Journal of Nanoparticle Research

, Volume 13, Issue 1, pp 331–346

Fluorescent polymeric nanocomposite films generated by surface-mediated photoinitiation of polymerization

Authors

  • Heather J. Avens
    • Department of Chemical and Biological EngineeringUniversity of Colorado
  • Erin L. Chang
    • Department of Chemical and Biological EngineeringUniversity of Colorado
  • Allison M. May
    • Department of Chemical and Biological EngineeringUniversity of Colorado
  • Brad J. Berron
    • Department of Chemical and Biological EngineeringUniversity of Colorado
  • Gregory J. Seedorf
    • Department of PediatricsPediatric Heart Lung Center, University of Colorado Denver
  • Vivek Balasubramaniam
    • Department of PediatricsPediatric Heart Lung Center, University of Colorado Denver
    • Department of Chemical and Biological EngineeringUniversity of Colorado
Research Paper

DOI: 10.1007/s11051-010-0034-z

Cite this article as:
Avens, H.J., Chang, E.L., May, A.M. et al. J Nanopart Res (2011) 13: 331. doi:10.1007/s11051-010-0034-z

Abstract

Incorporation of nanoparticles (NPs) into polymer films represents a valuable strategy for achieving a variety of desirable physical, optical, mechanical, and electrical attributes. Here, we describe and characterize the creation of highly fluorescent polymer films by entrapment of fluorescent NPs into polymer matrices through surface-mediated eosin photoinitiation reactions. Performing surface-mediated polymerizations with NPs combines the benefits of a covalently anchored film with the unique material properties afforded by NPs. The effects of monomer type, crosslinker content, NP size, and NP surface chemistry were investigated to determine their impact on the relative amount of NPs entrapped in the surface-bound films. The density of entrapped NPs was increased up to 6-fold by decreasing the NP diameter. Increasing the crosslinking agent concentration enabled a greater than 2-fold increase in the amount of NPs entrapped. Additionally, the monomer chemistry played a significant role as poly(ethylene glycol) diacrylate (PEGDA)-based monomer formulations entrapped a 10-fold higher density of carboxy-functionalized NPs than did acrylamide/bisacrylamide formulations, though the latter formulations ultimately immobilized more fluorophores by generating thicker films. In the context of a polymerization-based microarray biodetection platform, these findings enabled tailoring of the monomer and NP selection to yield a 200-fold improvement in sensitivity from 31 (±1) to 0.16 (±0.01) biotinylated target molecules per square micron. Similarly, in polymerization-based cell staining applications, appropriate monomer and NP selection enabled facile visualization of microscale, sub-cellular features. Careful consideration of monomer and NP selection is critical to achieve the desired properties in applications that employ surface-mediated polymerization to entrap NPs.

Keywords

Fluorescent polymersPolymeric nanocompositesPhotopolymerizationSurface-mediated polymerizationFluorescent immunocytochemistryProtein microarrays

Introduction

Polymeric nanocomposites are of significant interest, heralded for achieving further enhancements in material properties as compared to microcomposite approaches (Allegra et al. 2008). Incorporation of nanomaterials in polymeric matrices has been employed in a variety of applications including generation of materials with enhanced optical, magnetic, electrical, thermal, and mechanical properties (Balazs et al. 2006; Dhibar et al. 2009; Durán et al. 2008; Luo et al. 2009; Verma et al. 2009). Additionally, nanocomposite coatings have been developed that achieve various desirable surface modifications including superhydrophobicity (Xu et al. 2009), corrosion resistance (Olad and Rashidzadeh 2008), reduced gloss (Balan et al. 2008), and enhanced mechanical strength (Zhu et al. 2007).

Recently, the use of fluorescent nanocomposite films has been reported for sensitive and photostable biodetection in microarrays and immunocytochemical staining (Avens and Bowman 2010; Avens et al., submitted to Journal of Histochemistry and Cytochemistry; Hansen et al. 2008a). This approach, coined “fluorescent polymerization-based amplification” (FPBA), utilizes biological probes selectively labeled with eosin photoinitiators. After the probe has bound its target on the microarray or within the cell, a solution of monomer, coinitiator, and fluorescent nanoparticles is applied and the test surface is exposed to light of wavelengths greater than 480 nm to initiate polymerization. This approach results in the formation of crosslinked polymer films specifically in regions of the microarray or cell where the probe has bound its target. As the polymer film forms, it entraps the fluorescent nanoparticles, rendering the film highly fluorescent. In this manner, the presence and localization of a biological target is evidenced by a highly fluorescent nanoscale polymer film. During the development of FPBA, other strategies for generating fluorescent films were considered, such as using monomers covalently attached to organic fluorophores; however, this approach was unsuitable because the organic fluorophores themselves non-specifically initiated polymerization and/or underwent extensive photobleaching (Avens and Bowman 2010). To overcome these challenges, it was necessary to use polystyrene nanoparticles with fluorophores embedded in the interior which enabled facile formation of fluorescent films with fewer problematic side reactions between the monomer and the fluorophores. FPBA has been demonstrated to yield approximately 100-fold brighter signals and a 100-fold improvement in detection sensitivity compared to using a molecular fluor-labeled probe (Avens and Bowman 2010; Avens HJ et al., submitted to Journal of Histochemistry and Cytochemistry). Additionally, in cell staining applications, FPBA compared favorably to the highly sensitive tyramide signal amplification (TSA) approach which relies on peroxidase enzymes. Moreover, while both TSA and FPBA generated similarly intense fluorescent signals, FPBA is advantageous in that it is not affected by endogenous cellular peroxidase enzymes which cause non-specific TSA staining (Avens HJ et al., submitted to Journal of Histochemistry and Cytochemistry).

Though a variety of surface-mediated initiation strategies are suitable for generation of NP-polymer composite films, the method presented here employs surface-immobilized eosin as a type II photoinitiator for polymerization. Upon photoexcitation with visible light, eosin undergoes energy, charge, and electron transfer with a coinitiator, commonly a tertiary amine such as N-methyldiethanolamine (MDEA), to yield a relatively stable eosin radical and an MDEA radical capable of initiating polymerization (Avens and Bowman 2009). Despite the fact that the initiating MDEA radicals are not tethered to the surface, the reaction is still considered surface mediated because the initiating radicals are generated only at the surface. Covalent attachment of the films to the surface is believed to occur through a variety of chain transfer reactions and other processes including termination of the polymer chains with the eosin radical which is reported to be inactive for initiation, but capable of reacting by termination (Kizilel et al. 2004). Eosin has been used for surface-mediated polymerization for an assortment of purposes including modification of nanoparticle surfaces to enhance dispersion (Satoh et al. 2005), islet cell encapsulation (Cruise et al. 1998), creation of protective surface coatings for arterial walls (An and Hubbell 2000), and generation of macroscopically visible and fluorescent films for sensitive signal amplification of biodetection (Hansen et al. 2008a; Avens and Bowman 2010). The fact that eosin uses visible light which is less damaging to biological systems than UV light, and the observation that eosin is less sensitive to oxygen inhibition than other photoinitiators (Avens and Bowman 2009) makes eosin an excellent choice for numerous surface modification applications. Importantly, photoinitiation, as opposed to other initiation strategies, readily enables precise spatial and temporal control of the polymerization process, factors which are often critical for achieving the desired material properties. (Bowman and Kloxin 2008). Though photoinitiation is the focus of this work, the NP incorporation results presented here are expected to be generalizable to other surface-mediated polymerization processes including other polymerization-based detection strategies (He et al. 2008; Lou et al. 2005).

For FPBA and other applications that employ surface-mediated polymerization with NPs, it is necessary to optimize the NP and monomer attributes to achieve the required polymer properties. To more aptly employ this type of nanocomposite film formation for biodetection and other applications, a detailed study is presented here investigating the effects of NP size, NP surface functionality, monomer type, and crosslinking agent content on the fluorescence and thickness of the polymer films. For FPBA, it is generally desired to incorporate as high a density of NPs as possible to maximize the subsequent fluorescent signal emanating from the surface. It is hypothesized that chemical and mechanical interactions occurring at the polymer–monomer–NP interface during the polymerization strongly influence the successful entrapment of NPs into the growing polymer matrix. To identify trends that are relevant across a broad range of initiation rates, surface-mediated polymerizations were carried out using a microarray format in which each row of spots comprises a different eosin surface density, such that a wide range of eosin photoinitiator densities were evaluated simultaneously. Reactions were performed with both an acrylamide-based monomer formulation and a poly(ethylene glycol) (PEG) acrylate-based formulation where the acrylamide polymerization is known to occur more rapidly and the PEG-based formulation is more hydrophilic. Trends that are common between these two monomer formulations are expected to be pertinent to other monomer types as well, while differences are correlated to the differences in the chemistry and reaction behavior of the formulations.

The significance of the trends presented here was investigated in the context of two specific applications. First, it was shown that through selection of the appropriate monomer type, crosslinker content and NP size one is able to significantly increase the absolute amount of fluorescent NPs immobilized on the surface, yielding a 200-fold improvement in detection sensitivity in a polymerization-based microarray biodetection platform compared to using sub-optimal reaction conditions. Second, by selecting parameters to optimize the NP density entrapped in the film, rather than just the total NP amount immobilized to the surface, it was possible to create highly fluorescent, yet thin and spatially restricted films suitable for visualization of microscale sub-cellular features in polymerization-based fluorescent immunostaining of cells. The principles outlined here are expected to be useful for tailoring monomer and NP attributes to achieve desired properties in additional applications in which NPs are being entrapped in surface-mediated polymerizations, including in the context of different polymer matrices and different types of NPs.

Experimental

Materials

Poly(ethylene glycol) diacrylate, Mn = 575 (PEGDA), poly(ethylene glycol) acrylate, Mn = 375 (PEGA), 40 wt% acrylamide in water, N,N′-methylenebisacrylamide (bisacrylamide), N-methyldiethanolamine (MDEA), vinyl pyrrolidone (VP), 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI), streptavidin (SA), biotin-labeled anti-goat IgG (bi-antibody) produced in rabbit, bovine serum albumin (BSA), and 10× PBS were purchased from Sigma–Aldrich. Eosin–isothiocyanate, carboxylate-functionalized yellow/green NPs (20, 40, 100, and 200 nm diameter FluoSpheres), carboxylate-functionalized dark red NPs (20, 40, and 200 nm diameter FluoSpheres), amine-functionalized yellow/green NPs (200 nm diameter FluoSpheres), and carboxylate-functionalized crimson NPs (20 nm diameter FluoSpheres) were purchased from Invitrogen. Table 1 provides the actual NP diameters and relative quantum yields as provided by Invitrogen for each of the NPs utilized. TEM images of FluoSpheres have been published previously (Kobayashi et al. 2004; Lansalot et al. 2005; Popielarski et al. 2005). Epoxy-functionalized glass slides (SuperEpoxy 2), 2× Protein Printing Buffer and solid printing pins were purchased from Telechem International, Inc. Biotin-labeled anti-mouse IgG produced in goat was purchased from Pierce Biotechnology while anti-nuclear pore complex proteins IgG (anti-NPC) produced in mouse was purchased from Covance. Streptavidin–eosin conjugate (SA–eosin) was prepared in-house as described previously (Hansen et al. 2008b) by reacting eosin isothiocyanate with amines on the protein’s surface. A Cy3 Scanner Calibration slide was purchased from Full Moon BioSystems. Water was purified using a Milli-Q system. The cells used for immunofluorescent imaging are endothelial colony forming cells whose isolation has been described previously (Baker et al. 2009).
Table 1

Characteristics of NPs used for these studies

Color

Surface functionality

Nominal diameter (nm)

Actual diameter (nm)

Relative QYa

Relative absorbanceb

Yellow/green

Carboxylate

20

24 ± 4

0.33

1.00

Yellow/green

Carboxylate

40

43 ± 6

0.47

1.37

Yellow/green

Carboxylate

100

100 ± 6

0.39

0.85

Yellow/green

Carboxylate

200

210 ± 10

0.17

0.93

Yellow/green

Amine

200

190 ± 11

0.18

0.93

Dark red

Carboxylate

20

24 ± 3

1.60

1.00

Dark red

Carboxylate

40

36 ± 5

1.45

0.46

Dark red

Carboxylate

200

210 ± 10

0.46

0.46

Crimson

Carboxylate

20

24 ± 4

1.00

N/A

These data are provided by Invitrogen on the certificate of analysis for each NP Fluosphere product

aFor yellow/green NPs, the QY is relative to a solution in methanol of the dye used to prepare the NPs. For dark red NPs and the crimson NPs, the QY is relative to a solution in chloroform of the dye used to prepare the NPs

bFor NPs of the same color, but different sizes or surface functionalities, relative absorbance values were determined. The absorbance values are per mass of NPs, and the 20 nm NP absorbance values are set to 1.0

Compare photoluminescence of NPs in solution and in polymer

To verify that encapsulation in polymer films does not dramatically alter NP photoluminescence, an Ocean Optics USB4000-FL detector was used to measure emission spectra of free 20 nm yellow/green NPs in water (0.05 wt% NPs) as well as NPs that had been entrapped in a polyacrylamide film analogous to the ones formed on the surfaces. The excitation source was a 370 nm CrystaLaser. The samples were prepared in 1 cm wide UV/VIS cuvettes.

Preparation of eosin dilution chips

Streptavidin–eosin was printed onto epoxy-functionalized glass slides using a VersArray Chip WriterTM Pro (Bio-Rad) with approximately 45% humidity and a solid pin yielding spots of approximately 500 μm diameter. 9 × 5 arrays of spots were prepared containing five replicate spots of 8 decreasing eosin concentrations, and a 9th row with no eosin that served as a negative control suitable for evaluating non-specific polymerization. The highest print concentration was 0.27 mg/mL SA–eosin (12 μM eosin, 1.3 eosin per streptavidin) in a final concentration of 1× printing buffer. Lower print concentrations were prepared by serial 1:3 dilutions into 1× print buffer with unmodified streptavidin such that all spots were printed with a total protein concentration of 0.27 mg/mL. Unbound SA–eosin was removed by three 15 min rinses in water with rapid rocking. Since eosin is a fluorophore, which also serves as the initiator, surface density was characterized with an Agilent Technologies Microarray Scanner. A Cy3 Scanner Calibration slide was used to convert fluorescence readings into surface density of eosin. The slides were then polymerized as described subsequently.

Preparation of biotin dilution chips

Microarray printing as described for SA–eosin was likewise performed to create slides with a dilution series of biotin-α-goat IgG in a 9 × 5 array containing five replicate spots of 8 decreasing antibody concentrations, and a 9th row with no antibody that served as a negative control suitable for evaluating non-specific polymerization. The highest print concentration was 85 μg/mL biotin-α-goat IgG in 1× printing buffer and 0.5 mg/mL BSA. Lower print concentrations were prepared by serial 1:3 dilutions into 1× print buffer with 0.5 mg/mL BSA, and the 9th row contained only 0.5 mg/mL BSA. The slides were stored unprocessed until they were used for binding and detection reactions. The surface concentration of bound antibodies was estimated as described previously (Avens and Bowman 2010).

Binding reactions with the biotin dilution chips

The biotin-α-goat IgG dilution chips were rinsed with water to remove unbound antibody, then the slides were incubated with 10 mg/mL BSA in 1× PBS for 1 h, followed by three 2 min rinses in PBS with rapid rocking. The dilution chips were then reacted 1 h with 1 μg/mL SA–eosin, using Chip Clips from Whatman to form wells on the chips and gentle rocking was used, followed by three 2 min rinses in PBS with rapid rocking and one rinse in water. The slides were then polymerized as described subsequently.

Cell staining procedures

Cells in micro-chamber slides were rinsed with 1× PBS then fixed with 4% paraformaldehyde for 15 min. The slides were then stored in 1× PBS and refrigerated until staining. At the time of staining, slides were rinsed 3 min with 1× PBS, permeabilized with 0.1% Triton-X in PBS for 5 min, then rinsed three times with PBS. Next, the slides were blocked for 1 h with 2% horse serum and 0.1% BSA in PBS, and then rinsed three times with 0.1% BSA in PBS (PBSA). Anti-NPC IgG at a dilution of 1:1000 in PBSA was applied for 1 h, followed by three rinses with PBSA. Next, biotin-anti-mouse IgG was applied for 1 h at a dilution of 1:400 in PBSA, followed by three rinses with PBSA. The slides were then contacted with 10 μg/mL SA–eosin for 30 min, followed by three rinses with PBS and one rinse with water. Polymerization reactions were then carried out as described in the following section. After polymerization, the nuclei of the cells were stained 2 min with 1 μg/mL DAPI in water.

Polymerization reactions

The microarray or cell slide surfaces were contacted with the desired monomer solution. For 5 min prior to and throughout the entire light exposure, the slides were placed in a plastic bag with argon flow to reduce oxygen in the atmosphere. The light source was an Acticure (Exfo) high pressure mercury lamp with an in-house internal bandpass filter (350–650 nm) and an external 490 nm longpass filter (Edmund Optics) positioned at the end of a light guide and a collimating lens. The light intensity was measured using an International Light radiometer. After polymerization, unreacted monomer was removed with three-5 min water rinses, followed by air drying. To obtain sufficient replicates in the microarray experiments, each condition investigated was polymerized on at least 2–3 arrays in 2–3 separate polymerization sessions.

Film thickness measurements

Polymer film thicknesses were measured using a Dektak 6 M surface profilometer with 12.5 μm diameter tip and a stylus force of 1 mg.

Fluorescent imaging of films

Cells stained by fluorescent polymerization-based signal amplification were imaged by confocal scanning laser microscopy (Zeiss LSM 510 instrument) with a 40× oil objective. Yellow/green NP fluorescence on the microarray surfaces was measured using a Leica MZ FLIII stereomicroscope (Leica Microsystems, Wetzlar, Germany) with the blue filter set. Exposure times in the range of 5 to 60 s were used, depending on the intensity of the fluorescent signal. A Cy3 Scanner Calibration slide was imaged at each exposure time to identify appropriate scaling factors for the different exposure times, such that all data could be plotted using the same arbitrary fluorescence scale. Crimson and dark red NP fluorescence on the microarray surfaces were measured using the red channel of an Agilent Technologies Microarray Scanner. Significant positive signals are those that have a signal to noise (S/N) greater than 3, where [S/N = (signal − background signal)/(standard deviation of the background signal)]. The film fluorescence values have been normalized to account for the differences in relative quantum yield between NPs of different sizes or different surface chemistries, utilizing the relative quantum yields provided in Table 1. Additionally, for NPs of the same color, but different sizes or surface functionalities, the film fluorescence values were normalized to account for the slight differences in the NP fluorophore content per mass, as determined by absorbance measurements (Table 1). In this way, the reported fluorescence intensities are more indicative of the mass of NPs that has been immobilized in the films, rather than simply variations in the manufacturing of these NPs.

Statistical analysis

In cases where a “gain value” is reported, linear regression with a least squares parameter estimation was performed to best fit a line of the form y = mx (intercept set to 0) to the data. The slope “m” is termed “gain” and the 95% confidence interval on the slope is indicated.

Mesh size determinations

Mesh sizes were estimated by measuring the equilibrium swelling ratio (Q) of the gels as described by Bryant and Anseth (2006). Photoinitiation of the indicated monomer formulations was achieved with 4 μM eosin–isothiocyanate to generate disc shaped hydrogels of approximately 1 mm thick and 8 mm in diameter. The gels were rinsed 24 h with several changes of water. The swollen weight (Ms) was recorded and then the gels were dried under vacuum at 37°C for 3 days after which the dried weight (Md) was recorded. Q was calculated as shown in Eq. 1, where ρs is the solvent density (1 mg/mL) and ρp is the polymer density, 1.12 mg/mL for PEG gels and 1.35 mg/mL for polyacrylamide gels.
$$ Q = 1 + {\frac{{\rho_{\text{p}} }}{{\rho_{\text{s}} }}}\left( {{\frac{{M_{\text{s}} }}{{M_{\text{d}} }}} - 1} \right) $$
(1)
The average molecular weight between crosslinks (Mc) is calculated as shown in Eq. 2, where V1 is the molar volume of water (18 mL/mol), v2,s is the equilibrium polymer volume fraction (1/Q), X12 is the solvent-polymer interaction parameter, 0.43 for PEG-based gels and 0.48 for polyacrylamide gels. Except for the gels composed of PEGDA with no PEGA, the term 2/Mn is neglected as Mc ≪ Mn.
$$ {\frac{1}{{\bar{M}_{\text{c}} }}} = - {\frac{{\left( {{1 \mathord{\left/ {\vphantom {1 {\rho_{\text{p}} V_{1} }}} \right. \kern-\nulldelimiterspace} {\rho_{\text{p}} V_{1} }}} \right)\left[ {\ln \left( {1 - v_{{2,{\text{s}}}} } \right) + v_{{2,{\text{s}}}} + X_{12} v_{{2,{\text{s}}}}^{2} } \right]}}{{\left[ {v_{{2,{\text{s}}}}^{{{1 \mathord{\left/ {\vphantom {1 3}} \right. \kern-\nulldelimiterspace} 3}}} - {\frac{{v_{{2,{\text{s}}}} }}{2}}} \right]}}} - {\frac{2}{{\bar{M}_{\text{n}} }}} $$
(2)
Finally, the average mesh sizes (ε) for the swollen hydrogels are calculated as shown in Eq. 3, where Cn is the characteristic ratio of the polymer (4 for PEG-based gels and 14.8 for the polyacrylamide gels), l is the bond length, and n is the number of bonds between the crosslinks.
$$ \varepsilon = v_{{2,{\text{s}}}}^{{ - {1 \mathord{\left/ {\vphantom {1 3}} \right. \kern-\nulldelimiterspace} 3}}} C_{n}^{{{1 \mathord{\left/ {\vphantom {1 2}} \right. \kern-\nulldelimiterspace} 2}}} n^{{{1 \mathord{\left/ {\vphantom {1 2}} \right. \kern-\nulldelimiterspace} 2}}} l $$
(3)

The observation that PEGA with no added crosslinker readily forms a crosslinked gel highlights the fact that a significant amount of chain transfer occurs in the PEGA and PEGDA systems, which creates ambiguity concerning the chemical composition of the chains between crosslinks. Thus, the average mesh sizes for the PEG-based gels were calculated by two distinct methods to provide reasonable bounds on the probable average mesh sizes: (1) n = 2Mc/Mr, where Mr is the molecular weight of the repeating monomer unit assuming no chain transfer; and (2) assuming extensive chain transfer, n = 3Mc/Mr, where Mr is the molecular weight of the PEG repeat unit. The value of three in the second method arises from the fact that the PEG repeat unit (COO) contains three bonds. In the case of polyacrylamide, n = 2Mc/Mr, where Mr is 71 g/mol.

Results and discussion

Compare the photoluminescence of NPs in solution and in the polymer film

The photoluminescent properties of these NPs are not expected to change upon encapsulation in polymer films, as the fluorophores in the NPs are already embedded in the NP-polymer matrix. To verify that encapsulation in polymer does not dramatically alter NP photoluminescence, emission spectra were compared between free 20 nm yellow/green NPs in water (0.05 wt% NPs) and NPs that had been entrapped in a 1 cm thick polyacrylamide film (Fig. 1). Entrapment in polyacrylamide resulted in only a 9% drop in the maximum fluorescent signal and the shape of the emission spectra was not significantly altered. These results indicate that NP photoluminescence is not altered in a meaningful manner either by the polymerization reaction or by being embedded in additional polymer matrix.
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Fig. 1

Comparison of the fluorescence emission spectra for 20 nm yellow/green NPs either in water (solid line) or entrapped in a polyacrylamide gel (dotted line). NPs were 0.05 wt%. The monomer formulation was 5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, and 35 mM VP in water. The polymerization was photoinitiated using 500 ng/mL eosin and exposure to 40 mW/cm2 light of wavelengths greater than 480 nm

Effect of NP size

In order to ascertain the impact of NP diameter on the ability of fluorescent NPs to become encapsulated in films generated from surface-mediated photoinitiation, polymerizations were performed with monomer solutions containing fluorescent NPs of a variety of distinct diameters ranging from 20 to 200 nm. Additionally, a microarray format was employed such that a wide range of eosin photoinitiator surface densities were evaluated simultaneously, enabling investigation of the effect of NP size at various initiator surface densities. Using various eosin initiator surface densities ensures that the observed trends for NP size are generalizable to varying initiation rates. Finally, two different monomer formulations were employed, an acrylamide-based formulation and a PEGDA-based formulation.

In the context of generating fluorescent films via NP entrapment, one can envision achieving improved fluorescent signals in two ways: first, by generating thicker films; or, second, by incorporating a higher density of NPs into the films. To take into account both these contributions, the data presented here are evaluated not only for overall fluorescence generated as a function of eosin initiator surface density, but also for both how thick the films are and how much fluorescence is seen as a function of film thickness. Figure 2 shows a typical data set comprising the results from photopolymerizations of an acrylamide monomer formulation containing 0.05 wt% 100 nm yellow/green NPs. Three terms are introduced to facilitate analysis of these data: (1) “overall gain” refers to the slope on the plot of eosin surface density (x) versus film fluorescence (y) (Fig. 2a) and is indicative of the increase in film fluorescence observed as photoinitiator surface density increases; (2) “fluorescence gain” denotes the slope on the plot of film thickness (x) versus film fluorescence (y) (Fig. 2b) and signifies the amount of increase in film fluorescence observed with an increase in film thickness; and (3) “thickness gain” refers to the slope on the plot of eosin surface density (x) versus film thickness (y) (Fig. 2c) and describes the increase in film thickness associated with a particular increase in photinitiator surface density. In all cases, the gain is reported to be the least squares fit of the proportionality constant for the two relevant factors. The three types of gains are related in that the fluorescence gain multiplied by the thickness gain is roughly equal to the overall gain; however, due to the error associated with linear regression, this relationship is not exact. Nearly all the data sets were sufficiently linear such that the p values for the slopes were many orders of magnitude less than one. The exceptions were the overall gain and the fluorescence gain for the 200 nm dark red NPs depicted in Fig. 4, whose p values were greater than 0.05. The fact that the film fluorescence typically increases monotonically with film thickness suggests that the NPs are evenly distributed throughout the thickness of the films, which is consistent with previously published results that also reported a linear relationship between film thickness and film fluorescence (Avens and Bowman 2010). A comparison of the gain values for polymerizations performed under different conditions enables determination of which conditions yield improved fluorescence amplification, and whether that amplification is a result of increased film thickness, a higher density of NP incorporation, or both.
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Fig. 2

Introduction of terminology for characterization of fluorescent films generated from a range of eosin photoinitator surface densities. Here, 0.05 wt% 100 nm yellow/green carboxylate-functionalized NPs were used in an acrylamide monomer formulation (5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, and 35 mM VP in water). a The term “Overall Gain” is introduced to describe the total increase in film fluorescence seen with an increase in eosin photoinitiator surface density. b The term “Fluorescence Gain” is introduced to indicate the increase in the observed fluorescence as a result of an increase in film thickness. c The term “Thickness Gain” is introduced to describe the change in film thickness achieved by an increase in eosin photoinitiator surface density. Polymerization was initiated by a 30 min exposure to 40 mW/cm2 light at wavelengths greater than 480 nm

Surface-mediated initiation of polymerization of an acrylamide monomer formulation with varying sizes of yellow/green NPs (0.05 wt%) reveals that the smallest NP diameter, 20 nm, results in 4- to 6-fold greater overall gain compared to the larger NP sizes. These results indicate that the 20 nm NP size favors immobilization of a higher mass of NPs onto the surface (Fig. 3a). Further, as the fluorescence gains are 3- to 6-fold higher for the 20 nm NPs (Fig. 3b), it is concluded that polymerization with 20 nm NPs enables a higher density of NPs (on a mass-basis) to become entrapped in the films. The ability of 20 nm NPs to incorporate better into the films is the dominant contribution to the overall gain, as there is no clear trend between thickness gain and NP diameter (Fig. 3c). Indeed, Fig. 3c demonstrates that film thickness is determined by the eosin photoinitator surface density and shows no clear correlation with NP size.
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Fig. 3

Comparison of the overall gain (a), fluorescence gain (b), and thickness gain (c) measured for polymerizations performed with carboxylate-functionalized NPs of various sizes (0.05 wt%) added to an acrylamide monomer formulation (5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, and 35 mM VP in water). Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. Films generated in the absence of NPs were not fluorescent. The letters denote groups of conditions that are statistically different (α = 0.05)

Similar experiments were also performed using a PEGDA monomer formulation with varying sizes of dark red NPs. To facilitate a more concise presentation of the data, the gains were normalized to a scale of 0–1 by dividing each gain by the largest gain in the data set. In this way, it is possible to plot the overall gain, fluorescence gain, and thickness gain all on a single chart, as shown in Fig. 4. Online Resource 1 contains tables of all the non-normalized gain values for the figures in this chapter that display normalized gains. In agreement with the acrylamide results, it is found that the 20 nm NP size favors improved fluorescence signal compared to the 40 nm NP size, primarily as a result of better incorporation of the 20 nm NPs into the films, rather than by growth of thicker films (Fig. 4). The increase in overall gain and fluorescence gain associated with the 20 nm particles in PEGDA is not as dramatic as was observed with the acrylamide formulation. This behavior suggests that factors other than NP size are more critical in determining NP incorporation in the PEGDA system than in the acrylamide system. Additionally, the large error associated with the 200 nm NPs is attributed to sporatic non-specific binding of the 200 nm particles to the surface, and indicates that this size NP is unsuitable for use with the PEGDA monomer formulation in biodetection applications that require quantitation and specificity.
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Fig. 4

Normalized overall gain (solid squares), normalized fluorescence gain (open circles), and normalized thickness gain (open triangles) for polymer films generated by surface-mediated initiation with eosin using a PEGDA monomer formulation (420 mM PEGDA, 210 mM MDEA, and 35 mM VP in water) containing carboxylate-functionalized dark red NPs of various sizes (0.05 wt%). Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. Within each gain type, normalization is performed by dividing each value by the highest gain value. For the overall gain and fluorescence gain, the 20 nm condition is significantly different (α = 0.05) than the 40 nm condition. Also, the thickness gain for the polymerization without NPs (0 nm) is significantly different (α = 0.05) from the 20, 40, and 200 nm conditions

The enhanced incorporation of smaller NPs likely arises from the interfacial nature of the surface-initiated polymerization and the evolving crosslinked network that is forming. Though the polymerization is initiated from a uniform surface, heterogeneities and differential extension of the polymer film into the bulk monomer will undoubtedly arise, particularly in these crosslinked materials that are notorious for microgel formation and heterogeneity (Hutchison and Anseth 2001; Kloosterboer 1988). The smaller the NP is, the more likely it is to become entrapped in this evolving structure.

Effect of NP surface chemistry

In light of the evolving interface between the polymerizing mixture and the bulk monomer, it is anticipated that chemical interactions between the polymer and functional groups on the NP surface are likely to impact incorporation of fluorescent NPs into polymer films. To investigate the effect of surface modifications, a comparison was made between carboxylate versus amine modification of the fluorescent NPs. Figure 5 shows that amine-functionalized NPs are preferentially incorporated into the polyacrylamide films compared to the carboxylate-functionalized NPs, as evidenced by their higher fluorescence gain; however, the amine-functionalized NPs yield thinner films than the carboxylate-functionalized NPs. The net effect is that the amine-functionalized NPs trend towards having higher overall gains than the carboxylate NPs, though this difference is not found to be significant (α < 0.05). The polymerization is performed at pH = 9, much higher than the pKa of carboxylic acid (~5) and lower than the pKa of ammonium ions (~11) (Jones 1997); therefore, during polymerization, the amine-functionalized particles have positive surface charges, while the carboxy-functionalized particles have negative surface charges. It is not immediately apparent that either of these functional groups would react preferentially with polyacrylamide, as the carboxyl group on acrylamide has a partial negative charge, while the amine on acrylamide has a partial positive charge. However, amines are known to undergo chain transfer more readily than carboxylates, even being capable coinitiators for a variety of photosensitization systems, including eosin. Generation of radicals on the amine-functionalized NPs would likely lead to covalent bond formation with the polymer network through polymerization and subsequent crosslinking from the radical formed as a result of the chain transfer process. This hypothesis also explains the thinner films associated with the amine-functionalized particles as the additional chain transfer will slow the polymerization.
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Fig. 5

Normalized overall gain (solid squares), normalized fluorescence gain (open circles), and normalized thickness gain (open triangles) for polymer films generated by surface-mediated initiation with eosin using an acrylamide monomer formulation (5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, and 35 mM VP in water) with 0.05 wt% 200 nm yellow/green nanoparticles. Use of carboxy- versus anime-functionalized NPs is compared. Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. Within each gain type, normalization is performed by dividing each value by the highest gain value. For fluorescence gain and thickness gain, the observed difference in gain between the amine- and carboxy-functionalized surfaces is statistically significant (α = 0.05)

Effect of crosslinker concentration

Increasing the crosslinking density of a polymer network increases the polymer modulus and decreases the average mesh size. If the average mesh size is on the order of the particle diameter, then changes in the crosslinking content of the monomer formulation would be expected to impact the ability of these gels to entrap fluorescent NPs effectively. To investigate how crosslinking density affects the incorporation of nanoparticles during a surface-mediated polymerization, polymerizations were performed with varying bisacrylamide crosslinker concentrations (0, 1, and 2 wt%) added to an acrylamide monomer formulation. It was found that the higher crosslinking agent content favors incorporation of NPs into the polymer films, yielding a more than 2-fold increase in fluorescence gain, and ultimately resulting in a greater number of fluorescent species immobilized on the surface (i.e., greater overall gain) (Fig. 6). When bisacrylamide is absent, no polymer film is detected by profilometry or by fluorescence. In the absence of any bisacrylamide crosslinking agent, the polymerization rate is too slow to generate detectable films under these conditions.
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Fig. 6

Normalized overall gain (solid squares), normalized fluorescence gain (open circles), and normalized thickness gain (open triangles) for polymer films generated by surface-mediated initiation with eosin using different concentrations of bisacrylamide added to 5.2 M acrylamide, 210 mM MDEA, 35 mM VP, and 0.05 wt% 20 nm carboxylate-functionalized yellow/green NPs in water (2 wt% bisacrylamide is 130 mM). Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. Within each gain type, normalization is performed by dividing each value by the highest gain value. Within each gain type, all the values are significantly different (α = 0.05)

A similar trend is seen in the gels formed from varying ratios of PEGDA and PEGA that achieve significant variations in crosslinking density and network mesh size. As the PEGDA content is increased, the fluorescence yield and overall yield also increase (Table 2), indicating more efficient trapping of the fluorescent NPs. On the other hand, the effect of crosslinker content on thickness gain does not exhibit any clear trend. This behavior is likely due to the fact that a more crosslinked gel has two conflicting influences: (1) it leads to an increased polymerization rate as termination becomes less facile; and (2) in a surface-mediated eosin polymerization, the more crosslinked gel limits diffusion of MDEA to the surface and hinders its surface initiated radicals, when formed, from reaching the monomer-rich regions, thereby slowing the rate. Also, unlike the polyacrylamide system, PEGA polymerized without any additional crosslinking agent does form a detectable, insoluble film, most likely due to a small amount of diacrylate impurity present in the monoacrylate and/or chain transfer reactions which lead to crosslink formation in combination with termination by combination.
Table 2

Gain values measured for fluorescent films formed from varying concentrations of PEGA and PEGDA

Diacrylate/Monoacrylate (wt%/wt%)

Overall gain (NP fluorescence/(eosin/μm2))

Fluorescence gain (NP Fluorescence/nm)

Thickness gain (nm/(eosin/μm2))

0/22

26 (±3)A

220 (±30)A

0.094 (±0.023)A,B

1/21

27 (±4)A

390 (±30)B

0.066 (±0.013)B

11/11

55 (±6)B

370 (±60)B

0.13 (±0.02)A

22/0

57 (±5)B

570 (±110)C

0.084 (±0.014)B

Varying PEGDA and PEGA concentrations are added to 210 mM MDEA, 35 mM VP, and 0.05 wt% 20 nm Crimson NPs in water (22 wt% PEGDA is 420 mM, and 22 wt% PEGA is 650 mM). Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. Capital letters denote groups of conditions that are statistically different (α = 0.05)

In order to investigate further the effect of crosslinking density on the incorporation of fluorescent NPs, the molecular weight between crosslinks (Mc) and the average mesh size were determined for hydrogels created using each monomer formulation described in Fig. 6 and Table 2. In the polyacrylamide system, 2 wt% bisacrylamide resulted in approximately 60% smaller average mesh sizes (9 vs. 14 nm) than the gels made with 1 wt% bisacrylamide (Table 3). For the PEG gels, variations in the crosslinker content yielded a range of mesh sizes from 2 to 19 nm (Table 4), demonstrating the possibility for up to a 10-fold variation in mesh size in gels generated from these monomer formulations. The diameters of the NPs employed in the crosslinker experiment (24 ± 4 nm for both the yellow/green and the Crimson NPs) are slightly greater than the largest average mesh size determined here, indicating that it is unlikely these NPs diffuse out of the gel once they are entrapped; however, at the monomer–polymer interface where NP incorporation occurs, the NPs are interacting with nascent polymer networks with incomplete mesh structures, which are likely larger than the final mesh size within the bulk of the gel. It seems probable that average internal mesh sizes in the range of 2–19 nm would correspond to incomplete mesh structures at the polymer–monomer interface that are on a size scale that would impact the efficiency of trapping 24 nm diameter NPs. It is posited that gels with smaller internal mesh size have incomplete mesh structures at the polymer–monomer interface that are smaller, thereby limiting diffusion of the NPs away from the interface into the monomer. In addition, the gels with smaller mesh sizes necessarily have a faster rate of mesh formation for a given polymerization rate, providing less time for NPs at the polymerization front to diffuse away before becoming entrapped. Thus, the enhanced NP entrapment observed with increased crosslinker content is likely attributable to a combination of changes in the average mesh size at the polymer–monomer interface and differences in the rate of mesh formation.
Table 3

Estimated molecular weight between crosslinks (Mc) and mesh size in polyacrylamide gels prepared with varying amounts of bisacrylamide crosslinker

Bisacrylamide content (wt%)

Mc (g/mol)

Mesh size (nm)

1

5.5 (±0.1) × 103

14 (±0)

2

2.9 (±0.1) × 103

9.1 (±0.2)

Gels prepared with 5.2 M acrylamide, 210 mM MDEA, 35 mM VP, and 4 μM eosin isothiocyanate in water. 2 wt% bisacrylamide is 130 mM. Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm

Table 4

Estimated molecular weight between crosslinks (Mc) and mesh size in PEG gels prepared with varying amounts of PEGDA crosslinker

Diacrylate/monoacrylate (wt%/wt%)

Mc (g/mol)

Mesh size range (nm)a

0/22

1.2 (±0.1) × 104

5.6–19

1/21

7.9 (±0.7) × 103

4.4–14

11/11

2.2 (±0.1) × 103

2.0–6.5

22/0

2.4 × 102–1.5 × 103 b

2.0–5.0

Gels prepared with 210 mM MDEA, 35 mM VP, and 4 mM eosin–isothiocyanate in water. 22 wt% PEGDA is 420 mM, and 22 wt% PEGA is 650 mM. Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm

aThe low end of the range is obtained by assuming that the chain between crosslinks is mainly composed of carbon–carbon linkages. The high end of the range is obtained by assuming that the chain between crosslinks mostly consists of PEG-backbone

bMc is calculated with and without taking into account chain ends

Effect of monomer type

Monomer selection has been shown to have a significant effect on the thickness of polymer films generated from surface-mediated eosin initiation of polymerization (Avens et al. 2008). In addition to yielding films of differing thicknesses, monomer choice is also expected to impact how well NPs incorporate into the growing films, based on chemical interactions between the polymer and the NP surface, as well as the mechanical properties of these films. To determine how the two monomer formulations employed here affect the immobilization of fluorescent nanoparticles, surface-mediated polymerizations were carried out with both an acrylamide and PEGDA monomer formulation in the presence of 0.05 wt% 20 nm Dark red NPs. Not unexpectedly, the acrylamide formulation has a more than 20-fold greater thickness gain as compared to the PEGDA system (Fig. 7), which is consistent with previous reports on similar monomer systems (Avens et al. 2008). The overall gain is also larger for the acrylamide system though by less than 2-fold. Strikingly, the fluorescence gain of the PEGDA system is 10-fold higher than the acrylamide formulation, highlighting that the PEGDA formulation is much more efficient at trapping fluorescent NPs though its polymer films are significantly thinner. This difference is likely attributable to the smaller average mesh size of the PEGDA formulation (2–5 nm) as compared to the acrylamide formulation (9 nm) (Tables 2, 3), leading to more facile NP entrapment. In addition, the Nps are composed of a polystyrene core, surrounded by a PEG shell which is functionalized with carboxylate groups. Thus, it is reasonable to expect that the NPs are more compatible chemically with the PEGA/PEGDA environment, interacting more favorably with PEG chains in these polymers than with the relatively more hydrophobic polyacrylamide.
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Fig. 7

Normalized overall gain (solid squares), normalized fluorescence gain (open circles), and normalized thickness gain (open triangles) for polymer films generated by surface-mediated initiation with eosin using either an acrylamide monomer formulation (5.2 M acrylamide with 130 mM bisacrylamide) or a PEGDA formulation (420 mM PEGDA), each with 210 mM MDEA, 35 mM VP, and 0.05 wt% 20 nm carboxylate-functionalized Dark red NPs. Polymerizations were initiated by a 30 min exposure to 40 mW/cm2 light of wavelengths greater than 480 nm. For all three types of gain, the observed difference between the acrylamide and PEGDA monomer formulations is statistically significant (α = 0.05)

A summary of the posited critical factors that affect NP incorporation into films generated by surface-mediated initiation of polymerization are shown in Fig. 8. Smaller particle sizes are seen to incorporate more efficiently into the films, likely due to their ability to become entrapped in the heterogeneous monomer–polymer interface. Additionally, surface functionality of the NPs and monomer choice also affect the entrapment of fluorescent NPs through non-covalent and possibly also covalent interactions. Finally, it is hypothesized that the favorable influences of smaller mesh sizes and faster rates of mesh formation at the monomer–polymer interface contribute to the improved NP incorporation seen with higher crosslinker content. Knowledge of these factors enables selection of NP and monomer attributes that are most suitable for specific applications that rely on surface-mediated initiation of polymerization to immobilize fluorescent NPs.
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Fig. 8

Summary of the posited factors influencing NP incorporation into films generated by surface-mediated initiation of polymerization

Impact on sensitivity in a biodetection application

Fluorescent polymerization-based amplification (FPBA) has been demonstrated previously to be a sensitive array-based biodetection platform for signal amplification that enables a 100-fold improvement in sensitivity as compared to traditional fluorescence detection methods (Avens and Bowman 2010; Hansen et al. 2008b). FPBA relies on surface-mediated initiation of polymerization to entrap fluorescent NPs within PEGDA or polyacrylamide polymer films. Specifically, the assay employs DNA or protein probes that are coupled to eosin photoinitiators, thereby immobilizing eosin photoinitiators on the surface wherever the probe has bound its target. In the last step of the assay, the surface is contacted with monomer, MDEA coinitiator, and fluorescent NPs and exposed to visible light, generating a highly fluorescent film specifically in regions of the surface where target is present. To achieve the highest sensitivity for fluorescence detection, it is necessary to maximize the mass of fluorescent NPs immobilized per surface area in response to the eosin-initiated polymerization.

The FPBA array-based biodetection platform was used to assess the combined effect of selecting monomer and NP attributes that are more or less favorable for immobilization of fluorescent NPs to the surface. As a model system for biodetection, microarray slides were used that contain a range of biotin-labeled antibody surface densities. These slides were contacted with eosin-labeled streptavidin which specifically binds to biotin, thereby immobilizing eosin photoinitiators to the surface. The biotin–streptavidin system is a commonly used label-probe combination employed in many detection assays. Two different monomer formulations were compared: (1) an acrylamide monomer formulation containing 2 wt% crosslinker and 20 nm yellow/green NPs; and, (2) a formulation containing 1 wt% PEGDA, 21 wt% PEGA and 100 nm yellow/green NPs. Based on the aforementioned NP-incorporation studies, the first formulation is expected to generate significantly stronger amplified fluorescence signals and allow more sensitive detection due to the smaller NP diameter, the higher crosslinker content, and the ability of acrylamide to grow thicker films.

Indeed, the acrylamide formulation with 20 nm NPs is found to enable detection of as few as 0.16 biotin-labeled antibodies per square micron, or 40 zeptomole surface-bound target molecules, while the PEGA/PEGDA formulation with 100 nm NPs is approximately 200-fold less sensitive with a detection limit of 31 biotin-labeled antibodies per square micron (Fig. 9). A direct comparison of the magnitude of fluorescence intensity generated by the two formulations is not readily obtained from these data, as the signal generated from the acrylamide formulation has started to saturate at the antibody surface densities that are required to see significant signal from the PEGDA/PEGA system. Additionally, it should be noted that the threshold for significant signal (S/N > 3) was higher in the PEGA/PEGDA system because the non-specific polymerization background on these surfaces was higher and more variable. These results demonstrate that selection of the appropriate monomer and NP combination is critical for designing a biodetection platform that relies on entrapment of NPs by surface-mediated initiation of polymer film growth.
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Fig. 9

Two different monomer/NP formulations with different anticipated aptitudes for immobilizing fluorescent NPs are compared for their suitability in the fluorescent polymerization-based signal amplification platform for biodetection. Solid squares represent 5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, 35 mM VP, and 0.05 wt% 20 nm yellow/green NPs in water. Open circles represent 620 mM PEGA, 19 mM PEGDA, 210 mM MDEA, 35 mM VP, and 0.05 wt% 100 nm carboxylate-functionalized yellow/green NPs in water. Polymerizations were initiated by a 30 min exposure to 40 mW/cm2light of wavelengths greater than 480 nm. The limit of detection (S/N > 3) for each system is indicated by an asterisk

Impact on the ability to obtain localized staining of a small cellular feature

Recently, FPBA has been extended to fluorescent immunocytochemistry, enabling enhanced fluorescence and more sensitive detection compared to traditional fluorescent immunocytochemistry methods which employ direct fluorescent labeling of the antigenic or protein probe (Avens HJ et al. submitted to Journal of Histochemistry and Cytochemistry). In this format, cells fixed onto glass slides are probed with antibodies that specifically bind a target of interest within the cell. Eosin photoinitiators are either coupled to the primary or secondary antibody, or alternatively, eosin-labeled streptavidin is used in conjunction with biotin-labeled antibodies. In the last step of the assay, the cells are contacted with a monomer formulation containing fluorescent NPs and are exposed to light, yielding the formation of highly fluorescent films specifically on structures in the cell where the antigenic probe has bound its target protein. Unlike the microarray platform in which generation of thick films is acceptable, for a cell imaging application, the generation of excessively thick films is undesirable because it reduces the spatial resolution of the staining process. Therefore, it is expected that a PEGDA formulation would be preferable to an acrylamide formulation since the former is capable of immobilizing a higher density of fluorescent NPs into the polymer film. This hypothesis was tested by performing FPBA with each formulation to detect nuclear pore complex proteins (NPC) which are localized within the nuclear envelope. As shown in Fig. 10, the acrylamide formulation yields an unacceptably diffuse ring of staining around the nucleus that reaches nearly 10 microns thick in some regions. Conversely, the PEGDA formulation results in staining that is more spatially confined around the targeted features at the edge of the nucleus.
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Fig. 10

Comparison of using an acrylamide (a) versus a PEGDA (b) monomer formulation for FPBA-based staining of the nuclear pore complex of endothelial cells. The acrylamide monomer formulation comprises 5.2 M acrylamide, 130 mM bisacrylamide, 210 mM MDEA, 35 mM VP in water, and 0.05 wt% yellow/green fluorescent NPs in water. The PEGDA monomer formulation comprises 420 mM PEGDA, 210 mM MDEA, 35 mM VP, and 0.05 wt% yellow/green fluorescent NPs in water. Polymerizations were initiated by a 20 min exposure to 30 mW/cm2 light of wavelengths greater than 480 nm. The scale bar indicates 10 μm

These two biodetection examples demonstrate the importance of understanding the NP incorporation behavior and their impact on polymerization as each different application places distinct requirements on the polymerization and detection process. Here, the optimal NP–monomer combination is dramatically different depending on whether one requires maximum sensitivity or whether one desires maximum spatial resolution.

Conclusions

The results presented here highlight the fact that careful consideration of monomer and NP selection is essential to achieving the desired properties in applications that employ surface-mediated polymerizations to entrap NPs. In choosing a monomer formulation, higher crosslinker content tends to be more efficient at entrapping NPs, while consideration of the interactions between the polymer and the NP surface is also important. Choice of suitable NP attributes requires consideration of both NP size and surface chemistry, with smaller sizes tending to favor entrapment in polymer matrices. Although this study employed relatively low NP concentrations (0.05 wt%) such that light attenuation of the NPs would not excessively hinder photoinitiation, it is expected that the trends outlined here will be similar and perhaps even more pronounced with increased NP loading. It is additionally anticipated that these findings will be helpful in designing approaches for surface-mediated generation of polymeric nanocomposites composed of other polymers, polymerization types, and NP types.

Acknowledgments

This material is based upon work supported by National Institutes of Health R21 CA 127884, a National Science Foundation Graduate Research Fellowship to HJA, and NSF Research Experience for Undergraduates funding to KRV. Also, this work has been supported by the State of Colorado and the University of Colorado Technology Transfer Office.

Supplementary material

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© Springer Science+Business Media B.V. 2010