Study of Layer-by-Layer Films on Thermoresponsive Nanogels Using

Nov 16, 2009 - Fax: +49.(0)241.8092327. E-mail: [email protected]. Tel. ... While a few studies have reported on the layer-by-layer (LbL) assembl...
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J. Phys. Chem. B 2009, 113, 15907–15913

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Study of Layer-by-Layer Films on Thermoresponsive Nanogels Using Temperature-Controlled Dual-Focus Fluorescence Correlation Spectroscopy John E. Wong,* Claus B. Mu¨ller, Ana M. Dı´ez-Pascual, and Walter Richtering* Institute of Physical Chemistry, RWTH Aachen UniVersity, Landoltweg 2, 52056 Aachen, Germany ReceiVed: April 29, 2009; ReVised Manuscript ReceiVed: October 19, 2009

While a few studies have reported on the layer-by-layer (LbL) assembly of polyelectrolytes on soft and porous templates, none have really demonstrated direct proof that the layers are actually on the template. Thermoresponsive nanogels present challenges that render a quantitative proof of successful polyelectrolyte deposition extremely difficult. Additionally, the fate of the polyelectrolyte has never been investigated during the phase transition of the coated nanogel. Here, the auto- and cross-correlation functions of a labeled polyelectrolyte assembled via the LbL technique onto soft and porous thermoresponsive labeled nanogels using dual-focus fluorescence correlation spectroscopy (2f-FCS) are presented. Performing 2f-FCS as a function of temperature, hydrodynamic radii of nanogels coated with various numbers of layers are determined, which are found to be in excellent agreement with values obtained from dynamic light scattering. This study presents irrefutable quantitative evidence of successful LbL assembly on thermoresponsive nanogels and demonstrates that the layers are not stripped off during the phase transition of the nanogels. Fo¨rster Resonance Energy Transfer (FRET) detection also supports our findings. Introduction Layer-by-Layer (LbL) assembly of polyelectrolyte multilayers has been demonstrated on various templates, from hard and planar substrates1 to rigid particles2,3 as well as hard and porous templates.4,5 Recently, we have taken the LbL technique to new dimensions by coating soft and porous templates.6-8 Indeed, we qualitatively demonstrated the feasibility of the LbL assembly of labeled polyelectrolyte multilayers on thermoresponsive poly(N-isopropylacrylamide) (PNiPAM) microgels in the submicrometer size range.8 One of the most significant challenges with substrates of such dimensions resides in the fact that they are also deformable and penetrable to the adsorbed polyelectrolyte. For such reasons, charge reversal alone is not sine qua non of successful polyelectrolyte assembly since the adsorbing layer could be argued to be stripping off the underlying layer onto which it is being deposited. The same argument applies for the LbL assembly of polyelectrolyte multilayers on rigid particles. However, from the very first time that buildup of layers was successfully demonstrated by an increase in the film thickness using single particle light scattering (SPLS)9 and, as a complementary technique, by charge reversal using zeta potential measurements, the latter technique has been widely accepted as the common tool to monitor successful LbL deposition of polyelectrolytes on rigid particles. Thermosensitive microgels are soft, porous, solvent-penetrable, cross-linked polymeric networks which are characterized by the so-called volume phase transition temperature (VPTT), at which temperature the microgel undergoes a conformational transition from a swollen to a collapsed state during heating.10 Because of all these reasons, SPLS is not suited for microgels, and even the notion of zeta potentials which is a more common notion to quantify surface charge on hard and rigid particles needs a new theoretical model.11 Due to the submicrometer size * Corresponding authors. Tel.: +49.(0)241.8098624. Fax: +49.(0)241. 8092327. E-mail: [email protected]. Tel.: +49.(0)241.8094760. Fax: +49.(0)241.8092327. E-mail: [email protected].

range of microgels, often termed as nanogels, confocal laser scanning microscopy (CLSM) is of no help for visualization of the polyelectrolyte layers on the particles. Additionally, we previously successfully demonstrated and established the systematic increase and decrease (also termed as the “odd-even” effect)7,12-14 in the hydrodynamic radius, Rh, of the coated microgel depending on the nature of the last adsorbed polyelectrolyte and which complicates the matter for a direct measurement of the layer thickness on such templates. One needs a characterizing tool that would be sensitive enough to changes in the overall size of the nanogel by capitalizing on the infinitesimal change in the size upon layer deposition. A fundamental parameter describing the diffusion of a species in solutions is its diffusion coefficient which is correlated to its Rh by the Stokes-Einstein relation.15 Any perceptible conformational change is associated with a change, no matter how feeble, in the Rh and is sometimes very difficult to detect by methods such as dynamic light scattering (DLS), pulsed-field gradient NMR, or size-exclusion electrophoresis. Fluorescence correlation spectroscopy (FCS)16-18 is a technique originally introduced in the early seventies to measure, among others, the diffusion coefficient of fluorescently labeled species at nanomolar concentrations by exploiting the detection of the fluctuation in the fluorescent intensity as the species enter and leave the detection volumesthe so-called autocorrelation function, ACF. We recently reported8 the use of single focus FCS to provide qualitative evidence of successful LbL deposition on submicrometer size microgels with a system composed of poly(Nisopropylacrylamide) (PNiPAM) and the use of two differently labeled polyelectrolytes: fluorescein-labeled poly(diallymethylammonium chloride), PDADMACFITC, and rhodamine-labeled poly(L-lysine), PLLRh. It was possible to discriminate between specific association of the fluorescent signals with the microgel due to the labeled polyelectrolyte and the free (unbound) labeled polyelectrolyte. A single diffusion of the ensemble composed of the assembled polyelectrolytes on the microgel was demon-

10.1021/jp903941c CCC: $40.75  2009 American Chemical Society Published on Web 11/16/2009

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strated by combining time-correlated single photon counting19 (TCSPC) and pulsed interleaved excitation20,21 (PIE) of two wavelengths. The approach relies on the two-color cross-correlation between two differently labeled polyelectrolytes, PLLRh and PDADMACFITC, assembled at a definite location within the multilayer on the microgel. Two-color cross-correlation unambiguously proved that the polyelectrolytes sit atop each other, but one could still argue that the latter were not anchored to the microgel and just coincidentally have the same diffusion coefficient as the microgel. The present work addresses the need for evidence of successful LbL assembly of polyelectrolyte multilayers on soft and thermoresponsive nanogels. To definitely remove any doubt about whether the polyelectrolytes are anchored to the nanogel or not, we make use this time of a charged nanogel which is chemically tagged with a fluorescent species and achieve LbL buildup using a different fluorescently labeled polyelectrolyte. Should we then detect both fluorescences with a common diffusion coefficient, it would be evidence of both species moving as one single entity. Furthermore, we want to investigate the fate of the polyelectrolytes during the VPTT of the nanogel, an issue which to our knowledge has never been explored simply due to technical and experimental limitations. Here, the nanogel used is poly(N-isopropylacrylamide-co-acrylic acid-co-rhodamine), denoted hereafter as NGRh, and the labeled polyelectrolyte is FITC-labeled poly(L-lysine) (PLLFITC). Unlabeled Poly(Llysine), PLL, and polystyrene sulfonate, PSS, are also employed during the LbL assembly to increase the distance between the two fluorescent species. We aim at a quantification of the particle size using 2f-FCS at temperatures below and above the VPTT. Experimental Section Materials. Poly(sodium 4-styrenesulfonate) (PSS), MW ) 70 000 g/mol, poly(L-lysine) bromide salt (PLL), MW ) 32 600 g/mol, fluorescein isothiocyanate (FITC)-labeled poly(L-lysine) (PLLFITC) (MW ) 32 600 g/mol), ultrapure tris(hydroxymethylaminomethane) (TRIS), and 2-(N-morpholino)-ethanesulfonic acid (MES) were purchased from Sigma-Aldrich. The monomer N-isopropylacrylamide (NiPAM) was purchased from Acros organics, and acrylic acid (AA) was purchased from Lancaster. Potassium peroxodisulfate (KPS), N,N′-methylenebis(acrylamide) (BIS), and sodium chloride (NaCl) were purchased from Merck. Sodium dodecyl sulfate (SDS) and methacryloyloxyethyl-thiocarbamoyl-rhodamine B (MW ) 683.24) were purchased from Fluka and Polysciences, respectively. All chemicals from commercial origin were used without further purification. Ultrapure water obtained from double-distilled Milli-Q water (resistivity g 18 MΩcm) was used to prepare all solutions. Synthesis of P(NiPAM-co-AA-co-rhodamine) Nanogel. The nanogel (NGRh) was synthesized by dispersion polymerization. In summary, polymerization was performed in a 1 L reaction vessel equipped with a mechanical stirrer, reflux condenser, thermometer, and gas inlet. Amounts of 12.943 g of the monomer NiPAM, 0.935 g of the cross-linker N,N′-methylenebis(acrylamide) (BIS), 0.784 g of the acrylic acid AA, 0.601 g of the surfactant SDS, and 0.0125 g of the methacryloyloxyethyl-thiocarbamoyl-rhodamine B were dissolved in 0.75 L of water at 70 °C and purged with nitrogen for at least 1 h. Polymerization was initiated with 0.703 g of potassium peroxodisulfate (KPS) dissolved in 10 mL of water and carried out for 6 h under a nitrogen stream and constant stirring at 400 rpm. The dispersion was passed through glass wool to remove

Wong et al. particulate matter and further purified by dialysis against deionized and double-distilled water for ∼1 week, using the Spectra-Por 10 000 MW cutoff membrane. Then the solution was centrifuged for 45 min at 50 000 rpm and 25 °C. Between each centrifugation, the supernatant was removed and replaced by deionized water to redisperse. After four cycles of centrifugation, the solution was freeze-dried overnight for storage. LbL Assembly on Nanogels. The assembly of polyelectrolytes was done7 by slowly adding 3 mL of an aqueous dispersion of the nanogel (0.02% by wt) to 12 mL of an aqueous 1 g/L solution of the polyelectrolytes in buffer (25 mM TRIS, 10 mM MES, and 1.15 M NaCl), pH 7.4. The nanogel being negatively charged, the first layer is always a polycation (PLL or PLLFITC). The mixture was kept under constant stirring for 4-6 h, and after each deposition, the excess polyelectrolyte and buffer was removed by using three ultracentrifugation cycles at 50 000 rpm and 25 °C, followed at each step by decantation and redispersion in water by vigorous shaking during at least 4 h. This sequence was repeated until the desired number of layers and configuration were achieved. Throughout this work, the pH was not adjusted (unless specifically stated). Before any characterizations, the coated nanogels were always redispersed in water overnight then filtered through a 1.20 µm Minisart hydrophilic filter in a laminar-flow box. Dynamic Light Scattering. The Rh was determined by dynamic light scattering (DLS) performed on highly diluted aqueous solutions of the sample using an ALV goniometer equipped with an avalanche photodiode. Temperature was increased from 293 to 325 K in steps of 2 K and then decreased again to 293 K, to get a complete cycle. The samples were allowed to equilibrate for 20 min before each temperature, and three sets of recordings were measured at each temperature. Scattered light was detected at 60° with an integration time of 120 s and computed with a digital ALV 5000E autocorrelator using an ALV Software version 5.3.2. The particle size was calculated by cumulant fits. Electrophoretic Measurements. The electrophoretic mobility (µ) measurements were performed with a Malvern Zetasizer 3000HSA. For each layer, a temperature dependence study was performed (heating from 293 to 325 K and cooling down to 293 K, with a temperature increment of 2 K). Each point is the average of 10 measurements, and the sample was allowed to equilibrate for 10 min for each temperature. Although zeta potential (ζ) is a more common notion to quantify surface charge on hard and rigid particles, no attempt has been made to convert the mobility values (µ) into ζ because nanogels are soft, porous, and solvent-penetrable particles11 that do not conform to the usual hard-sphere model hypothesized in standard theories relating µ to ζ. Dual-Focus Fluorescence Correlation Spectroscopy (2fFCS). 2f-FCS experiments were performed on a modified MicroTime200 (PicoQuant GmbH, Berlin, Germany). A detailed description of our 2f-FCS setup has been described elsewhere.22-24 Briefly, we use two identical 470 nm lasers (LDH-P-C-470B) and one 532 nm laser (PicoTA 530N) which is modified to obtain two time-delayed pulses22 (both PicoQuant GmbH, Berlin, Germany). The fluorescence emission light is split from the excitation by using a major triple-band dichroic (z470/532/ 638rpc). After passing some clear-up filters (HQ505/30m for λex ) 470 nm and HQ580/70m for λex ) 532 nm, all filters purchased from AHF-Analysentechnik, Tu¨bingen, Germany), the light is split by a nonpolarizing beam splitter cube and focused onto two single photon avalanche diodes (SPAD, PDM series, Micro Photon Devices, Bolzano, Italy). The temperature

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was regulated by a custom-made temperature-controlled cell with an absolute temperature accuracy of (0.1 K within the detection volume with a precision of (0.05 K.23 For 2f-FCS, solutions of the nanogel particles were prepared using LiChrosolv water for chromatography (No. 115333, Merck KGaA, Darmstadt, Germany), and during measurements, the sample chamber was sealed to prevent solvent evaporation and convection. Results and Discussion To quantify the particle size using FCS, we upgraded our single-focus FCS to a dual-focus FCS, 2f-FCS,25-28 which provides (a) two laser foci of precisely known distance apart and (b) an overlapping detection volume which allows for the quantitative measurement of absolute values of the diffusion coefficient with extremely high accuracy and precision.29 So, additional to the ACF curves, one can also correlate the signal recorded from the first laser focus with the signal from the second laser focus and vice versa. By measuring the ACF for each detection volume separately, and the dual-focus crosscorrelation function (CCF) between both volumes, and then comparing and analyzing the delay of the CCF decay to that of the ACFs, one can calculate absolute values of the diffusion coefficients of the fluorescent species in solution. An adequate model27 ACF/CCF for a diffusion-generated correlation decay is given by

g˜(t, δ, v) )

c 4

[

 Dtπ ∫ dz ∫ dz 8Dt + w (z ) + w (z ) × κ(z1)κ(z2)

1

2

2

2

1

2

(δ - Vxt)2 + Vy2t2 (z2 - z1 - Vzt)2 exp -2 4Dt 8Dt + w2(z1) + w2(z2)

]

(1)

where δ is the lateral distance between detection volume centers; ε1 and ε2 are factors proportional to overall excitation intensity and detection efficiency in each laser; c is the concentration of fluorescent molecules or particles; and D is their diffusion coefficient. The functions κ(z) and w(z) are given by

[ ( )]

w(z) ) w0 1 +

and

λexz

πw20n

2 1/2

(2)

κ(z) ) 2

(

2

2F exp - 2 ∫0a RdFF 2 (z) R (z)

)

(

) 1 - exp -

2a2 R2(z)

)

(3)

with

[ ( )]

R(z) ) R0 1 +

λemz

πR20n

2 1/2

(4)

where λex and λem are excitation and center emission wavelengths; n is the sample refractive index; a is the confocal pinhole radius; and w0 and R0 are fit parameters. For calculating the ACF of each focus, one has to set δ to zero in eq 1 and to replace ε1,2 by either ε12 or ε22, respectively. The integration in eq 1 is performed numerically. Fitting of experimental data is done globally for both the ACFs and the CCF, where one has the free fit parameters ε1 · c1/2, ε2 · c1/2, w0, R0, and D.22,27 The fluorescently labeled nanogel, NGRh, is a stimuliresponsive 3D soft and porous polymeric network30 wherein the NiPAM lends thermosensitively to the system, the acrylic acid allows for pH tunability (hence charge and different swelling behavior), and the rhodamine brings about the fluorescent moiety. A dilute aqueous solution of NGRh has a pH ∼ 4.7, has an Rh of 272 nm in the swollen state, and collapses to 122 nm at 50 °C, with a VPTT around 38 °C (Figure 1a). From electrophoretic measurements, the -COOH group of AA in NGRh is partially deprotonated conferring a negative charge to the nanogel particles (Figure 1b). Multilayer assembly was done from polyelectrolytes in buffer (25 mM TRIS, 10 mM MES, and 0.15 M NaCl), pH 7.4. After each deposition step, the excess polyelectrolyte and buffer were removed by ultracentrifugation at 50 000 rpm at 25 °C for 30-60 min.13 The coated nanogel was redispersed in water and washed three times before deposition of the next polyelectrolyte. Figure 1a compares the Rh, as determined by DLS, of the uncoated NGRh with that of PLLFITC-terminated NGRh wherein the number of bilayers of (PLL/PSS)n between the fluorescent species is varied (n ) 0, 1, 2). The buildup of (PLL/PSS)n onto the nanogel, NGRh, confirms the “odd-even” effect of the Rh as recently reported.13,14 Whenever the nanogel is terminally coated with the polycation, the Rh of the resulting nanogel is smaller than that of the uncoated nanogel, and whenever the nanogel is terminally coated with the polyanion, the Rh is bigger. The thermoresponsive behavior of all PSS-terminated nanogels (data not shown) shows reversible heating and cooling curves, while that of all PLL-

Figure 1. (a) Monitoring of the Rh by DLS of (A) NGRh, (B) NGRh/PLLFITC, (C) NGRh/(PLL/PSS)1.0/PLLFITC, and (D) NGRh/(PLL/PSS)2.0/PLLFITC as a function of temperature. (b) The electrophoretic mobility of A, B, C, and D during the heating and cooling cycles measured at T ) 20 and 52 °C.

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Figure 2. Fluorescence intensity as a function of temperature for the system NGRh/(PLL/PSS)n/PLLFITC, with n ) 0, 1, and 2. The same samples are characterized in a UV-vis spectrometer to determine the fluorophore concentration by which the fluorescence intensity is normalized. Open symbols are the control measurements at 20 °C measured on cooling.

terminated as well as PLLFITC-terminated (Figure 1a) nanogels exhibits some hysteresis between heating and cooling curves. Electrophoretic measurements (data not shown) confirm clear charge reversal during buildup of (PLL/PSS)n, with all PLLterminated (and PLLFITC-terminated) nanogels bearing positively charged surfaces and with all PSS-terminated nanogels bearing negatively charged surfaces. For clarity, Figure 1b only shows the electrophoretic mobilities for all PLLFITC-terminated nanogels measured at temperatures below and above the VPTT. As can be seen from Figure 1b, the hysteresis in the size of all nanogels which are PLLFITC-terminated is also observed in the surface charge when the samples are subjected to heating and cooling cycles, while all PSS-terminated nanogels show no such hysteresis. PLL is known for its ability to diffuse within the entire film when used during multilayer construction,31,32 while PSS is known for its inability to do so. The hysteresis observed is indicative of PLL or PLLFITC diffusing or interpenetrating with the entire system. It is not an easy task, by any technique, to prove that PLL diffuses in the nanogel. One could think of using microgel beads or core-shell microgels in the micrometer range to visualize the diffusion of PLL in the templates. Even then, hasty direct comparisons may be erroneous as changing the system size can inherently lead to differences in the

Wong et al. architectural morphology of the templates. In any case, charge reversal alone does not prove successful buildup of layers on such nanosize templates, hence the need for new characterization techniques adapted for the quantitative monitoring of LbL assembly on soft and deformable nanogels. Since NGRh is thermoresponsive, it is important to investigate the fate of the polyelectrolyte multilayer when the ensemble is heated beyond the VPTT, an issue which to our knowledge has never been addressed before by any technique whatsoever. To confirm that the polyelectrolyte multilayers are not stripped off the nanogel during the volume phase transition, we performed 2f-FCS as a function of the temperature.23 Another special feature about our 2f-FCS setup is a custom-made temperaturecontrolled cell with an absolute temperature accuracy of (0.1 K within the detection volume. However, prior to the temperature-controlled 2f-FCS experiment, fluorescence intensities of NGRh/PLLFITC, NGRh/(PLL/PSS)1.0/PLLFITC, and NGRh/(PLL/ PSS)2.0/PLLFITC nanogels were monitored with a JASCO FP6500 spectrofluorometer coupled to a PCT-163T temperature control unit. Figure 2 shows the normalized fluorescence intensity as a function of temperature. NGRh/PLLFITC nanogels show a very slight decrease in intensity with increasing temperature (closed symbols), and on cooling (open symbols) the initial intensity is recovered. This observation rules out any photobleaching effect. However, NGRh/(PLL/PSS)1.0/PLLFITC and NGRh/(PLL/PSS)2.0/ PLLFITC nanogels show a drastic decrease in intensity for T < 35 °C, indicating that fluorescence quenching could be responsible for the decrease in intensity of the fluorescence emission due to electrostatic interaction within the polymeric network between polyelectrolytes and nanogel alike. It has been reported that the fluorescence of the fluorophore is quenched in the presence of PSS within the multilayer system.9 The fluorescence intensity drops nearly to zero (around 48 °C) even before the nanogel is fully collapsed at T ∼ 50 °C. On cooling, the intensity is significantly less than the initial one in both systems containing PSS layers, suggesting permanent conformational restructuration of the network. In our system, this decrease in fluorescence intensity complicates not only its detection, especially at T > 35 °C, but also the data analysis. For all of the mentioned reasons, for the 2f-FCS measurements, (a) T ) 40.7 °C corresponding to the near collapsed state of the coated nanogel is chosen, and depending on the high fluorescent content

Figure 3. 2f-FCS curves of the uncoated NGRh as detected in the rhodamine channel at (a) T ) 25 °C and (b) T ) 40.7 °C. Using the particle size effect (PSE) model, an Rh of 276 and 174 nm was calculated for the uncoated NGRh in the swollen and collapsed state, respectively. These values are in good agreement with the Rh values obtained by DLS, which were, respectively, 265 and 168 nm.

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Figure 4. 2f-FCS auto- and cross-correlation curves of the NGRh/PLLFITC, NGRh/(PLL/PSS)1.0/PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC as detected in the rhodamine- and FITC-channel at T ) 25 °C and T ) 40.7 °C. Experimental data points are fitted (line) using the dual particle size effect (PSE) model, and the Rh obtained is in good agreement with that obtained from DLS.

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TABLE 1: Comparison of Rh Obtained from 2f-FCS and DLS for the Various Systems Studied T ) 20 °C

T ) 40.7 °C

Rh(2f-FCS)/nm Rh(DLS)/nm Rh(2f-FCS)/nm Rh(DLS)/nm NGRh NGRh/PLLFITC NGRh/(PLL/PSS)1.0/ PLLFITC NGRh/(PLL/PSS)2.0/ PLLFITC

276 245 254

265 242 250

174 170 168

168 165 170

258

252

166

172

on the nanogel, (b) the applied excitation power is reduced by roughly 2 orders of magnitude27 (in the range of nanowatts) as compared to a single molecule experiment, where excitation intensities of up to 30-50 µW (depending on the fluorophore used) can be applied before photobleaching is observed.33 Figure 3 shows the normalized 2f-FCS curves of NGRh, as detected separately at two different foci (ACF1 and ACF2) and cross-correlated (CCF) for two different temperatures: T ) 25 °C when NGRh is fully swollen (Figure 3a) and T ) 40.7 °C (Figure 3b). It is noteworthy that the coated nanogel is fully collapsed at T g 50 °C, and at T ) 40.7 °C it is only near collapsed. In Figure 3, the experimental data (points) were fitted (line) using the particle size effect (PSE) model which takes into account the fact that the size of the labeled particles investigated is not negligibly small as compared to the size of the detection volume.24 The Rh values calculated for NGRh at T ) 25 and 40.7 °C are 276 and 174 nm, respectively, in excellent agreement with those obtained from DLS (265 and 168 nm, respectively). The insertion of a differently labeled polyelectrolyte such as PLLFITC within the multilayer assembly on NGRh enables the detection of this bound dye using the 2f-FCS, thereby distinguishing the nanogel core from the multilayer shell when collecting data for the auto- and cross-correlation in the respective channels. Figure 4 shows the normalized 2f-FCS autoand cross-correlation curves of NGRh/PLLFITC, NGRh/(PLL/ PSS)1.0/PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC nanogels for

two different temperatures: T ) 25 and 40.7 °C. Some correlation curves reveal a broad decay at long delay times, indicating the presence of some large aggregates in addition to the individual coated nanogels. The size of the coated nanogels was obtained from fitting (line) the experimental data (points) with a bimodal PSE model24 for the ACFs and CCFs obtained from the rhodamine moieties contained in the nanogel core and the FITC moieties contained in the multilayer shell. At 25 °C, the ACFs and CCFs curves for NGRh/PLLFITC, NGRh/(PLL/ PSS)1.0/PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC nanogels could be nicely fitted, and the Rh values obtained are 245, 254, and 258 nm, respectively. These results are in excellent agreement with those obtained using DLS (see Table 1), indicating clearly that the polyelectrolyte shell is bound to the nanogel core and the whole ensemble is moving as one entity. At T ) 40.7 °C, the Rh values obtained for NGRh/PLLFITC, NGRh/(PLL/PSS)1.0/ PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC nanogels are 170, 168, and 166 nm, respectively. Although there are some discrepancies between the experimental and the fit curves concerning the steepness of the decay (observed only at such high temperatures), the fits yield Rh which are in good agreement with those obtained by DLS. These results point to the fact that in the collapsed state the polyelectrolyte multilayer shell is still bound to the collapsed nanogel core. At such high temperatures, water is a poor solvent for the system. During the collapse of the nanogel core, through a combination of electrostatic interaction and hydrophobic-hydrophobic interactions as well as physical entanglement, the polyelectrolyte multilayer shell is “pulled in” with the nanogel core as the water is expelled out. The collapsed core-shell ensemble moves as one entity as evidenced by the correlation curves. In the 2f-FCS, during the heating and cooling cycles, no polyelectrolyte release is observed because ACFs and CCFs show no extra decay which could have been correlated to the free labeled polyelectrolyte. Since the labeled polyelectrolyte always constitutes the outermost layer, and is not stripped off, it is very much unlikely that the unlabeled polyelectrolyte layers underneath could be stripped off leaving the labeled one still

Figure 5. 2f-FCS auto- and cross-correlation curves of the NGRh/PLLFITC, NGRh/(PLL/PSS)1.0/PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC as detected in the FRET channel at T ) 25 °C and T ) 40.7 °C.

Study of LbL Films on Thermoresponsive Nanogels anchored to the nanogel.8 Besides, the very fact that the intensity of the fluorescence emission is decreased (Figure 2) for NGRh/ (PLL/PSS)1.0/PLLFITC and NGRh/(PLL/PSS)2.0/PLLFITC nanogels as compared to NGRh/PLLFITC is indicative of the presence of the unlabeled underlying layers between the NGRh and PLLFITC, yet more evidence of successful buildup of polyelectrolyte multilayers on nanogels. So far, 2f-FCS has been used to quantitatively determine the diffusion of the coated ensemble as one entity from the ACFs and CCFs from both rhodamine and FITC signals. More readily available information that could be obtained from our 2f-FCS setup is the detection of fluorescence resonance energy transfer (FRET) in the FRET channel, the two fluorophores chosen in the present study constituting a donor-acceptor pair. However, it is very difficult to obtain quantitative FRET. Qualitative analysis of FRET can be done on two levels: when (1) the distance between NGRh and PLLFITC is varied and when (2) the temperature is varied. Figure 5 shows the FRET-ACFs and FRET-CCF of NGRh/PLLFITC, NGRh/(PLL/PSS)1.0/PLLFITC, and NGRh/(PLL/PSS)2.0/PLLFITC nanogels for two different temperatures: T ) 25 and 40.7 °C. The fact that FRET is detected in all the systems (NGRh/(PLL/PSS)n/PLLFITC with n ) 0, 1, 2) studied is clearly indicative of the close proximity of the fluorescent species. At 25 °C, the FRET signals detected are from the fully swollen systems, indicating that the layer thickness is still within the Fo¨rster radius and/or the PLLFITC is able to interdigitate with the underlying layers and reach NGRh. The ability of PLL to diffuse over the entire multilayer film has already been demonstrated even on planar substrates.31,32,34 Another study reported that, in a multicompartment film, two PSS/poly(allylamine) bilayers separating two compartments of multilayer film containing PLL are sufficient to act as a barrier to prevent further PLL diffusion over the entire film.32 Our templates are 3D soft and porous polymeric networks that are chemically (the nanogel) and physically (polyelectrolyte multilayers) interacting with each other and into which PLL can readily diffuse. In the future, we envisage using synthetic polyelectrolytes to provide insight into the blocking, transportation, or diffusion ability of the polyelectrolyte in microgels. At 40.7 °C, the availability and accessibility of rhodamine fluorophores from the nanogel and FITC fluorophores from the PLL increase as they are brought closer in vicinity to one another concomitant to the collapse of the network, and FRET signals are detected as expected. The spatiotemporal proximity of acceptor and donor molecules at 40.7 °C also proves that the polyelectrolytes do not get detached from the nanogel and that the nanogel core and the multilayer shell ensemble are still diffusing as one entity in the near collapsed state. Conclusion In summary, it is fair to say that study of the layer-by-layer (LbL) on microgels is at the same stage as when it first started on hard and rigid substrates some 15 years ago. The milestone in the present study resides in the fact that it is the first time a direct observation and calculation of the hydrodynamic radii from dual-focus fluorescence correlation spectroscopy (2f-FCS) is performed on polyelectrolyte-coated thermoresponsive submicrometer nanogels, thereby conclusively proving the motion of core and shell as one entity. Furthermore, performing 2fFCS as a function of temperature reveals, for the first time, that the polyelectrolyte multilayer shell is still bound to the nanogel during the volume phase transition. This makes not only 2fFCS a powerful technique to study submicrometer particles but

J. Phys. Chem. B, Vol. 113, No. 49, 2009 15913 also colloidal and supramolecular systems.30,35,36 Just like single particle light scattering renders studies by charge reversal qualitatively sine qua non to successful buildup of layers on hard and rigid particles, 2f-FCS accomplishes the same aim for soft and porous submicrometer particles and makes this technique an adequate tool to monitor LbL assembly on nanogels. The achievement of this study extends the possibility of assembling any charged species (polyelectrolytes, nanoparticles, or proteins) on microgels toward their potential exploitation as drug storage, transport, target, and controlled delivery. Acknowledgment. We are indebted to Jo¨rg Enderlein for stimulating and fruitful discussions. Financial support from the Deutsche Forschungsgemeinschaft (DFG) is gratefully acknowledged. References and Notes (1) Decher, G. Science 1997, 277, 1232. (2) Donath, E.; Sukhorukov, G. B.; Caruso, F.; Davis, S. A.; Mo¨hwald, H. Angew. Chem., Int. Ed. 1998, 37, 2202. (3) Caruso, F.; Caruso, R. A.; Mo¨hwald, H. Science 1998, 282, 1111. (4) Wang, Y.; Yu, A.; Caruso, F. Angew. Chem., Int. Ed. 2005, 44, 2888. (5) Wang, Y.; Caruso, F. Chem. Mater. 2005, 17, 953. (6) Greinert, N.; Richtering, W. Colloid Polym. Sci. 2004, 282, 1146. (7) Wong, J. E.; Richtering, W. Prog. Colloid Polym. Sci. 2006, 133, 45. (8) Wong, J. E.; Mu¨ller, C. B.; Laschewsky, A.; Richtering, W. J. Phys. Chem. B 2007, 111, 8527. (9) Sukhorukov, G. B.; Donath, E.; Lichtenfeld, H.; Knippel, E.; Knippel, M.; Budde, A.; Mo¨hwald, H. Colloids Surf. A 1998, 137, 253. (10) Pelton, R. H.; Chibante, P. Colloids Surf. 1986, 20, 247. (11) Ohshima, H. Colloid Polym. Sci. 2007, 285, 1411. (12) Wong, J. E.; Richtering, W. Curr. Opin. Colloid Interface Sci. 2008, 13, 403. (13) Wong, J. E.; Dı´ez-Pascual, A. M.; Richtering, W. Macromolecules 2009, 42, 1229. (14) Wong, J. E.; Dı´ez-Pascual, A. M.; Richtering, W., to be submitted. (15) Einstein, A.; Fu¨rth, R. InVestigations on the Theory of the Brownian MoVement; Dover: New York, 1956. (16) Madge, D.; Webb, W. W.; Elson, E. Phys. ReV. Lett. 1972, 29, 705. (17) Elson, E. L.; Madge, D. Biopolymers 1974, 13, 1. (18) Madge, D.; Elson, E. L.; Webb, W. W. Biopolymers 1974, 13, 29. (19) O’Connor, D. V.; Phillips, D. Time Correlated Single Photon Counting; Academic Press: London, 1984. (20) Mu¨ller, B. K.; Zaychikov, E.; Bra¨uchle, C.; Lamb, D. C. Biophys. J. 2005, 89, 3508. (21) Lamb, D. C.; Mu¨ller, B. K.; Bra¨uchle, C. Curr. Pharm. Biotechnol. 2005, 6, 405. (22) Mu¨ller, C. B.; Weiβ, K.; Richtering, W.; Loman, A.; Enderlein, J. Opt. Express 2008, 16, 4322. (23) Mu¨ller, C. B.; Richtering, W. Colloid Polym. Sci. 2008, 286, 1215. (24) Mu¨ller, C. B.; Loman, A.; Richtering, W.; Enderlein, J. J. Phys. Chem. B 2008, 112, 8236. (25) Schwille, P.; Meyer-Almes, F.-J.; Rigler, R. Biophys. J. 1997, 72, 1878. (26) Schwille, P.; Heinze, K. G. ChemPhysChem 2001, 2, 269. (27) Dertinger, T.; Pacheco, V.; von der Hocht, I.; Hartmann, R.; Gregor, I.; Enderlein, J. ChemPhysChem 2007, 8, 433. (28) Dertinger, T.; Loman, A.; Ewers, B.; Mu¨ller, C. B.; Kra¨mer, B.; Enderlein, J. Opt. Express 2008, 16, 14353. (29) Mu¨ller, C. B.; Loman, A.; Pacheco, V.; Koberling, F.; Willbold, D.; Richtering, W.; Enderlein, J. Europhys. Lett. 2008, 83, 46001. (30) Nayak, S.; Lyon, L. A. Angew. Chem., Int. Ed. 2005, 44, 7686. (31) Picart, C.; Mutterer, J.; Richert, L.; Luo, Y.; Prestwich, G. D.; Schaaf, P.; Voegel, J.-C.; Lavalle, P. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 12531. (32) Garza, J. M.; Schaaf, P.; Muller, S.; Ball, V.; Stoltz, J.-F.; Voegel, J.-C.; Lavalle, P. Langmuir 2004, 20, 7298. (33) Loman, A.; Dertinger, T.; Koberling, F.; Enderlein, J. Chem. Phys. Lett. 2009, 459, 18. (34) Porcel, C.; Lavalle, P.; Ball, V.; Decher, G.; Senger, B.; Voegel, J.-C.; Schaaf, P. Langmuir 2006, 22, 4376. (35) Cohen Stuart, M. A. Colloid Polym. Sci. 2008, 286, 855. (36) Liu, R. X.; Fraylich, M.; Saunders, B. R. Colloid Polym. Sci. 2009, 287, 627.

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