preparation. For spin-probing CW EPR, 1 v/v% of a 20 mM solution of the probe 16-DSA (4,4-dimethyl-oxazolidine-N-oxyl stearic acid) in ethanol was added to the polymer in phosphate buffered saline (PBS) to yield a spin probe concentration of 0.2 mM. For both approaches, with spin-labeled polymers and with spin probes, the polymer concentration was 3 mg/mL in all cases. Liposomes were prepared according to literature and extruded with a filter of 100 nm in size.24 EPR Data Analysis. All spectral simulations were performed with home-written programs in MATLAB (The MathWorks) employing the EasySpin toolbox for EPR spectroscopy.25 Simulations of CW EPR spectra in fluid solution were performed by using a model, which is
mmol) TEA were added to the reaction mixture, and the reaction was continued as described above. Purification from unbound dye was achieved using SEC (Sephadex G25) chromatography. For the synthesis of the spin-labeled polymer, 1.6 mg (9.3 × 10−6 mol) of 4-amino-TEMPO ((2,2,6,6-tetramethyl-piperidin-1-yl)oxyl) with 2 mg (2.0 × 10−5 mol) TEA was dissolved in dry DMSO and added to a mixture of 50 mg precursor polymer (Mn = 23 000 g/mol) in dioxane. After stirring for 4 h at 60 °C, 9.5 mg (0.16 mmol) HPA and 36.2 mg (0.36 mmol) TEA were added to the reaction mixture, and the reaction was continued as described above for the nonlabeled polymer. Fluorescence Correlation Spectroscopy. FCS experiments were performed using a commercial FCS setup (Zeiss, Germany) consisting of the module ConfoCor 2 and an inverted microscope model Axiovert 200 with a Zeiss C-Apochromat 40/1.2 W water immersion objective. The fluorophores were excited by an argon laser (λ = 488 nm), and the emission was collected after filtering with a LP505 long pass filter. For detection, an avalanche photodiode that enables single-photon counting was used. Eight-well, polystyrenechambered cover glass (Laboratory-Tek, Nalge Nunc International) was used as sample cell. The fluorescent-dye-labeled polymers were dissolved in phosphate buffer containing 1% abs DMSO (c = 1 mg/ mL) and diluted to the final concentration of 100 μg/mL. To calibrate the FCS observation volume, a dye with known diffusion coefficient was used, that is, Alexa Fluor 488. Each sample was tested in 10 measurement cycles with a total duration of 30 s. The time-dependent fluctuations of the fluorescence intensity dI(t) were recorded and analyzed by an autocorrelation function G(t) = 1 + /2. As has been shown theoretically for an ensemble of m different types of freely diffusing fluorescence species, G(t) has the following analytical form:23
⎡ fT −t / τ ⎤ 1 m ∑ G(t ) = 1 + ⎢1 + e T⎥ ⎢⎣ ⎥⎦ N i = 1 ⎡1 + 1 − fT ⎣⎢
t ⎤ ⎥ τDi ⎦
fi 1+
t S 2τDi
(1) 4067
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Table 1. Overview of HPMA-Based Homo- (H1) and Random Copolymers (C1−C10) and the Respective Spin-Labelled Polymers (C1e-C10e) (See Scheme 1) polymer
structure
H1 C1 C1e C5 C5e C10 C10e
homopolymer random copolymer random copolymer random copolymer random copolymer random copolymer random copolymer
mol % spin label
5 5 5
HPMA/LMA ratioa
Mn(RE) (g/mol)b
Mw(RE) (g/mol)b
Mn (g/mol)c
PDIb
100:0 99:1 99:1 95:5 95:5 90:10 90:10
24 000 23 000 23 000 23 500 23 500 23 000 23 000
28 000 28 000 28 000 28 000 28 000 27 000 27 000
13 600 13 700 13 700 13 500 13 500 14 000 14 000
1.16 1.21 1.21 1.20 1.20 1.18 1.18
a
Monomer ratio determined by 1H NMR spectroscopy after post-polymerization modification. bDetermined by GPC in THF as solvent for the reactive ester polymers. cCalculated from the molecular weights of the reactive ester precursor polymers as determined by GPC in THF as solvent.
Figure 1. (a) Normalized FCS autocorrelation curves for solutions of H1 (dark blue diamonds), C1 (light blue triangles), C5 (orange squares), and C10 (red circles). The solid lines represent the corresponding fits with eq 1. (b) Fluorescence intensity versus time traces from FCS measurements for solutions of p(HPMA)-co-p(LMA) copolymers with 1 (C1), 5 (C5), and 10 (C10) mol % LMA. based on the slow-motion theory and a program developed by Schneider and Freed as implemented in EasySpin.26 These simulations can account for the effect of intermediate or slow rotational diffusion of the radical on the EPR spectra. All reported values for hyperfinecoupling parameters and rotational correlation times were obtained from simulating the experimental CW spectra. The hyperfine-coupling constants are given in megahertz throughout this article. One megahertz corresponds to 0.0357 mT at a magnetic field of 336 mT.
were analyzed by applying FCS and EPR spectroscopy. Both techniques require tracer compounds that can either be covalently attached to the polymer (labeling) or interact noncovalently with the system, for example, via hydrophobic or electrostatic interactions. Labeling of the polymers was performed either with the fluorescent dye Oregon green (FCS) or with the spin label 4-amino-TEMPO ((2,2,6,6tetramethyl-piperidin-1-yl)oxyl) (EPR). The combination of RAFT-copolymerization to reactive precursor polymers with post polymerization modification was used for the labeling of polymers with either 1 mol % fluorescent dye or 5 mol % spin label starting from the same polymer precursor.5,12 For the present study, a homopolymer p(HPMA) (H1) and three random amphiphilic copolymers p(HPMA)-co-p(LMA) (C1, C5, and C10) with increasing amounts of hydrophobic LMA were synthesized. (Scheme 1, Table 1) RAFT polymerization of PFPMA yielded the reactive precursor polymer of homo p(HPMA) while copolymerization of PFPMA with LMA resulted in the precursor of random amphiphilic copolymers p(HPMA)-co-p(LMA). Because both copolymerization parameters are smaller than 1, the formation of microblocks of lauryl-methacrylate is rather unlikely.8 These completely hydrophobic and therefore well-characterizable functional precursors were transformed into amphiphilic copolymers using postpolymerization modification reactions. In the case of the fluorescently labeled polymers, pentafluorophenyl groups are first substituted with one mol % Oregon Green cadaverine and in a second step with 2-hydroxypropyl
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RESULTS AND DISCUSSION Here we first describe the synthesis of the diverse p(HPMA)co-p(LMA) copolymers, followed by FCS experiments, showing the dependence of the self-assembly behavior of the amphiphilic copolymers on the amount of hydrophobic side chains and the presence of guest molecules. Furthermore, EPR results elucidate the internal constitution of the p(HPMA)-cop(LMA) micelles and aggregates as well as their interaction with liposomes as model membranes. Finally, it is shown how FCS and EPR complement each other to yield a sophisticated picture of the conformations and interactions of p(HPMA)-cop(LMA) Polymer Synthesis and Characterization. To investigate the influence of the content of hydrophobic LMA on the selfassembly and aggregation behavior of amphiphilic p(HPMA)co-p(LMA) copolymers, we synthesized polymers composed of only hydrophilic p(HPMA) and copolymers containing 1, 5, and 10 mol % LMA. The size and nature of the self-assembled structures in aqueous solution of the copolymers alone and after loading with the hydrophobic model drug domperidone 4068
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the formation of aggregates composed of several micelles. (See Figure 2.) The detection of two distinct species shows that the aggregates coexist together with single chains, most likely in a dynamic equilibrium.
amine (H1; C1−C10). For EPR spectroscopy, 5 mol % of the reactive ester groups was substituted by 4-amino-TEMPO (C1e−C10e) and the remaining pentafluorophenyl groups were substituted by 2-hydroxypropyl amine. Different welldefined polymers with low polydispersities (PDI: 1.16 to 1.21) and similar molecular weights (Mn) were obtained independent of the amount of hydrophobic groups and functionalization. (Scheme 1, Table 1) Fluorescence Correlation Spectroscopy: Relation between Hydrophilic/Hydrophobic Composition and Aggregation. FCS is a powerful tool for the investigation of the dynamic properties of fluorescent molecules, macromolecules, or nanoparticles in various environments.23 The method is based on monitoring fluctuations of the fluorescence intensity originating from species diffusing through a very small observation volume, typically the focus of a confocal microscope. A correlation analysis of these fluctuations yields information on the diffusion coefficients and hydrodynamic radii of the species. (See the Materials and Methods for details.) Because of the high sensitivity and selectivity of the method, very small species (down to 1 nm) at nanomolar concentrations can be studied. In a first step, the size and self-assembly of H1 and C1−C10 in aqueous solution were analyzed by FCS. Typical fluorescence intensity versus time traces and the corresponding autocorrelation curves are shown in Figure 1a,b. The obtained hydrodynamic radii are listed in Table 2.
Figure 2. Schematic sketch of different polymer arrangements in aqueous solutions.
The copolymers C1 and C5 are intermediate systems to the above-described extremes. The correlation function obtained for the random amphiphilic copolymer with one mol % LMA (C1) is still in good agreement with the diffusion of a single species. The hydrodynamic radius of C1 with Rh = 4.0 nm is slightly larger than that of H1. Overall, C1 has too few hydrophobic side chains to form intramolecular micelles or aggregates that were not detected in either the fluorescence intensity versus time traces or in corresponding autocorrelation curves (Figure1a,b). The results show that an only slightly hydrophobized copolymer such as C1 is still dissolved as swollen polymer chains. The interpretation of the FCS data obtained for the random amphiphilic copolymer p(HPMA)-co-p(LMA) with 5 mol % LMA (C5) is more complex. The fluorescence intensity versus time traces show some peaks of high count rate that can originate from bigger aggregates of C5 (Figure 1). However, because there are very few such aggregates, their contribution to the autocorrelation curve is almost negligible and it can be represented by eq 1 using a single-component fit. The fit yields a hydrodynamic radius for C5 of Rh = 3.7 nm, which is slightly smaller compared with the radius of C1. This observation might be surprising with respect to the observed larger aggregates in Figure 1b. One possible explanation can be found considering that a key characteristic of polysoaps is the formation of intramolecular micelles above a critical alkyl group content (CAC).13 Copolymers with alkyl contents exceeding the CAC are no longer dissolved as random coils of swollen polymer chains but form intramolecular micelles within one polymer chain with a collapsed hydrophobic core, primarily composed of clustered alkyl groups, and a shell of swollen hydrophilic loops anchored to the core. (See Figure 2.) Such polymers remain in solution in a more globular state, as it is, for example, known for polymers in a poor solvent.27 This coil-toglobule-like transition is accompanied by a shift to low viscosities of polymer solutions and a reduction of hydrodynamic radii.13 Consequently, one possible explanation for the unexpected small Rh of C5 could be that at 5 mol % of hydrophobic LMA the intrachain CAC of p(HPMA)-cop(LMA) copolymers of this molecular weight is reached. Thus, intramolecular micelles are formed, which reduces the detected radius. At the same time, aggregation of some micelles
Table 2. Hydrodynamic Radii As Determined by FCS polymer H1 C1 C5 C10 C1+Dom C5+Dom C10+Dom
Rh1 (nm)a
Rh2 (nm)a
± ± ± ± ± ± ±
43.5 ± 2.5
3.4 4.0 3.7 3.5 3.9 5.7 3.5
0.2 0.25 0.23 0.21 0.24 0.34 0.21
62.5 ± 3.5
a
Determined via FCS in PBS (pH 7.4) with c(polymer) = 0.1 mg/mL and c(domperidone) = 0.05 mg/mL.
The correlation function for the homopolymer p(HPMA) (H1) can be well-represented by eq 1, assuming only one type of diffusing species with Rh = 3.4 nm. This shows that the completely hydrophilic polymer is dissolved as single swollen polymer chains in a random coil conformation. On the contrary, an LMA content of 10 mol % (C10) clearly results in interchain aggregation (Figure 1a,b). The fluorescence intensity versus time trace (Figure 1b) shows many peaks of high count rate that originate from the appearance of large aggregates. This is further reflected by the fit to the correlation function of C10 that requires two components: one giving a hydrodynamic radius of Rh = 3.5 nm for single polymer chains and a second component corresponding to Rh = 43.5 nm for the polymer aggregates. The higher number of hydrophobic side chains clearly provokes interchain aggregation toward bigger self-assembled objects, which have been described as a “string of connected intrachain micelles” in previous publications concerning polysoaps.15 Thus, for C10, the content of hydrophilic HPMA is not large enough to stabilize unimeric, intramolecular micelles. The random nature of the copolymers leads to conformations where hydrophobic LMA can be close to the surface of the core−shell micelles, provoking 4069
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toward bigger aggregates takes place, as observed in Figure 1b. Because the number of aggregates is very small compared with the intrachain micelles, FCS detects only the Rh of the latter. The fluorescence intensity versus time traces (Figure 1b) obtained from FCS for C1, C5, and C10 reveal that the onset of hydrophobic clustering seems to happen around 5 mol % of LMA for p(HPMA)-co-p(LMA) copolymers. This first evidence found via FCS measurements could be verified by subsequent EPR experiments, which will be discussed below. Compared with CAC values published for other polysoaps with lauryl side chains, this range is smaller, for example, for poly(4vinyl-N-ethylpyridinium) bromide and quaternized poly(ethylenimine) polymers, a modification with 10 mol % lauryl side chains is necessary to obtain typical polysoap behavior.13,20,28 However, it is important to consider that in these two cases both copolymers feature ionic hydrophilic parts in contrast with the neutrally charged p(HPMA)-co-p(LMA) copolymers studied here. Because repulsive forces between the positive charges on the polymer backbone partially loosen the inner hydrophobic clusters, it is likely for p(HPMA)-cop(LMA) to exhibit smaller CAC values compared with charged polymers. Despite the charge, HPMA is less hydrophilic compared with the above-mentioned polymers. Both effects can explain that for p(HPMA)-co-p(LMA)copolymers hydrophobic self-assembly is already observed for copolymers carrying only 5 mol % LMA. Fluorescence Correlation Spectroscopy: Effect of Encapsulating a Hydrophobic Model Drug. In the next step, we analyzed the influence of encapsulating a hydrophobic model drug, domperidone, into the copolymers C1, C5, and C10 on their aggregation behavior. When amphiphilic copolymers exhibit hydrophobic nano- or microdomains they are capable of solubilizing hydrophobic, low-molecular-weight substances. The hydrodynamic radii are listed in Table 2, and the respective autocorrelation curves for the copolymers and the copolymers loaded with 50 wt % domperidone are shown in Figure 3. The attempt to encapsulate domperidone into C1 was barely successful; consequently, no change of the hydrodynamic radius was detected. Therefore, the hydrodynamic radius remained almost constant at Rh = 3.9 nm. This result is in good agreement with the observation that C1 is dissolved as swollen polymer chains (see above) because the amount of LMA is below the CAC. Consequently, no hydrophobic microdomains are formed to solubilize the model drug. Upon loading with domperidone, the hydrodynamic radius of C5 increases from Rh = 3.7 to 5.7 nm. This significant increase in Rh can be explained by the before-mentioned picture of C5 self-assembling into intrachain micelles: hydrophobic molecules can be solubilized by these unimeric, hydrophobic microdomains, leading to an effective size increase due to encapsulation of (one or more) drug molecules. These findings further support our assumption that the CAC of p(HPMA)-cop(LMA) copolymers is ∼5 mol % of LMA. Encapsulation of domperidone into C10 provokes a strong increase in the detected hydrodynamic radius of the aggregates from Rh = 43.5 to 62.5 nm, and the small component representing the single polymer chains is almost undetectable anymore. The interchain aggregates exhibit a huge loading capacity, and the aggregates are stabilized by the hydrophobic interactions with domperidone. CW EPR Results and Discussion. Continuous-wave (CW) EPR is an intrinsically local magnetic resonance technique.
Figure 3. Normalized FCS autocorrelation curves for solutions of copolymers in the absence of domperidone (triangles, squares, circles) and of copolymers loaded with 50 wt % domperidone (diamonds). Solid lines represent the respective fits to eq 1.
Hence it is sensitive to changes in the direct environment of a paramagnetic center (here an unpaired electron in a spin-probe molecule) in a p(HPMA)-co-p(LMA) solution. In general, there are two ways to add electron spins to an otherwise diamagnetic system: spin probing, that is, the addition of a stable, spin-carrying molecule that interacts noncovalently with the system, or spin labeling, that is, covalent attachment of a stable, spin-carrying label to a specific site in the system, as shown in Scheme 1. The advantage of the spin-probing strategy is that there is no chemical modification of the tested system. However, one cannot be sure a priori about the localization of the probes inside a sample. In contrast, spin labeling changes a system chemically, but the localization of the labels is well known.29,30 Here we performed both types of experiments. For spin-probing, we employed the fatty acid derivative 16-DSA (16-DOXYL stearic acid; see Figure 4a). This probe is rather hydrophobic and therefore is predominantly located in hydrophobic domains when exposed to aqueous solutions of amphiphilic copolymers. For spin labeling, we attached 5 mol % of 4-amino-TEMPO ((2,2,6,6-tetramethyl-piperidin-1-yl)oxyl) to p(HPMA)-co-p(LMA). The probing approach was chosen for the elucidation of the composition of the self-assembled aggregates. Spin labeling was used to study p(HPMA)-cop(LMA)−liposome interactions because this approach allows for the exclusive observation of the polymers in an otherwise quite complex system. CW EPR: Influence of Hydrophobic Attraction on the p(HPMA)-co-p(LMA) Aggregate Composition. In Figure 4, the CW EPR spectra of 16-DSA in solution with C1, C5, and C10 are shown for two cases: (i) spin probing with 16-DSA on p(HPMA)-co-p(LMA) copolymers loaded with domperidone (Figure 4d) and (ii) spin probing with 16-DSA on p(HPMA)co-p(LMA) copolymers in the absence of domperidone (Figure 4e). In most cases, one can observe two spectral components, a fast rotating one, A, as indicated by three sharp lines, and a slowly rotating one, B, as indicated by broad lines (see Figure 4a−c; Figure 4c: schematic sketch of simulated CW EPR 4070
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Figure 4. (a) Molecular structure of 16-DSA spin probes. (b) Schematic depiction of the origin of the two DSA species A (in hydrophilic environment) and B (in hydrophobic environment) through partial incorporation of DSA into LMA-rich domains. (c) CW EPR spectra corresponding to 16-DSA in hydrophilic environment (yellow, denoted “A” in panel b), in hydrophobic environment (blue, denoted “B” in panel b), and the spectrum stemming from superposition of A and B (black), as observed for 16-DSA in p(HPMA)-co-p(LMA) solutions due to partitioning of the probes. (d) CW EPR spectra of 0.2 mM 16-DSA in solution with 3 mg/mL p(HPMA)-co-p(LMA) (1% LMA (C1): red; 5% LMA (C5): yellow; 10% LMA (C10): blue) loaded with 1.5 mg/mL domperidone. The separation of the dashed lines correlates with the strength of hyperfine interaction in the DSA probes. As a consequence of a decreasing hydration of the polymer chains and accordingly of Azz with increasing LMA content, the strength of hyperfine interaction decreases from top to bottom. (e) CW EPR spectra of 1 mM 16-DSA in solution with 3 mg/mL p(HPMA)-co-p(LMA) (1% LMA: red; 5% LMA: yellow; 10% LMA: blue) without domperidone. The fraction of incorporated 16-DSA probes is significantly reduced compared with the spectra derived from domperidone-containing samples. Note that the two peaks marked with the asterisk stem from baseline artifacts that could not be completely removed by baseline subtraction.
component A, that is, the component stemming from fast rotating DSA probes. In the presence of domperidone, the simulations for C1 yield χA = 0.08, for C5 χA = 0.05, and for C10 χA = 0.04, yet note that the extraction of exact values for χA from the simulation of two-component nitroxide spectra with at least one component in the slow-tumbling regime can be rather difficult due to tensor framework-dependent effects of electron Zeeman (g-) and hyperfine (A-)anisotropy on the line shape.32 Because for spin probing p(HPMA)-co-p(LMA) with 16-DSA the spectral component A is always close to the fast rotation regime and component B is always in the slow tumbling regime, the above values for χA might feature a non-negligible margin of errors. (This, however, does not imply the impossibility of exact simulations, only a non-negligible error propensity of such simulations.) However, the tendency of decreasing χA with rising LMA content of the polymers is clearly supported by the spectral simulations. Therefore, the spin-probing approach with 16-DSA reveals that the mole fraction of the slow spectral species B increases with rising LMA content from C1 to C10 in the presence of domperidone (Figure 4). Moreover, the hyperfine splitting of the nitroxide EPR signal (due to coupling of the unpaired electron to the nuclear spin of the nitrogen in the NO moiety, in principle, the splitting between the EPR lines) decreases with increasing LMA content, which can be clearly observed in the presence of domperidone.33 The hyperfine splitting of DSA is sensitive to
spectra corresponding to species A and B and their superposition). One can directly observe that the fraction of the slow species (component B) increases with rising LMA content of the polymers in the presence and in the absence of domperidone. A slow spectral component B coexists with a fast one, A, only if the exchange rate of the probes between polymer-rich phases, which slow down the DSA rotation, and the aqueous phase is slow on EPR time scales, which is at Xband frequencies of 9.4 GHz between approximately 10−9 and 10−5 s.31 If exchange takes place with frequencies above 1/ (10−9 s), then one observes a spectrum averaged between species A and B. In contrast, exchange frequencies below 1/ (10−5 s) lead to superposition of pure spectral components A and B. (The condition for the slow-exchange regime in a twosite system can be expressed as: τi−1 ≪ Δω. τ1 denotes the lifetime of a species i at site i, for example, in an LMA aggregate, and Δω denotes the spectral separation between the lines of species A and B.)31 Hence, the mole fraction of B is a measure of the extent of long-lived incorporation of the spin probes into hydrophobic volume fractions. For the spin-probing approach, the fraction of the spectral component B, that is, the contribution of species B to the overall experimental spectrum, was extracted from spectral simulations. (See the Materials and Methods and the Supporting Information.) The experimental spectra, Sexp, were simulated as a weighted sum of doubleintegral normalized CW EPR spectra: Sexp = χASA + (1 − χA)SB, where χA can be regarded as the mole fraction of the spectral 4071
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the environment of the NO group.34−36 In hydrophobic or apolar regions the hyperfine splitting is smaller than in hydrophilic or polar environments. Thus, with increasing LMA content, the environment of DSA species B (in polymer-rich domains) becomes more and more hydrophobic in p(HPMA)-co-p(LMA) copolymer solutions. Also, the tendency of decreasing hyperfine splitting with increasing LMA content is supported by the spectral simulations. From the spectral simulations, characteristic values for Azz can be found, which denotes the hyperfine coupling component in the molecular z direction that is most strongly affected by changes in the polarity of the environment of a probe. (In Figure 4, this is indicated by the separation of the dashed lines of the outermost extrema in a spectrum.33) For domperidonecontaining solutions of C1, spectral simulations yield Azz = 95.5 MHz (Aiso = tr(A)/3 = 43.5 MHz), for solutions of C5 Azz = 95 MHz (Aiso = 43.3 MHz), and for solutions of C10 Azz = 93.5 MHz (Aiso = 42.8 MHz). In the absence of domperidone, the 16-DSA CW EPR spectra show only a significant contribution of species B in the case of C10. This species could be simulated with the same parameters as those used for the simulation in the presence of domperidone, with a slightly faster rotational diffusion in the absence of domperidone (Diso,dom = 0.9 × 107 s−1/Diso,withoutdom = 2.3 × 107 s−1, with Diso denoting the isotropic rotational diffusion coefficient). From the decreasing Azz values of the spectral species B (Figure 4d) with increasing LMA content in p(HPMA)-co-p(LMA) copolymer solutions containing domperidone, one can deduce that a rising LMA amount leads to the pronounced expelling of residual water from hydrophobic clusters. Reference measurements on 16-DSA in different solvents show that the observed isotropic hyperfine splitting (42.8 to 43.5 MHz) is representative of a mean hydrophobicity of mixtures of isopropanol and chloroform (Aiso,i‑propanol = 44.2 MHz/ Aiso,chloroform = 41.4 MHz). Hence, aggregates of p(HPMA)-cop(LMA) copolymers clearly provide a more hydrophobic environment than a purely or PBS-buffered aqueous solution. In the absence of domperidone, one does not observe interaction of 16-DSA with C1. No pronounced water-depleted regions in the system are found, which is in full agreement with the FCS data, giving a Rh comparable to the completely hydrophilic homo p(HPMA). For C5 and C10, a weak but significant incorporation of 16-DSA into hydrophobic regions is detected. These data support the assumption that the onset of hydrophobic interactions (CAC) toward the formation of intraand interchain aggregation of p(HPMA)-co-p(LMA) copolymers are around 5 mol % of LMA. Furthermore, the hydrophobic domperidone apparently supports the formation of hydrophobic cavities. In conclusion, with increasing amount of hydrophobic lauryl side chains, the volume fraction of the formed hydrophobic domains in p(HPMA)-co-p(LMA) solutions increases (decreasing χA). These domains are further stabilized by hydrophobic interactions with domperidone leading to pronounced formation of hydrophobic clusters. With increasing LMA content, the amount of residual water in hydrophobic clusters decreases (decreasing Azz), such that hydrophobic interactions in the clusters become stronger. This increasing strength of hydrophobic interaction with increasing LMA content correlates well with the pronounced formation of polymerrich domains with increasing LMA. These findings are in good agreement with the data gained from FCS, revealing a general trend of increasing hydrodynamic radii of C1 to C10 upon
loading with domperidone due to rising strength of the hydrophobic interactions and pronounced intermolecular aggregation. Therefore, the observation of larger hydrophobic regions found with increasing LMA content in FCS is further mirrored in the results from EPR spectroscopy. CW EPR: Interactions of Amphiphilic p(HPMA)-cop(LMA) Copolymers with Model Membranes. For a spinlabeling approach, one covalently introduces additional hydrophobic side-chains to the system, hence one can expect slightly different observations than those in the spin-probing approach. For the spin-labeled analogue to C1, C1e, we observed only one component, A, that can be attributed to the spin-labeled polymer in the presence and absence of domperidone. (Note that due to the low polarity of the spin labels, which leads to pronounced hydrophobic interaction among the chains, domperidone does not generally lead to significant changes in CW EPR spectra of labeled polymers (Azz or χA) because the drug’s interaction with the polymers is primarily of a hydrophobic nature. See the Supporting Information for the spectra of spin-labeled p(HPMA)-co-p(LMA) in the presence and absence of domperidone). Because the spin labels are polymer-bound, incorporation into hydrophobic clusters would need drastic conformational rearrangement of the polymer backbone. An incorporation of labels into only small hydrophobic patches, like observed for C1, is therefore unlikely. In contrast, for C5e and C10e, there are significant contributions of a spectral component B observable in the CW EPR spectra. However, in this case, the spectral component A does not stem from freely rotating, nitroxide-carrying molecules but from labels attached to the polymer backbone. These fast rotating labels are not incorporated in the hydrophobic center of p(HPMA)-co-p(LMA) aggregates or micelles, probably because they reside in polymer segments of low LMA density, which are more solvent-exposed segments with lower aggregation propensity. To elucidate the mechanism of membrane interaction of p(HPMA)-co-p(LMA) copolymers, we studied their interaction with liposomes, which can be regarded as model membranes. The liposomes are composed of lecithin and cholesterol in a molar ratio of 66/1. In Figure 5, the effect of binding of spinlabeled p(HPMA)-co-p(LMA) copolymers to liposomes on CW EPR spectra is shown. For C1e, one cannot observe any effect when liposomes are present in the solution. For C5e and C10e, however, one can observe a significant effect: The fraction of the spectral component B increases notably. Therefore, with increasing LMA content of the polymers the general interaction strength with liposomes increases. There are no significant additional differences in the CW EPR spectra of the spin-labeled polymers if the lecithin-cholesterol ratio is changed. (See the Supporting Information.) Note that we did not observe any significant changes in the CW spectra of spinlabeled p(HPMA)-co-p(LMA) when the domperidone concentration was varied. (See the Supporting Information; spectra in Figure 5 are all shown for presence of domperidone only.) The domperidone independence of the spectra is a consequence of the strong hydrophobic interactions induced among the copolymer chains by the spin labels. This is evident from significant amounts of species B even in the absence of domperidone. Such additional hydrophobicity provided by domperidone does not entail significant changes in hydrophobic interaction strength anymore. Therefore, we here focus only on the interaction between spin labeled p(HPMA)-co4072
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ASSOCIATED CONTENT
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FCS and CW EPR spectroscopy are two complementary techniques with regard to their sensitivity and the length scale of a measurement. Whereas FCS gives rise to information about the ensemble-average size on the length scale of a few to tens of nanometers, CW EPR spectroscopy yields data about the rotational dynamics and the local environment of a paramagnetic center on length scales of only a few nanometers. Combining these two methods, it is possible to obtain a unique picture of p(HPMA)-co-p(LMA) copolymers in aqueous solutions, ranging from the size of micelles and aggregates to the composition of the interior of these self-assembled systems. The picture of p(HPMA)-co-p(LMA) that arises from the combined study is the following: Above LMA contents of ∼5%, p(HPMA)-co-p(LMA) aggregates significantly, whereas at lower LMA contents, hydrophobic clusters may be rather small and only of transient nature. However, the presence of domperidone fosters the hydrophobic aggregation of p(HPMA)-co-p(LMA). EPR spectroscopy complements this picture: A generally observed trend is that with increasing LMA amount the tendency and strength of hydrophobic interactions increase among the copolymers alone (as can be observed by decreasing hyperfine-splitting of the DSA probes with increasing LMA content) or with hydrophobic guest molecules such as domperidone or model membranes (as can be deduced from the spectral contribution of species B). These observations shed light on the molecular mechanisms underlying the transport properties of p(HPMA)-co-p(LMA) copolymers across cell membranes and biological barriers as, for example, the BBB. Initially domperidone is hydrophobically bound inside the hydrophobic patches of the copolymers. Upon contact with (cell) membranes, p(HPMA)-co-p(LMA) copolymers are attracted toward the membranes because of hydrophobic interactions, as described for the liposomes above. Consequently, the hydrophobic patches of LMA may then no longer be available to bind domperidone, which may subsequently be transferred or diffuse to the cell membrane to minimize solvent exposure. Although all of our experimental results in this study and previous studies substantiate such a scenario, further in vitro experiments are needed to obtain a more detailed picture. In general, we find that the intra- and intermolecular aggregation propensity increases with the mole fraction of hydrophobic groups in the system. As such, it is possible to control the formation of intramolecular micelles or composite aggregates by the amount of LMA comonomers or the presence of hydrophobic cosolutes as domperidone.
Figure 5. CW EPR spectra of spin-labeled (5%) p(HPMA)-cop(LMA) with different fractions of LMA (1% LMA (C1e): red; 5% LMA (C5e): yellow; 10% LMA (C10e): blue) in solution with lecithin-cholesterol (1/66)-based liposomes (solid lines). The dashed lines depict the spectra before interaction with the liposomes. The strength of interaction, as indicated by the difference between spectra before and after binding of p(HPMA)-co-p(LMA) to the liposomes, increases with increasing LMA content of the copolymer.
p(LMA) and liposomes and neglect the effect of domperidone on this interaction. For C1e, no changes in the presence of the liposomes were observed. In contrast, the fraction of spectral component B and hence the mole fraction of the slowly rotating spin label increases when liposomes are added to a solution of C5e or C10e. This in return means that the copolymers interact with the model membranes and such hinder or at least significantly slow down the rotational dynamics of the spin-label side chains. This may be explained by physical interactions with the liposomes; that is, the copolymers adsorb onto the surface of the liposomes and the hydrophobic lauryl side chains as well as the amphiphilic spin labels insert into the hydrophobic alkyl chain part of the lipid bilayer. The large hydrophobic liposomes thereby lead to an increasing contribution of slowly rotating spin labels. These interactions between liposomes and hydrophobic lauryl chains must be due to hydrophobic attraction because the fraction of the spectral component B in the case of interaction with the liposomes becomes increasingly prominent as the LMA content of the polymers increases. These findings are in good agreement with a recent in vitro study testing the copolymers C1, C5, and C10 as carriers to enhance the delivery of rhodamine 123 (Rh123) over a BBB model.11 Rh123 is a substrate of the efflux transporter Pgp and cannot pass the BBB. The study showed no enhanced transport of Rh123 with C1 but increasing transport rates for C5 and even more pronounced for C10. Our results here give a molecular scale explanation of these former observations: C1 exhibits no membrane interaction, whereas C5 and C10 take the first step of adsorbing to the membranes, which can finally lead to inhibition of Pgp and enhanced transport of Rh123 to the brain side.
S Supporting Information *
Spectral simulation of C1−C10 with domperidone and C10 alone of CW EPR spectra with 16DSA, spectral parameters of species B of 16-DSA, CW EPR spectra of spin-labeled polymers C1e−C10e with and without domperidone, and CW EPR spectra for different amounts of LMA in HPMA-co-LMA and different types of liposomes. This material is available free of charge via the Internet at http://pubs.acs.org. 4073
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(26) Schneider, D. J.; Freed, J. H. Biological Magnetic Resonance Vol 8. Theory and Applications; Plenum Press: New York, 1889. (27) Rubinstein, M.; Colby, R. Polymer Physics; Oxford University Press: New York, 2004. (28) Straus, U. P.; Gershfeld, N. N.; Crook, E. V. J. Phys. Chem. 1956, 60, 577−584. (29) Ottaviani, M. F.; Favuzza, B.; Sacchi, B.; Turro, N. J.; Jockusch, S.; Tomalia, D. A. Langmuir 2002, 18, 2347−2357. (30) Pace, M. D.; Snow, A. W. Macromolecules 1995, 28, 5300−5305. (31) Atherton, N. M. Electron Spin Resonance; Ellis Horwood: Chichester, U.K., 1973. (32) Weil, J. A.; Bolton, J. R.; Wertz, J. E. Electron Paramagnetic Resonance; John Wiley & Sons: New York, 1994. (33) Likhtenshtein, G. I.; Yamauchi, J.; Nakatsuji, S.; Smirnov, A.I.; ; Tamura, R. Nitroxides; Wiley-VCH: Weinheim, Germany, 2008. (34) Saracino, G. A. A.; Tedeschi, A.; D’Errico, G.; Improta, R.; Franco, L.; Ruzzi, M.; Corvaia, C.; Barone, V. J. Phys. Chem. A 2002, 105, 10700−10706. (35) Kurzbach, D.; Kattnig, D. R.; Zhang, B.; Schlüter, A. D.; Hinderberger, D. Chem. Sci. 2012, 3, 2550−2558. (36) Akdogan, Y.; Heller, J.; Zimmermann, H.; Hinderberger, D. Phys. Chem. Chem. Phys. 2010, 12, 7874−7882.
AUTHOR INFORMATION
Corresponding Author
*Fax: +49-6131-3924778. Tel: +49-6131-3920361. E-mail:
[email protected] (R.Z.),
[email protected] (D.H.). Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was financially supported by the Deutsche Forschungsgemeinschaft (DFG) under grant number HI 1094/2-1 and the Max Planck Graduate Center with the University of Mainz (MPGC). We thank Christian Bauer for technical support and Prof. Hans W. Spiess for continuing support. D.K. acknowledges support by the Gutenberg Academy of the University of Mainz.
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