J. Phys. Chem. C 2008, 112, 1427-1434
1427
Fluorescence Boost in Polyelectrolyte Multilayer Architectures Stephan Meyer,† Paula Pescador,‡ and Edwin Donath*,† Institute of Medical Physics and Biophysics, Leipzig UniVersity, Ha¨rtelstrasse 16-18, D-04107, Leipzig, Germany, and Bioengineering and Bioelectrochemistry Group, Departament d’Enginyeria Quı´mica, UniVersitat RoVira i Virgili, AVda. Paı¨sos Catalans 26, 43007 Tarragona, Spain ReceiVed: August 15, 2007; In Final Form: October 23, 2007
Polyelectrolyte multilayer coated colloids were fabricated by means of layer-by-layer (LbL) deposition of poly(allylamine hydrochloride) (PAH) and poly(styrene sulfonate) (PSS) in alternating order. The top layer consisted of PAH fluorescently labeled with one of the following dyes: succinimidyl 6-(N-(7-nitrobenz-2oxa-1,3-diazol-4yl)amino) hexanoate, fluorescein isothiocyanate, or rhodamine B isothiocyanate. For all three PAH-dye conjugates, a manifold increase in fluorescence intensity was observed upon adsorption of a PSS layer on top of the labeled PAH layer. The degree of fluorescence increase varied for each of the fluorescent dyes and depended on the polyelectrolyte concentration, revealing the existence of different modes of adsorption of the incoming polyelectrolyte molecules. Dequenching as well as changes in the local environment following polyelectrolyte adsorption were identified as the possible causes for the fluorescence enhancement. These findings prove that the top layer has a distinct physicochemical quality that makes it intrinsically responsive toward polyelectrolyte interactions, opening the way for the design of new sensing devices.
Introduction Fluorescently labeled colloids constitute a versatile tool widely used in life sciences, medicine, and nanotechnology. The use of fluorescent dyes as marker molecules for colloidal particles allows their qualitative and quantitative analysis by techniques such as flow cytometry,1,2 confocal laser scanning microscopy,3 or fluorescence spectroscopy.4 An important application of these fluorescently labeled particles are cytometric bead based immunological assays.5 Fluorescent particles can be fabricated (i) by incorporating the dye molecules directly into the beads during their synthesis (internal labeling), (ii) by using dye-labeled polyelectrolyte molecules that are then adsorbed on the particles by a suitable means such as the layer-by-layer (LbL) approach,6 or (iii) by conjugation of label molecules with functional groups located at the surface of the colloids.7,8 The LbL coating of charged surfaces with oppositely charged polyelectrolytes, introduced by Decher et al. in 1991,9,10 is a powerful means to engineer interfaces. A charged surface is exposed in alternating order to solutions of polycations or polyanions. After each deposition step the surface charge is reversed since more than the stoichiometric equivalent of charges is adsorbed, which allows for the deposition of a new layer of oppositely charged molecules.11 Excess polyelectrolyte is removed by centrifugation or filtration following each deposition step.12,13 In this way a polyelectrolyte multilayer film can be grown. Later the LbL approach was also extended to colloidal particles.14,15 A variety of materials such as polystyrene latex beads, silica particles, cross-linked melamine formaldehyde colloids, organic crystals, and living cells have been used as templates for * To whom correspondence should be addressed. E-mail: edwin.
[email protected]. Fax: +49 (0)341 9715749. Tel: +049 (0)341 9715704. † Leipzig University. ‡ Universitat Rovira i Virgili.
multilayer assembly.15-20 The adsorbed species differ widely ranging from small ions21 to biological particles,22 but mostly polyelectrolytes such as poly(ethyleneimine) (PEI), poly(allylamine hydrochloride) (PAH), poly(diallyldimethylammonium chloride) (PDADMAC), and poly(styrene sulfonate) (PSS) are employed.14 The advantages of the LbL technique are on one hand its striking simplicity and on the other hand its extreme versatility. With simple equipment and easy preparation steps one can readily tailor thin polymer films with different functional molecules (e.g., proteins, carbohydrates, nucleic acids, antibodies, and lipids) by using them as adsorbing species.23-30 The resulting structures can be controlled at the nanoscale level.31 The procedure is applicable to any substrate configuration, allowing the design of devices based on functional layer components as well as the study of phenomena on the nanoscale. Despite the versatility and widespread use of polyelectrolyte multilayers in research and practical applications, not all details about film formation and resulting properties are understood. For example, a remarkable fluorescence increase occurring after adsorption of another polyelectrolyte layer onto a fluorescently labeled top layer was recently observed by Caruso and coworkers;32 however, to this date the phenomenon has been neither explored nor explained. The present work is an attempt to gain some insight on the nature and characteristics of this fluorescence increase effect. Our motivation stemmed both from a fundamental interest in the causes for this behavior and also from its potential practical applications. Understanding the physicochemical mechanisms underlying this effect might reveal valuable information about the formation and properties of polyelectrolyte multilayer films. In particular, the observed behavior suggests that the outermost layer has a distinct physicochemical quality as compared with the inner layers. This fact has obvious relevance since the top layer is the one exposed to contact and interaction with the environment. The practical uses of the fluorescence increase effect could include the
10.1021/jp076551u CCC: $40.75 © 2008 American Chemical Society Published on Web 01/16/2008
1428 J. Phys. Chem. C, Vol. 112, No. 5, 2008 development of a sensor system for low concentrations of polyelectrolyte. Therefore we set out to systematically study this effect aiming to reveal its causes and explore its potential use for sensing purposes. Experimental Methods Materials. Silica particles of 3.03 ( 0.17 µm diameter were purchased from microParticles GmbH (Berlin, Germany). PAH (MW 70 000), PSS (MW 70 000), N,N-dimethylformamide (DMF), dextran sulfate sodium salt (DXS; MW 500 000), fluorescein isothiocyanate (FITC), rhodamine B isothiocyanate (RITC), as well as Sephadex G-25 chromatography gel were supplied by Sigma (Deisenhofen, Germany). Succinimidyl 6-(N(7-nitrobenz-2-oxa-1,3-diazol-4yl)amino) hexanoate (S-NBD) was obtained from Molecular Probes (Eugene, Oregon). Spectra/ Por molecular porous membrane tubing (MWCO: 12 00014 000 Da) was from Spectrum Laboraties, Inc. (California). All other chemicals were obtained from Fluka (Neu-Ulm, Germany). Labeling of PAH. PAH was labeled with FITC according to the standard method used for protein labeling.33 RITC-labeled PAH was prepared as described by Schna¨ckel et al.6 The conjugation of S-NBD to PAH was carried out as follows: PAH was dissolved in 0.1 M carbonate buffer, pH 8.5, and S-NBD, dissolved in carbonate buffer containing 10% DMF (v/v), was added dropwise. After an incubation time of 48 h, the NBDlabeled PAH was separated from the free dye by gel chromatography, and afterward the remaining buffer was removed by dialysis. The labeled and purified polyelectrolytes were lyophilized and stored at 4 °C until further use. The degree of labeling defined as the ratio of dye molecules attached to one polyelectrolyte molecule was determined as follows: lyophilized labeled PAH was dissolved in 0.5 M NaCl, and the absorbance of the solution was compared to that of a calibration series of the respective dyes. Measurements were performed in a UV/vis spectrometer Cary 50 (Varian Instruments) using the molar extinction coefficients 484 ) 41 200 M-1 cm-1 for NBD, 490 ) 80 900 M-1 cm-1 for FITC, and 559 ) 62 100 M-1 cm-1 for RITC. Coating of Silica Particles. The bulk solution of particles (500 µL, 5 wt %, approximate surface area 0.025 m2) was centrifuged, and the supernatant was discarded. The particles were then redispersed in 200 µL of 0.5 M NaCl. This suspension was pipetted into a 1 mg/mL solution of PAH in 0.5 M NaCl and incubated under constant stirring for 10 min. Next, the colloids were centrifuged at 650 g for 2 min and washed with 0.1 M NaCl. The washing procedure was repeated 3 times to remove excess polyelectrolyte. Then a 1 mg/mL solution of PSS in 0.5 M NaCl was added to form the next polyelectrolyte layer. After PSS adsorption, washing was performed in distilled water. The centrifugation/washing steps and the sequential adsorption of oppositely charged polyelectrolytes were repeated to obtain silica particles with 8 polyelectrolyte layers ([PAH/PSS]4). Next, a layer of fluorescence labeled PAH was adsorbed. Finally, one further PSS or DXS layer was deposited. Characterization by Flow Cytometry. The fluorescence intensity, forward scattering (FSC) distributions, and sideward scattering (SSC) distributions of the coated silica particles were measured with a flow cytometer (FACS Calibur, Becton Dickinson, U.S.A.) equipped with a 15-mW, 488-nm, air-cooled argon ion laser. For each sample, 10 000 events were recorded, except in the case of the time-dependent quenching assay measurements. Time-dependent assays were conducted over 1-3 min with a
Meyer et al. time resolution of 1 s. Recording of the fluorescence started 3 s after addition of sodium dithionite. Sodium dithionite is known to be an irreversible quenching agent for NBD. For each measurement the solution of dithionite was freshly prepared (1 M) in 0.1 M Tris (pH 10.0). 20 µL of this solution were added to 1 mL of the silica suspension containing about 250 000 silica particles, which corresponds to a molar ratio between sodium dithionite and NBD of 1.6 × 108. The geometric mean values of the fluorescence intensity distribution within every second were calculated. The experimental error was calculated as follows: The particle solutions were diluted to a concentration of approximately 50 counts per second. The geometric mean of 50 consecutive counts was calculated. The highest and lowest calculated values of these geometric means were taken as the error limits of the method. The single particles were gated on the basis of their light scattering to separate them from the aggregates, i.e., doublets, triplets, and quadruplets, since there is a linear increase of forward scattering with the size of the aggregates.6 The data were analyzed with the WinMDI 2.8 software written by Joseph Trotter. Fluorescence Spectroscopy. Fluorescence spectroscopy measurements were performed using a Fluoromax 2 fluorescence spectrometer (Spec Industries., Inc., U.S.A.). The excitation wavelength was set at 467 nm for NBD, 492 nm for FITC, and 530 nm for RITC. Excitation and emission slits were adjusted at 3 nm. ζ-Potential Measurements. ζ potentials were measured in water using a Malvern Zetasizer 4. Each Measurement was repeated 5 times, and afterward the mean value and the standard deviation were calculated. Results and Discussion The first question we addressed is whether the fluorescence increase effect is specific for the particular combination of polyelectrolytes used by Caruso et al., namely, a PAH layer deposited onto a fluorescent PSS-rhodamine layer,32 or it constitutes a more general effect occurring also for other combinations of polyelectrolytes and dyes. To evaluate the dependence of the fluorescence increase on the nature of the fluorophore, a series of experiments with different PAH-labeled conjugates was conducted. RITC-PAH, NBD-PAH, or FITCPAH were adsorbed on a [PAH/PSS]4 support. Next, one more PSS layer was deposited on top of the fluorescent PAH layer. Figure 1 shows the respective fluorescence intensity distributions. Three fluorescence distributions are superimposed for each dye, corresponding in increasing order of fluorescence intensity to: (i) unlabeled particles with 8 polyelectrolyte layers, (ii) particles carrying fluorescently labeled PAH as the top layer, and (iii) particles with an additional PSS layer on top of the fluorescent layer. The geometric means of the three fluorescence intensity distributions (arbitrary units, a.u.) were 1.4, 8, and 79 for RITC; 2.5, 145, and 487 for NBD; and 1.5, 480, and 1330 for FITC. These results show that there is a remarkable fluorescence increase for all three dyes upon subsequent PSS adsorption. However, the degree of the fluorescence increase is specific for each particular fluorophore. To illustrate this aspect, the ratio of fluorescence intensity after and before PSS adsorption was calculated for each dye and taken as the basis for further considerations. The calculated ratios were 9.9 for RITC, 3.4 for NBD, and 2.8 for FITC, respectively. The next step in our study was to find out if the fluorescence increase is associated to the specific properties of the PAH/ PSS couple or if other polyelectrolyte species can also induce
Polyelectrolyte Multilayer Architectures
Figure 1. Histograms of flow cytometry measurements of particles coated with eight polyelectrolyte layers ([PAH/PSS]4, leftmost peaks), after adsorption of fluorescently labeled PAH ([PAH/PSS]4/labeled PAH, middle peaks), and after deposition of a further PSS layer on top of the labeled PAH layer ([PAH/PSS]4/labeled-PAH/PSS, rightmost peaks). The degree of labeling for RITC-PAH was 0.8 molecules per 100 monomers, for FITC-PAH 0.47 molecules per 100 monomers, and for NBD-PAH 0.16 molecules per 100 monomers.
Figure 2. Geometric mean of fluorescence intensity of particles coated with [PAH/PSS]4/NBD-PAH (Fl) and with one more adsorbed layer consisting of PSS (Fl+PSS) or DXS (Fl+DXS). The degree of labeling for NBD-PAH was 0.16 molecules per 100 monomers.
this effect. This question might have special relevance for practical applications. For example, when measuring fluorophore-conjugated biomarkers assembled into a multilayer film, a possible change of fluorescence intensity after interaction with other molecules should be taken into account. Particles were coated with a multilayer of [PAH/PSS]4/NBDPAH. Next, an outermost layer of either DXS or PSS was adsorbed on top of the NBD-PAH layer, and the fluorescence intensity of both sets of particles was compared. Figure 2 shows that after deposition of either a PSS layer or a DXS layer the fluorescence intensity increases. However the magnitude of the increase is different. The fluorescence intensity after adsorption of PSS increased from 145 to 487 a.u., whereas in the case of DXS, the increase was from 145 to 277 a.u.. Our results up to here demonstrate that the fluorescence increase effect occurs for the three different dyes RITC, NBD, and FITC covalently bound to PAH and that PSS, DXS, and PAH32 are, upon deposition, able to induce this effect. We also found a fluorescence increase when FITC-labeled enzymes where covered with a PSS layer (data not shown). Thus, this phenomenon seems to be rather general, although the degree
J. Phys. Chem. C, Vol. 112, No. 5, 2008 1429 of fluorescence increase depends on the fluorophore and the nature of the incoming polyelectrolyte layer. At this point, however, it would be preliminary to speculate about the causes for the observed quantitative differences, since the results obtained with different dyes refer to different degrees of labeling. Nevertheless, two possible reasons for the observed effect can be considered. On one hand, all three dyes used in this work are susceptible to self-quenching caused by label-label interaction, resulting in a smaller fluorescence intensity than the fluorophore concentration would suggest. For example, in the case of RITC, self-quenching is the basis of the widely used octadecylrhodamine dequenching assay as described by Bardelletti et al.34 Upon adsorption of further polyelectrolyte molecules, a dequenching process might take place leading to an increase of the observed fluorescence intensity. The dequenching itself could be caused by desorption of a fraction of previously adsorbed molecules upon interaction with the incoming polyelectrolyte. This desorption would lead to an overall reduced concentration of dye molecules per unit area, which in turn could result in dequenching and a subsequent increase of fluorescence. This mechanism may be described as “dequenching by desorption”. Alternatively, taking into account that self-quenching is a result of dye-dye interactions, e.g., two fluorophore molecules forming a dimer, the adsorbing polyelectrolyte species could facilitate dimer-monomer transitions by interfering with the dye-dye interaction. On the other hand the fluorescence yield of dyes is strongly dependent on the local environment. For example, fluorescein shows a pH-sensitive behavior: with increasing pH the thermodynamical equilibrium between the nonemitting and fluorescent forms is shifted toward the fluorescent state, which leads to a higher fluorescence yield.35 Moreover, the fluorescence yield often depends on the nature of the solvent. In this sense, the observed effect could be related to changes in the local environment caused by polyelectrolyte adsorption. Experiments were carried out in order to check these hypotheses. First the possibility of dequenching as a result of desorption was investigated. An evidence for a desorption process would be a fluorescent supernatant after adsorption of a further polyelectrolyte layer. Silica particles (surface area approximately 0.049 m2) were coated with a multilayer of [PAH/ PSS]4/NBD-PAH. The mean coverage for the PAH/PSS couple is about 1 mg m-2 per single layer.12 It is obvious that, in order to observe a dequenching effect, a significant part of the labeled top PAH layer should desorb during adsorption of PSS. Assuming desorption of 50% of the NBD-PAH upon PSS adsorption, 24.5 µg of NBD-PAH would remain in the supernatant after coating the particles. Accordingly, a control solution of 24.5 µg of NBD-PAH was prepared in 1.2 mL of 0.5 M NaCl and measured by fluorescence spectroscopy. Similarly, the supernatant after adsorption of the final PSS layer on the particles was collected and measured. A second control solution containing the same amount of PSS used for adsorption was also measured. The recorded spectra are presented in Figure 3. The spectrum of NBD-PAH can be clearly identified, with an emission maximum at 550 nm. Because all the samples are in a very low concentration range (although still above the lower limit of the spectrophotometer sensitivity), the Raman peak of the solvent can be seen superimposed on the NBD-PAH spectrum. Therefore the spectrum of the solvent was subtracted from the spectrum of the fluorophore leading to the typical emission spectrum of NBD, with a maximum at a 538 nm.
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Figure 3. Fluorescence spectra, from top to bottom: 24.5 µg of NBDPAH in 1.2 mL 0.5 M NaCl, the same sample after subtraction of the spectrum of the solvent, 1 mg/mL PSS in 1.2 mL 0.5 M NaCl, supernatant after PSS adsorption on colloids, solvent (0.5 M NaCl). The degree of labeling for NBD-PAH was 0.16 molecules NBD per 100 monomers PAH.
Further analysis shows that the spectrum of the supernatant is quite similar to that of PSS, again with the Raman peak of the solvent clearly visible. However there is no evidence of the presence of NBD-PAH in the supernatant after deposition of PSS. At most, only negligible amounts of fluorophore below the sensitivity of the photometer can be present in the supernatant, since it does not show a typical NBD spectrum. In view of these results, the hypothesis of dequenching by desorption can be ruled out as an explanation for the fluorescence increase upon subsequent polyelectrolyte adsorption onto a fluorescently labeled PAH layer. Within an adsorbed fluorescent PAH layer there may exist sites where the dye molecules are closely arranged or even directly interact with each other, resulting in self-quenching. This may be a consequence of intramolecular interactions of conjugated dye molecules within the PAH.36 The PAH molecule is highly flexible, with free rotation allowed by the σ-bonds between its carbon atoms.37 This property provides the condition for intramolecular and intermolecular dye-dye interactions even when the label density is comparatively low. Thus, the distribution of the fluorophores in dye-PAH conjugates may be characterized by reduced separating distances between them. Self-quenching may be caused either by Fo¨rster transfer or by π-π stacking interactions of two or more dye molecules within direct contact. The adsorption of PSS interacting with the amino groups of PAH might compete with the interaction of neighboring or aggregated fluorophores. Upon adsorption of PSS, PAH adopts a largely two-dimensional configuration leading on average to enhanced distances between the fluorophores. This would attenuate self-quenching and thus lead to an increase of the observed fluorescence. To find out whether dequenching is actually involved in the mechanisms of fluorescence increase upon layer adsorption, dye-PAH conjugates with different degrees of labeling were prepared. Particles carrying RITC-PAH as the top layer were fabricated, and their fluorescence was measured before and after adsorption of a further PSS layer. Figure 4 shows the initial fluorescence intensities as well as the fluorescence increase after adsorption of PSS (amplification factor) as a function of the degree of labeling of RITC-PAH. The higher the degree of labeling, the lower is the fluorescence intensity of the particles, proving the existence of selfquenching. However, the observed amplification factor follows
Meyer et al.
Figure 4. Amplification factor (ratio of fluorescence intensity after and before PSS adsorption, solid line) of mean fluorescence as a function of degree of labeling. The dashed line shows the corresponding fluorescence intensity before PSS adsorption as a function of the degree of labeling.
the opposite trend: with a higher degree of labeling the effect of fluorescence increase upon layer adsorption becomes more pronounced, until a saturation level is reached. There is thus a direct correlation between the degree of self-quenching and the fluorescence increase after the adsorption of another layer of PSS. These results support the hypothesis that dequenching is most likely involved and plays an important role in the effect of fluorescence increase upon subsequent layer adsorption, at least in the case of RITC. To identify the specific mechanism of dequenching, further investigations are required. At this point we consider the fact of self-quenching from a different angle, looking now for a direct proof of the existence of fluorophore molecules in a quenched state and of the influence of PSS on their occurrence. We focused on RITC-PAH, since Figure 4 showed a relation between self-quenching and fluorescence increase for this particular dye species. If a dequenching interaction between the fluorophore and PSS takes place, the absorption spectra of RITC-PAH in the presence and absence of PSS should differ. UV/visible spectroscopy is not straightforward in the case of colloidal suspensions because of the high increase of the background caused by light scattering. Therefore the spectrum of a solution of labeled PAH was compared to that after addition of equimolar amounts of PSS, which are supposed to form a polyelectrolyte complex with the PAH molecules.38 Both spectra are shown in Figure 5. The spectrum of RITC-PAH shows two peaks, one at 530 nm and a second one at 560 nm. After addition of PSS the first peak transforms into a shoulder at 530 nm, and the peak height at 560 nm increases from 0.12 to 0.21 units. Both spectra result from the superposition of two peaks, reflecting the existence of two absorption maxima and consequently two different states of the RITC molecules. The superimposed peaks were resolved into single peaks by Lorentz fitting. The ratio between the two extracted peaks (560 nm/530 nm) is 1:1 for RITC-PAH and 2.5:1 after addition of PSS. A correlation of the absorption maximum at 530 nm with RITC dimers and of the maximum at 560 nm with RITC monomers has been reported in the literature.39-42 It is also known that the fluorescence yield of the dimers is lower than that of the monomers. In the case of RITC-PAH, the presence of PSS leads to a shift of the distribution between monomers and dimers toward the monomers. This provides further support for the hypothesis
Polyelectrolyte Multilayer Architectures
Figure 5. UV/visible spectroscopy of RITC-PAH (black) and RITCPAH mixed with PSS (Mol% 1/1; gray) in aqueous solution of 0.5 M NaCl. The superimposed spectra were converted to simple peaks by Lorentz fitting (dashed lines). The label ratio of RITC-PAH was 0.8 molecules RITC per 100 monomers PAH.
Figure 6. Spectrophotometric measurements of fluorescence intensity of FITC, RITC, and NBD in different solvents. The ratio between the measured intensities in the three solvents and the intensity in water is shown.
that the adsorbing polyelectrolyte interferes with dye-dye interactions, inducing dequenching and leading finally to the observed fluorescence increase, at least in the case of the RITCPAH/PSS couple. When considering the possibility of developing a sensing tool, it would be of practical use to know whether dequenching is the only cause for fluorescence increase upon layer adsorption or if there are also other factors involved in the mechanism. In this sense, changes in the local environment after adsorption of PSS onto a labeled PAH layer might have an impact on the fluorescence increase. When trying to elucidate this question, one must keep in mind that the dye molecules incorporated into the top layer have a different location with respect to the bulk solution as compared to the same dye molecules after adsorption of one more polyelectrolyte layer. In the second situation the influence of the solvent might be reduced. To explore this point experimentally, the fluorescence of equal concentrations of nonconjugated dyes was measured in solvents of different polarity, namely, water with a dielectric constant () of 78, methanol ( ) 32), ethanol ( ) 24), and propanol ( ) 20). Figure 6 shows the fluorescence intensity of RITC, FITC, and NBD solutions in these four solvents. It is immediately apparent that the lowest fluorescence intensity corresponds always to the samples dissolved in water, while the fluorescence intensity increases for all dyes in the other three solvents. The
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Figure 7. Time-dependent geometric mean of the fluorescence intensity of [PAH/PSS]4/NBD-PAH (black line) and [PAH/PSS]4/NBD-PAH/ PSS (gray line) before and after addition of sodium dithionite. The perpendicular dashed line marks the moment of addition of sodium dithionite. The 3-second gap in the collected data is a consequence of the measuring conditions (see Experimental Methods).
factor of increase is specific and differs from dye to dye. In the case of NBD there is even a reciprocal dependency between dielectric permittivity of the solvent and fluorescence intensity of the sample. The solubility of each dye in each solvent was not investigated, a factor that could also have an influence on the fluorescence intensity in the different solvents. It is conceivable that for some solvents a particular micelle configuration may give rise to self-quenching effects, resulting in lower fluorescence intensity. Nevertheless it is clear that the changes in the local environment after adsorption of one further polyelectrolyte layer onto a PAH-labeled layer must be taken into account as a possible factor for fluorescence increase. Next we set out to determine to which extent the dye molecules are accessible for bulk water, and whether this accessibility changes after adsorption of another polyelectrolyte layer. Thus the effect of a water-soluble, polar, and hydrophilic quenching agent on the fluorescence of the multilayer films was examined. Sodium dithionite is known to be a quenching agent for NBD. Its mechanism of action is through chemical reduction of the 7-aminobenz into a 7-nitrobenz group, which leads to a complete and irreversible loss of fluorescence. The quenching assays performed for colloids with an outermost NBD-PAH layer and for particles with a further PSS layer on top are shown in Figure 7. It can be seen that the fluorescence of the sample with NBDPAH as the top layer is quenched in less than 3 s after addition of sodium dithionite. However, with one more PSS layer on top a much slower quenching effect is observed. An instantaneous loss of about 12% is followed by a slow decay over a time period of 3 min to 50% of the initial value of fluorescence intensity. This indicates that the inner fluorescent label is located at sites that have a reduced accessibility from the aqueous phase. These results again stress the fact that the local environment around the fluorophore is determined by the multilayer structure, in particular when an additional polyelectrolyte layer covers the labeled layer. Comparison of the two quenching assays reveals that the adsorption of PSS on top of the NBD-PAH layer induces a dramatic change of the label environment. In sum, the quenching assay points to the conclusion that the polarity around dye molecules in the top layer is different from that in inner layers. The shift toward a less polar environment during layer adsorption leads in turn to the creation of a less aqueous
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Figure 8. Amplification factor of fluorescence intensity (9) and ζ-potential (b) after adsorption of a PSS layer onto [PAH/PSS]4/RITCPAH as a function of the concentration of PSS in the solution. The incubation time was 10 min. The degree of labeling of RITC-PAH was 0.8 labels per 100 monomers.
Figure 9. Mean fluorescent intensity as a function of incubation time using three different PSS concentrations: 9, 1 mg/mL; b, 0.002 mg/ mL; 2, 0.0005 mg/mL.
environment around the fluorophore molecules. This could be an important factor for fluorescence increase, as suggested by the results shown in Figure 6. These findings may have relevant implications for practical applications, for example, the protection of a giving substance from water-soluble polar agents by incorporation within such multilayer films. In summary, the fluorescence increase upon subsequent layer adsorption can be the result of the concurrence of dequenching phenomena and changes in the local environment. The dequenching is not caused by desorption of fluorescent molecules during adsorption of the next layer but by decreased dye-dye interaction induced by the newly adsorbed polyelectrolyte. The deposition of polyelectrolyte molecules leads to a strong change of the label environment, especially with regard to its accessibility for water. The transfer into a more nonaqueous environment has also a relevant impact on the observed increase of fluorescence. The existence of this effect could be used to design a sensing device for detection of polyelectrolyte species. To explore this possibility, the concentration of the adsorbing polyelectrolyte species was varied, and the fluorescence increase of RITCPAH upon adsorption of PSS was monitored. The results are displayed in Figure 8, showing a sigmoidal curve for a half-log plot of the amplification factor of fluorescence increase and the concentration of PSS. A linear behavior over only one magnitude between 0.5 and 5 µg PSS/mL is observed. This small linear range is in concordance with a ratio of excess PSS from 1 to 10 in terms of the amount of PSS required to coat the total surface area of the sample. The change in the fluorescence behavior was correlated to a change of the ζ-potential from 0 mV at low PSS concentrations to -45 mV at higher PSS concentrations. While this system may have a limited use for concentration measurements due to its small dynamic range, it may find application as a sensing tool for the presence of polyelectrolytes above a threshold value. To establish the possible influence of the adsorption time on the final values of fluorescence intensity, [PAH/PSS]4/NBDPAH-coated colloids were incubated with different concentrations of PSS within and above the linear range and the fluorescence intensity of the particles was measured after defined time intervals. An increase of the fluorescence can be observed only within the first minute of incubation. There are no systematic changes in fluorescence intensity for longer incubation times, as illustrated in Figure 9. This finding is in contrast
to the overly simple interpretation of the small increase of fluorescence at low polyelectrolyte concentrations as being the result of insufficient adsorption within the 10 min exposure to the bulk polyelectrolyte. The adsorption time required to fully cover the particle surface with another polyelectrolyte layer can be estimated as follows: the amount of polyelectrolyte needed for full coverage of one colloid is distributed at a concentration of 1 mg polyelectrolyte/ mL solution in a volume of 2.8 × 10-11 mL. This volume would correspond to a shell with a thickness of 0.67 µm surrounding each particle. Taking a diffusion coefficient of 7 × 10-11 m2/s for PSS (MW 47 300)43 and recalculating this coefficient for PSS (MW 70 000), by inserting this value in the Einstein diffusion equation, one can calculate, assuming that diffusive transport is the rate-limiting step, that adsorption of a single PSS layer should take about 15 ms. The calculated adsorption times for the other concentrations would be 6.4 s for a concentration of 0.002 mg/mL and 17.5 s for 0.0005 mg/mL, respectively. On one hand these estimations can explain that there is no fluorescence increase above incubation times of 1 min, since the complete adsorption process would be finished after 17.5 s in the case of the lowest concentration or even faster for higher polyelectrolyte concentrations in the coating solution. On the other hand, these considerations are not able to explain the dependence of the fluorescence increase on the concentration of the coating solution. If full coating of the particles was the only condition involved, the same increase in fluorescence intensity would be expected for all the samples, since all should reach full coating within the time scale of the measurements. Other causes have to be involved. It could be that differences in the structure of the assembled layer as a result of concentrations of polyelectrolyte in the coating solution were responsible for the variation in the fluorescence behavior. A distance of 30 nm between the PSS molecules was calculated for a polyelectrolyte solution of 1 mg/mL, which ranges within the contour length of one PSS molecule (115 nm).37 In this situation the PSS molecules are forming a “PSS continuum“ rather than appearing as single entities. When adsorption of these molecules takes place, the structure of the resulting layer could be different than that of a layer assembled from a less concentrated solution of 0.002 mg/mL, with a calculated intermolecular distance of 240 nm. In this case the PSS molecules adsorb individually, separated both in time and space. To get a better understanding of the adsorption process at low and high PSS concentrations the following experiment was conducted. Particles with labeled PAH on top were incubated
Polyelectrolyte Multilayer Architectures
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Figure 10. Geometric mean of fluorescence intensities of flow cytometric measurements of particles: BG, background without any fluorescently labeled layers; Control, with fluorescence labeled PAH layer on top; low, with labeled PAH incubated in low PSS concentration (0.002 mg/mL); high, with labeled PAH incubated in high PSS concentration (1 mg/mL); low-high, with labeled PAH incubated first in low and afterward in high PSS concentration; high-low, with labeled PAH incubated first in high and afterward in low PSS concentration. The fluorescent layer was either RITC-PAH (white columns) or NBDPAH (hatched columns) with a degree of labeling of 0.8 and 0.16 labels per 100 monomers, respectively.
Figure 11. Geometric mean of fluorescence intensities of flow cytometric measurements of particles: BG, background without any fluorescence labeled PAH layers; CONTROL, a single fluorescently labeled PAH layer as the top layer; HIGH, a PSS layer adsorbed onto fluorescently labeled PAH from high PSS concentration (1 mg/mL). 0.1...1.7 cm2, increasing particle number at a constant, but low PSS concentration (0.002 mg/mL), expressed as total particle surface area. The degree of labeling was 0.8 labels per 100 monomers. Above the bars the fluorescence intensity histograms of particles is shown.
in low-concentration PSS solutions. Next, the colloids were washed 3 times and incubated again in PSS, but this time with a high concentration. The same procedure was also carried out in reversed order of PSS concentrations, first high and then low. The experiments were performed for two different dye-PAH conjugates, RITC-PAH and NBD-PAH. The results are shown in Figure 10. The fluorescence intensity increases after adsorption of the PSS layer onto the dye-labeled PAH layer, although at low PSS concentration the increase is smaller than at high PSS concentration. After a second incubation in PSS, the fluorescence intensity remains constant when switching from high to low concentration. By reversing the order of PSS incubation, first at low and afterward at high concentration, the fluorescence intensity increases after the second incubation but does not reach the same level observed for an initial incubation in high concentration. The quotient of both values is 73% for RITC-PAH and 78% for NBD-PAH, respectively. These data support the fact that PSS adsorption is irreversible, at least in the time scale of the experiment. Re-exposing the particles to a low PSS concentration after PSS was adsorbed from a 1 mg/mL solution did not lead to desorption. What might be the difference between adsorption from a low or a high PSS concentration? As calculated above, at low PSS concentration the PSS molecules adsorb individually separated in time and space over a time interval of the order of 101 s. This is a rather long time in terms of the molecular dynamics of these polymers, sufficient to adopt, upon interaction with PAH, an optimal conformation from the energetic and entropic points of view. It seems favorable that at low concentrations of adsorbing solution an intercalating arrangement between the adsorbed molecules and the existing PAH layer is adopted, since there is no or at least less interference of adsorbing PSS in the neighborhood. This assumption is in good agreement with the observed decrease of the ζ-potential, indicating a trend toward a neutralization of the surface rather than the overcharging characteristic for high PSS concentrations. Thus it is more likely that the PAH molecules remain in a conformation compromising between PSS interaction with label interaction. It seems also probable that in this case the structure on the top of the film resembles that of a layer of PSS/PAH complexes rather than a system of two more or less fuzzy, but still distinct, PAH and
PSS layers. This would explain the ongoing adsorption of PSS when the particles are re-exposed to the polyelectrolyte but this time in higher concentration and the corresponding fluorescence increase observed for these samples. In reversed order, the initial incubation with high PSS concentration would not allow major rearrangements of PAH/PSS complexes once the PAH layer is completely covered by PSS. Thus a further incubation at low concentration of PSS would have no or at least less effect on the fluorescence intensity. The different quality of the top layer assembled at low PSS concentrations results further in an increased accessibility of NBD for hydrophilic species, as it was inferred from the increasing initial drop of fluorescence upon exposure to dithionite. Adsorption with subsequent structural changes of the adsorbing species at interfaces is not uncommon. It has been demonstrated for proteins leading to a non-Langmuir adsorption behavior.44,45 So far, only the PSS concentration has been taken into account to explain these observations. However at very low PSS concentrations the total surface area of the particles becomes increasingly important. If the number of particles is too large, there simply might not be enough PSS available for the formation of a complete layer on all of the particles. Therefore, another experiment was performed where the amount of particles with RITC-PAH as top layer was varied, but the PSS concentration was kept constant at 0.002 mg/mL. The sample volume was such that at a concentration of 0.002 mg/mL the amount of PSS was sufficient to coat a total surface area of 2 cm2. This ensures an excess of PSS of 20:1 for a total surface area of 0.1 cm2 and a ratio of 1.18:1 for the highest area of 1.7 cm2. The results of the experiment are shown in Figure 11. A sample with particles incubated in 1 mg/mL PSS is also shown for comparison. On top of each column the histogram of fluorescence intensity as recorded by FACS is plotted. As expected, the fluorescence increase of the sample incubated in 1 mg/mL PSS (total surface area 0.4 cm2) reaches the degree typical for high PSS concentration. At 0.002 mg/mL PSS, the fluorescence intensity increase of samples with a total surface area between 0.1 and 0.4 cm2 is smaller but constant within the error of the measurements. However at higher surface areas the fluorescence intensity increase is even smaller, indicating that
1434 J. Phys. Chem. C, Vol. 112, No. 5, 2008 under these conditions the amount of adsorbed PSS was not sufficient to cover the surface. This is also evidenced by the fluorescence intensity distribution of the particles, which shows an increasing proportion of aggregates as seen from the pronounced structured right tail of the histograms. The degree of aggregation is 20 and 45% for the total surface area of 0.9 and 1.7 cm2, respectively. For all the other samples the fraction of aggregates is about 10%. The increasing aggregation proves the heterogeneity of the particle surface in terms of surface charge density, which is characteristic of an incomplete coverage. For ratios of excess PSS to surface area of 2.2:1 and 1.18:1, the limited amount of PSS interferes with the process of layer formation. In summary, these results suggest that the observed fluorescence increase effect can easily find applications in the development of new sensing tools. A bead-based assay or a device incorporating fluorescence assays on a chip could be used to detect threshold concentrations of polyelectrolyte species. An important finding is also the evidence that layers formed at low polyelectrolyte concentrations may be substantially different from layers formed at regular concentrations. This fact should be taken into account when building multilayers from species available only in small quantities, as is the case for many biologically relevant polyelectrolytes. Conclusions It was shown that fluorophore-fluorophore interactions and very likely changes in the local environment after deposition of one more polyelectrolyte layer onto a PAH-labeled layer are responsible for the fluorescence increase upon adsorption. When using fluorophores at interfaces for investigations of multilayer films or for quantification of incorporated biofunctionalizing molecules such as enzymes, the change of fluorescence intensity in the presence of other polyelectrolytes has to be kept in mind. The top layer has different physicochemical properties than the inner regions of the film in this kind of multilayer-coated colloids, at least in the areas immediately surrounding the dye molecules. Furthermore, the mode of adsorption of the next incoming layer depends on the polyelectrolyte concentration of the coating solution. On the basis of these new insights into the nature and mechanism of formation of multilayer films, the feasibility of a novel sensing tool for polyelectrolytes was demonstrated. Acknowledgment. P.P. thanks Universitat Rovira i Virgili for a PhD studentship and also acknowledges financial support from DURSI-Generalitat de Catalunya (Grant No. 2005BE 00512). References and Notes (1) Lawrie, G. A.; Battersby, B. J.; Trau, M. AdV. Funct. Mater. 2003, 13, 887. (2) Bele, M.; Siiman, O.; Matijevic´, E. J. Colloid Interface Sci. 2002, 254, 274. (3) Verhaegh, N. A. M.; Van Blaaderen, A. Langmuir 1994, 10, 1427. (4) Deng, G.; Markowitz, M. A.; Kust, P. R.; Gaber, B. P. Mater. Sci. Eng. C 2000, 11, 165.
Meyer et al. (5) Morgan, E.; Varro, R.; Sepulveda, H.; Ember, J. A.; Apgar, J.; Wilson, J.; Lowe, L.; Chen, R.; Shivraj, L.; Agadir, A.; Campos, R.; Ernst, D.; Gaur, A. Clin. Immunol. 2004, 110, 252. (6) Schna¨ckel, A.; Hiller, S.; Reibetanz, U.; Donath, E. Soft Matter 2007, 3, 200. (7) Tedeschi, C.; Caruso, F.; Mo¨hwald, H.; Kirstein, S. J. Am. Chem. Soc. 2000, 122, 5841. (8) Caruso, F.; Lichtenfeld, H.; Donath, E.; Mo¨hwald, H. Macromolecules 1999, 32, 2317. (9) Decher, G.; Hong, J. D. Makromol. Chem. Makromol. Symp. 1991, 46, 321. (10) Decher, G. Science 1997, 277, 1232. (11) Hoogeveen, N. G.; Cohen Stuart, M. A.; Fleer, G. J.; Bo¨hmer, M. R. Langmuir 1996, 12, 3675. (12) Bertrand, P.; Jonas, A.; Laschewsky, A.; Legras, R. Macromol. Rapid Commun. 2000, 21, 319. (13) Voigt, A.; Lichtenfeld, H.; Sukhorukov, G. B.; Zastrow, H.; Donath, E.; Ba¨umler, H.; Mo¨hwald, H. Ind. Eng. Chem. Res. 1999, 38, 4037. (14) Sukhorukov, G. B.; Donath, E.; Lichtenfeld, H.; Knippel, E.; Knippel, M.; Budde, A.; Mo¨hwald, H. Colloids Surf. A 1998, 137, 253. (15) Sukhorukov, G. B.; Donath, E.; Davis, S.; Lichtenfeld, H.; Caruso, F.; Popov, V. I. Polym. AdV. Technol. 1998, 9, 759. (16) Donath, E.; Sukhorukov, G. B.; Caruso, F.; Davis, S. A.; Mo¨hwald, H. Angew. Chem., Int. Ed. 1998, 37, 2202. (17) Neu, B.; Voigt, A.; Mitlo¨hner, R.; Leporatti, S.; Gao, C. Y.; Donath, E.; Kiesewetter, H.; Mo¨hwald, H.; Meiselmann, H. J.; Ba¨umler, H. J. Microencapsulation 2001, 18, 285. (18) Moya, S.; Da¨hne, L.; Voigt, A.; Leporatti, S.; Donath, E.; Mo¨hwald, H. Colloids Surf. A 2001, 183, 27. (19) Gao, C.; Moya, S.; Lichtenfeld, H.; Casoli, A.; Fiedler, H.; Donath, E.; Mo¨hwald, H. Macromol. Mater. Eng. 2001, 286, 355. (20) Donath, E.; Moya, S.; Neu, B.; Sukhorukov, G. B.; Georgieva, R.; Voigt, A.; Ba¨umler, H.; Kiesewetter, H.; Mo¨hwald, H. Chem.-Eur. J. 2002, 8, 5481. (21) Dai, Z.; Voigt, A.; Leporatti, S.; Donath, E.; Mo¨hwald, H. AdV. Mater. 2001, 13, 1339. (22) Lvov, Y.; Haas, H.; Decher, G.; Mo¨hwald, H.; Mikhailov, A.; Mtchedlishvily, B.; Morgunova, E.; Vainshtein, B. Langmuir 1994, 10, 4232. (23) Caruso, F.; Niikura, K.; Furlong, D. N.; Okahata, Y. Langmuir 1997, 13, 3427. (24) Lvov, Y.; Ariga, K.; Ichinose, I.; Kunitake, T. J. Am. Chem. Soc. 1995, 117, 6117. (25) Decher, G.; Lehr, B.; Lowack, K.; Lvov, Y.; Schmitt, J. Biosens. Bioelectron. 1994, 9, 667. (26) Sukhorukov, G. B.; Montrel, M. M.; Petrov, A. I.; Shabarchina, L. I.; Sukhorukov, B. I. Biosens. Bioelectron. 1996, 11, 913. (27) Hiller, S.; Leporatti, S.; Schna¨ckel, A.; Typlt, E.; Donath, E. Biomacromolecules 2004, 5, 1580. (28) Szyk, L.; Schwinte´, P.; Voegel, J. C.; Schaaf, P.; Tinland, B. J. Phys. Chem. B 2002, 106, 6049. (29) Caruso, F.; Schu¨ler, C. Langmuir 2000, 16, 9595. (30) Caruso, F.; Mo¨hwald, H. J. Am. Chem. Soc. 1999, 121, 6039. (31) Caruso, F.; Niikura, K.; Furlong, D. N.; Okahata, Y. Langmuir 1997, 13, 3422. (32) Johnston, A. P. R.; Zelikin, A.; Lee, L.; Caruso, F. Anal. Chem. 2006, 78, 5913. (33) Nargessi, R. D.; Smith, D. S. Methods Enzymol. 1986, 122, 67. (34) Bardeletti, G.; Gautheron, D. C. Arch. Virol. 1976, 52, 19. (35) Beyer, H.; Walter, W. Lehrbuch der organischen Chemie, 19th Ed.; S. Hirzel Verlag Stuttgart: Mainz, 1981; p 573. (36) Caruso, F.; Donath, E.; Mo¨hwald, H.; Georgieva, R. Macromolecules 1998, 31, 7365. (37) Donath, E.; Walther, D.; Shilov, V. N.; Knippel, E.; Budde, A.; Lowack, K.; Helm, C. A.; Mo¨hwald, H. Langmuir 1997, 13, 5294. (38) Petrak, K. In Polyelectrolytes: Science and Technology; Hara, M., Ed.; Marcel Dekker: New York, 1992; p 265. (39) Levshin, L. V.; Bocharov, V. G. Opt. Spectrosc. 1961, 10, 330. (40) Rohatgi, K. K.; Singhal, G. S. J. Phys. Chem. 1996, 70, 1695. (41) Selwyn, J. E.; Steinfeld, J. I. J. Phys. Chem. 1972, 76, 762. (42) Plant, A. L. Photochem. Photobiol. 1986, 44, 453. (43) Matsuoka, H.; Ogara, Y.; Yamaoka, H. J. Chem. Phys. 1998, 109, 6125. (44) Seigel, R. R.; Harder, P.; Dahint, R.; Grunze, M.; Josse, F.; Mrksich, M. Anal. Chem. 1997, 69, 3321. (45) Dahint, R.; Ros Seigel, R.; Harder, P.; Grunze, M.; Josse, F. Sens. Actuators, B 1997, 36, 497.
