pubs.acs.org/Langmuir © 2010 American Chemical Society
DNA-Mediated Two-Dimensional Colloidal Crystallization above Different Attractive Surfaces Sabrina Jahn,† Nienke Geerts,‡ and Erika Eiser*,†,§ †
Cavendish Laboratory, University of Cambridge, J.J. Thomson Avenue, Cambridge CB3 0HE, United Kingdom, ‡Yale University, Malone Center, 55 Prospect Street, New Haven, Connecticut 06511, United States, and §BP Institute, Madingley Road, Cambridge CB3 0EZ, United Kingdom Received August 11, 2010. Revised Manuscript Received September 19, 2010 We explore the formation of “floating” two-dimensional colloidal crystals above weakly attractive surfaces that are either positively or negatively charged. In particular, we studied crystal formation above positively charged poly-Llysine-poly(ethylene glycol) surfaces with and without short single-stranded DNA and above negatively charged bovine albumin serum-streptavidin multilayers. Confocal microscopy revealed the evolution of crystals several micrometers above all three surfaces. Interestingly, the “flying height” of crystals was found to depend on the surface coating. All crystalline structures remained remarkably stable over weeks, even under high salt conditions. Neither lifting the crystals nor lowering them by means of buoyancy forces destroyed them.
1. Introduction The possibility to assemble and direct individual colloidal building blocks into highly ordered two- or three-dimensional structures has gained practical relevance in the context of the development of novel photonic1 and plasmonic2 materials. Since the 1980s, when “hard-sphere” colloidal systems were first prepared3 and colloidal self-assembly became a common model system for crystallization in statistical physics, various routes of colloidal crystal formation have been investigated by experimentalists and theoreticians. Colloidal crystals either can form spontaneously in solution4,5 or can be induced by convective,6,7 gravitational,8 and electrohydrodynamic forces.9,10 Recognitionmediated assembly, which allows for a high degree of control and tunability of particle-particle interactions, can be achieved by cross-linking particles with biomolecules.11-14 The specific shortrange interactions that these biomolecular linkers provide offer the prospect to create unique colloidal assemblies that are otherwise entropically unfavorable. In particular, DNA linkers have the advantage that their base-pairing is sequence-specific and *To whom correspondence should be addressed. E-mail: ee247@ cam.ac.uk. (1) Urban, J. J.; Talapin, D. V.; Shevchenko, E. V.; Kagan, C. R.; Murray, C. B. Nat. Mater. 2007, 6, 115–121. (2) Lee, J.; Hernandez, P.; Lee, J.; Govorov, A. O.; Kotov, N. A. Nat. Mater. 2007, 6, 291–295. (3) Pusey, P. N.; Vanmegen, W. Phys. Rev. Lett. 1987, 59, 2083–2086. (4) Gast, A. P.; Russel, W. B. Phys. Today 1998, 51, 24–30. (5) Pusey, P. N.; Vanmegen, W. Nature 1986, 320, 340–342. (6) Denkov, N. D.; Velev, O. D.; Kralchevsky, P. A.; Ivanov, I. B.; Yoshimura, H.; Nagayama, K. Nature 1993, 361, 26–26. (7) Micheletto, R.; Fukuda, H; Ohtsu, M. Langmuir 1995, 11, 3333–3336. (8) vanBlaaderen, A.; Ruel, R.; Wiltzius, P. Nature 1997, 385, 321–324. (9) Holgado, M.; Garcia-Santamaria, F.; Blanco, A.; Ibisate, M.; Cintas, A.; Miguez, H.; Serna, C. J.; Molpeceres, C.; Requena, J.; Mifsud, A.; Meseguer, F.; Lopez, C. Langmuir 1999, 15, 4701–4704. (10) Lin, K. H.; Crocker, J. C.; Prasad, V.; Schofield, A.; Weitz, D. A.; Lubensky, T. C.; Yodh, A. G. Phys. Rev. Lett. 2000, 85, 1770–1773. (11) Alivisatos, A. P.; Johnsson, K. P.; Peng, X.; Wilson, T. E.; Loweth, C. J.; Bruchez, M. P., Jr.; Schultz, P. G. Nature 1996, 382, 609–611. (12) Biancaniello, P. L.; Kim, A. J.; Crocker, J. C. Phys. Rev. Lett. 2005, 94, 058302-1–058302-4. (13) Katz, E.; Willner, I. Angew. Chem., Int. Ed. 2004, 6042–6108. (14) (a) Schmatko, T.; Bozorgui, B.; Geerts, N.; Frenkel, D.; Eiser, E.; Poon, W. C. K. Soft Matter 2007, 3, 703–706. (b) Geerts, N.; Eiser, E. Soft Matter 2010, 6, 4647–4660.
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thermoreversible. Unlike other biological “linker” molecules such as biotin and avidin, which bind together with high affinity, the attraction between complementary DNA strands can vary significantly depending on the oligonucleotide sequence and the solution conditions.15 DNA-directed colloidal assembly has focused primarily on the linkage of binary colloidal systems via DNA hybridization. Most of the studies observed the formation of disordered DNA-linked nano- or microparticle assemblies rather than crystalline structures, even where theoretical studies predicted the formation of crystals.14b,16,17 In 2008, the groups of Mirkin and Gang experimentally showed sequence-recognition driven 3D crystallization of short oligonucleotide-modified gold nanocolloids.18,19 We have recently reported a novel concept to assemble colloids grafted with very long λ-phage DNA into quasi-two-dimensional crystals above a weakly attractive poly-L-lysine-graft-poly(ethylene glycol) (PLL-g-PEG) flat surface at a very low colloidal volume fraction.20 Compared to other studies reported in the literature,21 the crystallization was not driven by the DNA hybridization between complementary ssDNA-coated colloids, but by the polymeric nature of the DNA with its extreme contour length. We proposed a two step crystallization process: First the colloids with the negatively charged DNA corona need to diffuse through the density matched solvent to reach and be adsorbed by the sticky support surface. Because the DNA arms have a gyration radius of almost 1 μm, the anchored colloids still have enough freedom to diffuse locally above the surface, giving neighboring colloids the time to overcome the weak steric DNA corona (second step). Once this steric repulsion is overcome, the (15) Milam, V. T.; Hiddessen, A. L.; Crocker, J. C.; Graves, D. J.; Hammer, D. A. Langmuir 2003, 19, 10317–10323. (16) Lukatsky, D. B.; Mulder, B. M.; Frenkel, D. J. Phys.: Condens. Matter 2006, 18, S567–S580. (17) Tkachenko, A. V. Phys. Rev. Lett. 2002, 89, 148303-1–148303-4. (18) Nykypanchuk, D.; Maye, M. M.; van der Lelie, D.; Gang, O. Nature 2008, 451, 549–552. (19) Park, S. Y.; Lytton-Jean, A. K. R.; Lee, B.; Weigand, S.; Schatz, G. C.; Mirkin, C. A. Nature 2008, 451, 553–556. (20) Geerts, N.; Eiser, E. Soft Matter 2010, 6, 664–669. (21) Kim, A. J.; Biancaniello, P. L.; Crocker, J. C. Langmuir 2006, 22, 1991– 2001.
Published on Web 10/06/2010
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Figure 1. (A) Schematic representation of a glass surface modified with the cationic copolymer PLL-g-PEG, (B) PLL-g-PEG-streptavidinssDNA, and (C) protein multilayers of BSA-streptavidin-ssDNA. The schemes are not drawn to scale; in all three cases, the thickness of the “sticky” layer was about 2-3 nm. For the evaluation of the surface coverage, 1 μm large colloids coated with complementary ssDNA, which are able to bind to the ssDNA on the surface, hybridized (B1, C1). Scale bars refer to 5 μm.
colloids will come close enough to feel the van der Waals interactions between them. In this two-dimensional (2D) crystalline arrangement, the DNA arms remain bound to the colloid but will form a dense polymer layer below and above the colloidal membrane. These DNA layers above and below the membrane then give rise to an additional attraction due to the DNA depletion between the neighboring colloids. Conversely, colloids grafted with short oligonucleotides did not show any periodic arrangement. In the present paper, we compare the influence of surfaces with different molecular properties to mediate DNAdriven 2D crystallization. We examined three types of surfaces: a negatively charged protein multilayer composed of bovine serum albumin (BSA)-streptavidin-ssDNA and PLL-g-PEGmodified glass surfaces with or without ssDNA. Moreover, we studied the influence of size polydispersity on crystal formation and the stability of the crystalline structures over time.
2. Experimental Section 2.1. Surface Preparation. For sample preparations and surface modifications of scenarios A-C (Figure 1; A-C), 96well plates with a glass bottom were used (Sensoplate, Greiner bio-one) as sample chambers. Prior to the actual surface modifications, all wells were incubated overnight in 10% hellmanex solution (Hellma, U.K.) and subsequently rinsed extensively with double-distilled H2O (ddH2O). In scenario A, a poly-Llysine-poly(ethylene glycol) polymer solution (0.5 mg/mL, 50 μL; PLL-g-PEG, Surface Solutions) was added to the wells for at least 3 h. After removing the PLL-g-PEG excess, 15 μL of 0.5% (w/v) λ-DNA cos-biotin coated colloids density matched in sucrose-Tris buffer (150 mg mL-1 sucrose, 100 mM Tris-HCl, pH 8) were added to the sample cell. In scenario B, poly-Llysine-poly(ethylene glycol)-biotin incubated for 3 h, the excess was removed, and streptavidin solution (5 μg/μL) was added and left for 15 min, rinsed with ddH2O before cos-biotin λ-DNA incubated in a sample chamber for 12 h. Before the samples were 16922 DOI: 10.1021/la103192q
added, the modified surface was rinsed again with ddH2O. For scenario C, biotin-bovine serum albumin (BSA) solution (2.5 μg/μL) was prepared in 50 μL of 20 mM acetic acid/80 mM sodium acetate buffer pH 4.7. Charge neutral BSA adsorbs to glass surfaces. A 30 min incubation of biotin-BSA solution was followed by a removal and washing step with ddH2O after which the treatment with streptavidin (1 μg/μL) in 50 μL of 20 mM acetic acid/80 mM sodium acetate buffer pH 5.2 and a final rinsing step followed. This procedure was repeated four times. 2.2. Preparation of λ-DNA-cos-Biotin. In the present study, λ-phage DNA (New England Biolabs) with a contour length of 16 μm (48 502 bps) was used. Due to the length of the DNA, special care was taken while handling the DNA to prevent any damage resulting from shear forces. First, the λ-DNA was diluted in 100 mM Tris-HCl pH 8 (25 μg μL-1) and biotin-ssDNA (sequence: 50 -AGGTCGCCGCCC-30 , called cos-2; 5 μL, 20 μM) was added. Subsequently, the λ-DNA cos-2-biotin solution was heated up to 65 °C for 30 min in order to linearize the plasmid. After linearization, each λ-phage DNA strand exhibits 12 base complementary single stranded overhangs at each end, cos-1 and cos-2. For hybridization of cos-2-biotin with the free cos-1 overhang of the λ-phage DNA, a very slow cooling process (1 °C 500 s-1) from 65 °C to room temperature was chosen. T4 DNA ligase (New England Biolabs) was used to ligate the λ-phage DNA backbone. Thereafter, the cos-2-biotinylated λ-phage DNA was washed four times using a Microcon YM100 membrane (Millipore) with 100 mM Tris-HCl pH 8 to remove the excess of cos-2-biotin and ligase. 2.3. Grafting Colloids with Biotinylated λ-DNA. Prior to grafting DNA onto the colloids, the average size of the polystyrene particles was determined using a Malvern Zetasizer Nano instrument (Malvern Instruments, U.K.). Subsequently, DNAcos-biotin coated colloids were prepared by incubating biotinylated cos-modified λ-phage DNA with streptavidin-modified red (Rhodamine-B) or green (Pyrromethene 486) fluorescent polystyrene colloids (1 μm, Microparticles GmbH) in Tris-HCl buffer (100 mM, pH 8). The DNA and the colloids reacted overnight and Langmuir 2010, 26(22), 16921–16927
Jahn et al. were tumbled continuously. Following this, pelleting and washing the samples five times removed the surplus of unconjugated DNA. Between washings, samples were furthermore once heated for 10 min at 50 °C and washed with NaOH solution (0.15 μM), to remove poorly bound DNA. A 0.5% w/v suspension was obtained through the subsequent dilution of the DNA-coated colloids in a fresh mixture of Tris-HCl buffer (100 mM, pH 8). To obtain a colloidal volume fraction of ∼4 10-4, 15 μL of the colloidal suspension was added to 200 μL sucrose-Tris buffer (150 mg/mL sucrose, 100 mM Tris-HCl pH 8) and incubated in a surface-modified sample chamber at room temperature (RT). The suspension was imaged 4 h after injection of the sample. 2.4. Testing the Surface Coverage. To test the surface coverage of the PLL-g-PEG-biotin-cos-1 and the BSA-streptavidin-cos-1 layers, fluorescent streptavidin labeled beads were coated with complementary biotinylated cos-2 DNA. For this, 5 μL of cos-2 DNA (20 μM) was incubated with 15 μL of colloidal suspension (1% w/v) in 100 μL 100 mM Tris-HCl pH 8 buffer overnight. The washing and dilution steps are equivalent to the ones for the λ-DNA-coated colloids.
2.5. Evaluation of the Stability of 2D Crystalline Structures. After formation, crystalline monolayers were treated with different concentrations of the monovalent salt KCl over days to weeks in order to evaluate the stability of the crystals under extreme conditions. Stock solutions of 2 M KCl in sucrose-Tris buffer were prepared. Appropriate volumes were carefully added to the sample chambers. The samples incubated for 24 h in 100 μM KCl, 1 mM, 10 mM; 7 days in 100 mM and up to 6 weeks in 1 M KCl, consecutively. Confocal image series were taken after each incubation period. Treated samples at 100 mM KCl were imaged 24 h after addition of the salt as well as 7 days later. Samples incubating in 1 M KCl were imaged 24 h, 21 days, and 6 weeks after the addition of salt. Control samples that were not treated with KCl were imaged over the same time span. In addition to the salt treatments, the influence of buoyancy forces on the stability of the crystals was analyzed. As the sample solution (100 mM TrisHCl pH 8 sucrose 150 mg/mL) was density matched to the density of the polystyrene particles (1.055 g cm-3), a solution with a higher concentration of sucrose (300 mg mL-3) was prepared. The density in the sample chamber was elevated to 1.08 g cm-3 and confocal images were taken every 30 s over 1 h. After 1 h, 100 mM Tris-HCl without sucrose was used to bring the solution back to its original density. Confocal imaging was performed after 24 and 72 h. 2.6. Contact Angle Goniometry. Advancing and receding contact angles on PLL-g-PEG coated glass surfaces were measured using a KSV Cam 200 goniometer (KSV Instruments, U.K.). For this purpose, sample chambers with removable cover units (CultureWell chambered coverglass for cell culture, Invitrogen) were used instead of regular 96-well plates. Surface preparation was accomplished as described earlier. The measurements were performed with 6 μL starting and 8 μL oscillating volume of deionized water at a velocity of 0.5 μL/s. 2.7. Confocal Imaging. Confocal imaging was carried out using an inverted Leica TCS SP 5 microscope. Oil immersion objectives with a magnification of 63 and 100 were used for the analysis of the colloidal suspensions. Fluorescence was excited at 486 nm for the green and at 560 nm for the red colloids. Emission was detected at 506 and 584 nm, respectively. For 3D reconstructions and z-overlays of the images, z-stacks with a step size of 0.3 μm were taken. The determination of the “flying height” of the crystalline monolayers was accomplished by measuring the distance in the z-direction from the surface up to the focal plane of the selected area of the crystal. The size of the area of which the height was determined was chosen to be 15 15 μm2. Depending on the size of the crystal, up to 10 areas per crystal were measured. Statistical analysis was performed on eight crystals per sample across six independent samples per surface preparation scenario. 2.8. Data Treatment and 3D Reconstruction. z-Stacks of the confocal images were background subtracted and converted Langmuir 2010, 26(22), 16921–16927
Article to 8-bit files using Image J. With the aim to obtain the xyzcoordinates of the imaged colloids, an “in house” developed code using IDL 6.3 was applied. Briefly, images were converted into arrays with each element representing 1 pixel. Based on the intensity of the particles, local maxima were determined and rendered by a circle related to the diameter of the particle. A centroid algorithm was used to identify the center of mass. The output was a list of xyz-coordinates. The absolute number of particles was extracted from the script after determining the coordinates. Using the xyz-coordinates of the colloids, images were reconstructed by means of the molecular visualization software VMD. To calculate the radial distribution function, g(r), an IDL routine based on the method of Grier and Crocker was employed.22
3. Results and Discussion 3.1. The Formation of Crystals above Different Surfaces. We investigated the crystallization of DNA-grafted colloids into two-dimensional membranes above distinct chemically modified glass surfaces. Accordingly, fluorescent polystyrene streptavidin labeled particles (1 μm) were grafted with biotinylated λ-DNA (contour length 16 μm). As reported in previous studies, approximately 10 chains of λ-DNA attach to a micrometer-sized colloid from solution,14 forming a homogeneous monolayer of DNA coils, terminated with a cos-overhang. This steric polyelectrolyte layer prevented colloidal aggregation in the bulk. All experiments were performed at a colloidal volume fraction of 410-4. Figure 1 shows schematically the three different types of surface modifications we used, which were accomplished by the adsorption of (A) the cationic PLL-g-PEG copolymer, (B) PLL-gPEG-biotin-streptavidin-biotinylated-ssDNA, and (C) multiple BSA-biotin-streptavidin layers with biotin-ssDNA. The surface coverage in scenario B and C (Figure 1) was tested by incubating the modified surfaces with colloids coated with short complementary ssDNA (Figure 1B1, C1). The high number of uniformly distributed and hybridized colloids confirms the efficiency of the surface coating. A control sample composed of colloids without DNA shows a dramatically lower number of particles due to the surface passivation by means of PEG (data not shown). In order to exclude sedimentation effects, all experiments were prepared in a density-matched environment. We found that all three surfaces mediated λ-DNA-driven twodimensional crystallization (Figure 2). Their formation occurs up to several micrometers above the modified surfaces (Figure 2A1, B1, C1). No crystals were present in the bulk of the solution. The nature of the packed colloidal monolayers was unambiguously crystalline. Characteristic crystalline facets as well as grain boundaries and point defects in larger crystalline structures were found. Sizes of the 2D monolayers ranged from 15 to 6000 colloids per crystal. Up to 18 independent 2D closely packed colloidal monolayers per sample were identified in this study. The mean surface area covered by crystalline structures was analyzed to be 8.2%, 5.9%, and 3.4% for PLL-g-PEG, PLL-g-PEG-strepatvidin-ssDNA, and BSA-strepatvidin-ssDNA multilayers, respectively, across eight independent samples per surface preparation scenario. In scenario A, PLL-g-PEG copolymer layers adsorbed to the glass surface via electrostatic interactions. The quality and homogeneity of the PLL-g-PEG films was evaluated by water contact goniometry. Advancing (θa) and receding (θr) contact angles were measured to be 25.9° ( 1° and 20.3° ( 1°, respectively. Low hysteresis values confirmed the presence of a homogeneous PLL-g-PEG film. Similar values of PLL-g-PEG films on silica (22) Crocker, J. C.; Grier, D. G. J. Colloid Interface Sci. 1996, 179, 298–310.
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Figure 2. Colloidal crystallization above different surfaces. Confocal images of a 2D crystalline structure above (A) PLL-g-PEG, (B) PLL-gPEG-streptavidin-ssDNA, and (C) BSA-strepatvidin-ssDNA surfaces. On the right, histograms of the average flying height of the crystals above (A1) PLL-g-PEG, (B1) PLL-g-PEG-streptavidin-ssDNA, and (C1) BSA-streptavidin-ssDNA surfaces are presented. Scale bars refer to 10 μm.
have also been reported in other studies.23 Cleaned glass surfaces with θa = 16.8° ( 2° and θr = 14.3° ( 1° served as controls. The assembly of a comblike PLL-g-PEG layer is thought to occur at the surface on which positively charged primary amine groups of the PLL bind to the negatively charged glass surface, whereas hydrophilic and uncharged PEG side chains are exposed to the solution phase.24 In this scenario, anchoring of the DNA-coated colloids to the surface through Watson-Crick base pairing is not possible. Hence, adsorption of the grafted DNA to the PLL-g-PEG (23) Saravia, V.; K€upc€u, S.; Nolte, M.; Huber, C.; Pum, D.; Fery, A.; Sleytr, U. B.; Toca-Herrera, J. L. J. Biotechnol. 2007, 130, 247–252. (24) Kenausis, G. L.; V€or€os, J.; Elbert, D. L.; Huang, N.; Hofer, R.; RuizTaylor, L.; Textor, M.; Hubbell, J. A.; Spencer, N. D. J. Phys. Chem. B 2000, 104, 3298–3309.
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surface is likely to be governed by the Coulomb attraction between the highly negatively charged λ-DNA corona and the positively charged poly-L-lysine. These forces are strongly dependent on the ionic strength of the buffer and may compete with adverse interactions between the colloidal DNA corona and the PEG side chains. Schlapak et al. argued that the deposition of DNA onto the ∼2 nm thick PLL-PEG films may result in a deposition-related dissociation of water molecules from the hydration shells that surround the PLL-g-PEG, and DNA.25 Thus, the adsorption of λ-DNA will only be favorable if the electrostatic energy dominates the PEG-DNA interactions. (25) Schlapak, R.; Armitage, D.; Saucedo-Zeni, N.; Chrzanowski, W.; Hohage, M.; Caruana, D.; Howorka, S. Soft Matter 2009, 5, 613–621.
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Indeed, Schlapak et al. found long-range electrostatically driven DNA adsorption to a PLL-g-PEG monolayer only above 100 mM NaCl concentrations.25 Hence, the Coulomb attraction of the λ-DNA-coated colloids to the surface plays an important role in crystal formation. Owing to a radius of gyration of ∼0.8 μm, the λ-DNA remains mobile enough to continue diffusing locally above the surface. These surface interactions give approaching colloids sufficient time to overcome the steric polymer layer surrounding the colloids, and to be arrested in an attractive secondary van der Waals minimum. As reported earlier, it remains difficult to estimate the strength and location of this secondary van der Waals minimum. At 1 nm separation between the 1 μm large bare polystyrene beads suspended in water at room temperature, the van der Waals attraction can be estimated to be about -300kBT, using the standard expression for sphere-sphere interactions and a Hamaker constant A = 9.5 10-21 J K-1. The zeta potential of the neutravidin-coated PS colloids in Tris-HCl buffer at pH 8 was about -20 mV, giving rise to a Coulomb repulsion of about 80kBT. However, the measure for the steric repulsion caused by the grafted DNA remains elusive. Furthermore, calculations by Bhatia and Russel suggest that colloids that are grafted with polymers of similar size exhibit an attraction due to many-body interactions, which are not taken into account in common van der Waals force calculations that only consider twobody interactions.26 In scenario B, PLL was grafted to PEG chains, half of which were terminated with biotin. To create a multiple component surface following the adsorption of PLL-g-PEG-biotin, layers of streptavidin and biotinylated ssDNA (here cos-1) were added consecutively (Figure 1B). With the addition of short DNA strands, base-pairing interactions between the single-stranded DNA overhangs of the λ-DNA (cos-2) and the surface became possible. As the streptavidin has four binding sites with one blocked by the PLL-g-PEG biotin, up to three can be occupied by ssDNA, yielding a dense brush of ssDNA. Although we worked in a saturated regime, the overall charge per PLL-g-PEGbiotin-streptavidin-ssDNA molecule is hard to estimate. To understand the influence of the ssDNA brush attached to the surface on crystal formation, we performed separate experiments using λ-DNA-coated colloids with complementary cos-ends and noncomplementary ones. Interestingly, we were able to observe colloidal crystallization in both regimes. Thus, hybridization plays a minor role in the present scenario, supporting our hypothesis that electrostatics between the PLL-g-PEG-biotin and the λ-DNA are the driving forces. To further test our hypothesis that a positively charged polymer other than PLL-g-PEG is also able to mediate DNA-driven colloidal 2D-crystallization, we conducted preliminary experiments with the cationic polyelectrolyte poly(diallyldimethylammonium chloride) (pDADMAC). We found crystalline structures of sizes up to 60 colloids above pDADMAC-modified surfaces (data not shown). The limitation in size might be due to the molecular architecture of the polyelectrolyte or its inefficient absorption onto the glass surface. Crocker and co-workers studied the crystallization behavior of DNAgrafted micrometer-sized colloids with different surface chemistries induced by depletion forces.21 They compared carboxylated, neutravidin-modified, and PEGylated DNA-grafted particles with each other. Only PEGylated colloids formed crystals. The distinct polymeric comblike architecture and the DNA-PEG chain interactions present in scenarios A and B may thus influence the time scale and may be in favor to form larger crystalline structures compared to the surfaces modified with pDADMAC. (26) Bhatia, S. R.; Russel, W. B. Macromolecules 2000, 33, 5713–5730.
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In scenario C, we prepared protein-based multilayers of biotinylated BSA and streptavidin linkers with terminal biotinylated ssDNA that was complementary to the sticky overhangs of our λ-DNA colloids. These BSA-streptavidin-ssDNA layres were about 2 nm thick. After surface preparation, the experiments of crystal formation were performed at pH 8 at which the protein layers were negatively charged. Hence, we studied a negatively charged surface compared to the systems described above. Also under these conditions we observed 2D crystallization. Protein layers terminated with noncomplementary ssDNA did not show any crystalline or amorphous aggregation of colloids. This suggests that the hybridization energy is the crucial driving force for crystallization. We calculated the hybridization energy between complementary single-stranded cos-overhangs to be -25kBT based on a method of Santa-Lucia.27 3.2. Comparison of the Flying Heights. Crystalline monolayers were analyzed in terms of their “flying height” above the different surfaces (Figure 2). Figure 2 (A1, B1, C1) shows a relatively broad distribution of flying heights for all surfaces. Although in all three cases the average flying height is larger than the radius of gyration of a λ-phage DNA coils, the colloidal membranes formed above PLL-g-PEG surfaces are closer to the surface than the ones observed in scenarios B and C. Using a Gaussian fit, we estimate the average flying height above PLL-gPEG surfaces to be ∼1.7 μm. Crystals above PLL-g-PEG-cos-1 and BSA-streptavidin-cos-1 had an average flying height of ∼3.1 and ∼3.3 μm, respectively. This suggests that λ-DNA, which is grafted to the colloids in a monolayer of thickness comparable to its radius of gyration, undergoes a transition from a coil to a more stretched-brush configuration, once it is bound to the surface. Under consideration of the grafting density of the crystalline sheets, stretching by a factor of 2-3 is expected.20 The larger flying height, and hence stronger stretching, in the case of the binding through hybridization may be understood by comparing the binding energies. We assume that the binding between the negatively charged DNA and the oppositely charged PLL-g-PEG layer is weaker than the 25kBT hybridization energy between cos-1 and cos-2. This energy can oppose the stretching of the DNA layer trapped between the colloids and the surface that arises when the colloids come close to each other in the crystalline layers, hence generating an increased osmotic pressure. 3.3. Polydispersity. We also studied the effect of polydispersity of the colloids. Zetasizer measurements showed a size distribution of 1.018 ( 0.04 μm for particles which were able to crystallize. Particles with sizes of 0.925 ( 0.12 μm formed amorphous colloidal monolayers (data not shown). Size polydispersity strongly influences the stability, nucleation, and growth of crystals and thus the location of the freezing curve in 2D and 3D suspensions of hard spheres. Computer simulations predicted a suppression of 3D crystal formation for polydispersities (σ) > 5.7%.28,29 Pronk and Frenkel found that 2D crystals are able to support a higher degree of polydispersity than 3D crystals.30 We have recently reported an experimental study in which binary λ-DNA-coated colloids (nominal 1 μm) with a size ratio of 0.9 fractionate upon crystallization to form effectively two pure 2D crystals of larger and smaller spheres.31 Although the DNA grafted colloids in ref 31 and in the present study do not entirely (27) (28) (29) (30) (31) (5pp).
Santa-Lucia, J. Proc. Nat. Acad. Sci. U.S.A. 1998, 95, 1460–1465. Auer, S.; Frenkel, D. Nature 2001, 413, 711–713. Kofke, D. A.; Bolhuis, P. G. Phys. Rev. E 1999, 59, 618–622. Pronk, S.; Frenkel, D. Phys. Rev. E 2004, 69, 066123-1–066123-7. Geerts, N.; Jahn, S.; Eiser, E. J. Phys.: Condens. Matter 2010, 22, 104111
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Figure 3. Influence of different KCl concentrations as a function of the number of colloids indicating the stability of the crystalline structure. After incubating for 6 weeks in 1 M KCl, the number of particles decreased only 1.9% compared to an untreated control sample.
Figure 4. Pair correlation function of a crystal that formed above a PLL-g-PEG surface. The crystal stability was observed over time under the influence of KCl treatment (see legend).
act as hard spheres, the colloidal hard-sphere radius is governing the periodic arrangement. Thus, the underlying mechanism of size fractionation and crystal formation suppression ought to be the same as in a hard sphere regime. Our results agree well with observations made in simulations for hard spheres: colloids with a standard deviation of 4.0% crystallize, while batches with polydispersities of 12.9% do not. 3.4. Salt Treatments and the Influence of Buoyancy Forces. With the aim to evaluate the stability of the colloidal crystals under extreme conditions, crystalline monolayers were treated with increasing concentrations of KCl up to 1 M while maintaining density matching between colloids and solvent, over 6 weeks. Confocal imaging was accomplished after different aging times. Subsequent 3D reconstructions of the image series were performed, and the number of particles and the radial distribution function were analyzed. We found that crystals that were kept in 1 M KCl solution over 6 weeks showed a decrease in the total number of particles of only 1.9% compared to a control sample not treated with salt (Figure 3), indicating a high stability. Moreover, the lattice spacings between colloids were not altered by the addition of KCl once the crystalline structures formed (Figure 4). This finding strengthens our argument that the crystals are in an attractive secondary van der Waals minimum that is reinforced by depletion attractions caused by the surrounding DNA cloud. Furthermore, the stability of the crystalline sheets was investigated by lifting them off the surface using buoyancy forces. In a 16926 DOI: 10.1021/la103192q
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Figure 5. Confocal image of colloidal crystalline structures formed in a capillary above a PLL-g-PEG-modified surface. A z-projection of the image stack is shown. The scale bar refers to 10 μm.
typical crystallization experiment, the DNA-coated polystyrene colloids were density matched with 100 mM Tris-HCl-sucrose solution pH 8 at 1.055 g cm-3. Solutions with higher densities due to an increased amount of sucrose were prepared and used to lift the crystals. The final density in these experiments was calculated and measured to be 1.080 g cm-3. The crystals were monitored every 30 s over 1 h. After lifting, we used a defined volume of 100 mM Tris buffer pH 8 to drive the crystalline structures back to the original density state. We observed that neither lifting nor bringing down of the crystalline sheets destroyed them or led to obvious structural changes. Also the geometry of the sample chamber did not affect the formation of colloidal 2D crystalline monolayers. Instead of a 96-well chamber, we used an X-ray glass capillary with an inner diameter of 1 mm and coated it with PLL-g-PEG. Density-matched DNA-coated colloids were then injected and incubated for several hours. We observed massive 2D crystalline sheets growing along the capillary wall (Figure 5).
4. Conclusions We have shown that micrometer-sized λ-DNA-coated colloids are able to form two-dimensional crystalline structures above weakly attractive surfaces with different molecular properties. Glass surfaces modified with the cationic copolymer PLL-g-PEG, PLL-g-PEG-streptavidin-ssDNA were compared with a negatively charged protein multilayer composed of biotin-BSAstreptavidin-ssDNA. PLL-g-PEG and PLL-g-PEG-ssDNA with complementary or non-cDNA-treated surfaces have shown that the hybridization between colloid-bound DNA single-strand overhangs and oligonucleotide-modified surfaces is not necessary for the formation of crystals. Whereas we observed that DNAcoated colloids above a negatively charged protein layer form crystalline structures only in the presence of complementary ssDNA attached to the protein layer. Here, the hybridization energy is essential. The “flying height” was observed to differ depending on the surface coating: the higher binding energy between the ssDNA strands causes a denser DNA layer between the crystal and the surface, leading to a coil-brush transition. Crystalline structures above PLL-g-PEG surfaces were on average found to occur closer to the surface. An important factor in the formation of the crystalline structures is size polydispersity. DNA-coated colloids with a standard deviation (SD) in size of Langmuir 2010, 26(22), 16921–16927
Jahn et al.
4% crystallized, while colloids with a SD of 12% formed flat amorphous aggregates. Additionally, we found that the formation of the crystals does not depend on the geometry of the sample chamber, as crystals were also able to form in a capillary. With respect to the stability, once formed the crystalline structures remained stable in high salt concentrations over weeks as well as after lifting the structures off the surface by means of buoyancy forces.
Langmuir 2010, 26(22), 16921–16927
Article
Acknowledgment. We thank M. Dogterom for providing laboratory facilities and E. Kumacheva for helpful discussions. Thanks to Nikolas Jahn as well, who performed many salt studies. This work is part of the research program of the Stichting voor Fundamenteel Onderzoek der Materie (FOM), which is supported by the Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO), the Cavendish Laboratory, and the Mott environmental fund.
DOI: 10.1021/la103192q
16927
