Hybridization and Immobilization of Long ds-DNAs on Polystyrene

Here, we develop a system where long double-stranded DNAs (ds-DNAs) are ... hybridization as a unique method to attach long ds-DNA chains in solution ...
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Hybridization and Immobilization of Long ds-DNAs on Polystyrene Microspheres Debalina Ghosh,† Nicolas Faure,‡ Sharmistha Kundu,† Francis Rondelez,‡ and Dipankar Chatterji*,† Molecular Biophysics Unit, Indian Institute of Science, Bangalore 560012, India, and Laboratoire Physico Chimie Curie (PCC), UMR CNRS 168, Institut Curie - Section de Recherche, 11, rue Pierre et Marie Curie, 75231 Paris Cedex 05, France Received February 5, 2003. In Final Form: May 5, 2003 Here, we develop a system where long double-stranded DNAs (ds-DNAs) are immobilized on the surface of a polystyrene (PS) microsphere. A simple synthetic strategy is adopted in order to achieve this goal in which a single DNA chain is anchored by one of its extremities to a latex (PS) microsphere. We chose hybridization as a unique method to attach long ds-DNA chains in solution with oligonucleotides grafted on modified aminated polystyrene microspheres. The DNAs chosen were of various sizes and sources: T7A1 DNA (4.48 kilobase pair (kbp)), a plasmid DNA; and ∆DIIIT7 DNA (39.34 kbp), a mutant of bacteriophage T7 DNA. The temperature dependence of the kinetics of hybridization of T7A1 DNA (4.48 kbp) in solution with an appropriate oligonucleotide (20-mer sequence) grafted on modified aminated polystyrene microspheres yielded a value of activation energy of ∼5.3 kcal/mol, consistent with a nondiffusion-controlled mechanism.

Introduction Immobilized DNA on a solid support has several technological and biological advantages. This system is used as a tool in medical, pharmaceutical, and diagnostics applications.1,2 It is also important to a wide range of research areas including work on nanoparticles,3 DNA chip technologies,4-6 DNA computing,5,6 and biosensor arrays.1,7 Some of the common supports used for the immobilization of DNA are latex particles, glass surfaces, silicon wafers, and gold nanoparticles. However, the need for reproducible stable surfaces has placed increasing interest on preparation of more DNA-modified surfaces. Latex particles are receiving considerable attention in the biomedical field because they can serve as a support for the immobilization of biomolecules through physical adsorption or covalent binding.8-12 Attaching doublestranded DNA (ds-DNA) to latex particles allows DNA manipulation as in optical tweezers13 or magnetic tweezers14 and direct observation of the biological processes * To whom correspondence should be addressed. Phone: +91-80-394 2836. Fax: +91-80-360 0535. E-mail: dipankar@ mbu.iisc.ernet.in. † Indian Institute of Science. ‡ Institut Curie - Section de Recherche. (1) Southern, E.; Mir, K.; Shchepinov, M. Nat. Genet. 1999, 21, 5. (2) Fodor, S. P. A. Science 1997, 277, 393. (3) Storhoff, J. J.; Mirkin, C. A. Chem. Rev. 1999, 99, 1849. (4) Fotin, A. V.; Drobyshev, A. L.; Proudnikov, D. Y.; Perov, A. N.; Mirzabekov, A. D. Nucleic Acids Res. 1998, 26, 1515. (5) Wang, L. M.; Liu, Q. H.; Corn, R. M.; Condon, A. E.; Smith, L. M. J. Am. Chem. Soc. 2000, 122, 7435. (6) Liu, Q.; Wang, L.; Frutos, A. G.; Condon, A. E.; Corn, R. M.; Smith, L. M. Nature 2000, 403, 175. (7) Mir, K. U.; Southern, E. M. Nat. Biotechnol. 1999, 17, 788. (8) Kremsky, J. N.; Wooters, J. L.; Dougherty, J. P.; Meyers, R. E.; Collins, M.; Brown, E. L. Nucleic Acids Res. 1987, 15, 2891. (9) Miller, C. A.; Patterson, W. L.; Johnson, P. K.; Swartzell, C. T.; Wogoman, F.; Albarella, J. P.; Carrico, R. J. Clin. Microbiol. 1988, 26, 1271. (10) Kawaguchi, H.; Asai, A.; Ohtsuka, Y.; Watanabe, H.; Wada, T.; Handa, H. Nucleic Acids Res. 1989, 17, 6229. (11) Arshady, R. Biomaterials 1993, 14, 5. (12) Imai, T.; Sumi, Y.; Hatakeyama, M.; Fujimoto, K.; Kawaguchi, H.; Hayashida, N.; Shiozaki, K.; Terada, K.; Yajima, H. J. Colloid Interface Sci. 1996, 177, 245.

occurring along the DNA chain.15 The other common utilities for these kinds of systems are reusability of a matrix-bound DNA for its biological activity, floating them as a monolayer to form a condensed system and performing biological reactions on such a condensed system, or manipulating their size by various enzymes without much difficulty as they are in a heterogeneous phase. ds-DNA is generally attached to latex particles by covalent grafting. In this method, the DNA end is modified to attach a particular chemical group like biotin or -SH that can react with a group or a small oligomer on the latex particle. Hybridization is also a method of choice, which has the distinct advantage of not requiring DNA modification. So far, this method has been applied only on short single-stranded oligomers16 but mostly on nonlatex particle surfaces,17-19 in the bulk phase (solution),20,21 or at the air-water interface.22 Hybridization of long dsDNA to an oligomer in solution is the most commonly used technique today because of its wide application in the polymerase chain reaction.23 However, it seems less practical when either one of the hybridizing partners is immobilized. Thus, when polynucleotides are used as ligands for the beads, the capture reaction rates are slow16,24,25 as observed by various techniques such as (13) Wang, M. D.; Schnitzer, M. J.; Yin, H.; Landick, R.; Gelles, J.; Block, S. M. Science 1998, 282, 902. (14) Maier, B.; Bensimon, D.; Croquette, V. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 12002. (15) Bianco, P. R.; Brewer, L. R.; Corzett, M.; Balhorn, R.; Yeh, Y.; Kowalczykowski, S. C.; Baskin, R. J. Nature 2001, 409, 374. (16) Wolf, S. F.; Haines, L.; Fisch, J.; Kremsky, J. N.; Dougherty, J. P.; Jacobs, K. Nucleic Acids Res. 1987, 15, 2911. (17) Georgiadis, R.; Peterlinz, K. P.; Peterson, A. W. J. Am. Chem. Soc. 2000, 122, 3166. (18) Peterson, A. W.; Heaton, R. J.; Georgiadis, R. J. Am. Chem. Soc. 2000, 122, 7837. (19) Lin, Z.; Strother, T.; Cai, W.; Cao, X.; Smith, L. M.; Hamers, R. J. Langmuir 2002, 18, 788. (20) Podder, S. K. Eur. J. Biochem. 1971, 22, 467. (21) Kuwahara, M.; Arimitsu, M.; Shigeyasu, M.; Saeki, N.; Sisido, M. J. Am. Chem. Soc. 2001, 123, 4653. (22) Shastry, M.; Ramakrishnan, V.; Pattarkine, M.; Gole, A.; Ganesh, K. N. Langmuir 2000, 16, 9142. (23) Mullis, K.; Faloona, F.; Scharf, S.; Saiki, R.; Horn, G.; Erlich, H. Cold Spring Harbor Symp. Quant. Biol. 1986, 51, 263.

10.1021/la0341963 CCC: $25.00 © 2003 American Chemical Society Published on Web 06/10/2003

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Figure 1. Sequence of reactions followed for grafting a DNA to a polystyrene microsphere.

surface plasmon resonance,26,17 fluorescence,4 or other methods.27 Considering the importance of grafting a long double-stranded DNA chain on a solid matrix like latex beads, we decided to work out the chemistry of attaching a small oligomer of DNA to the beads and subsequently hybridizing a long DNA chain with a complementary sequence at one end to this oligomer. Our interest was to quantitate the barrier of hybridization due to the presence of the heterogeneous phase. Perhaps one of the most exciting possibilities, yet unexplored, is the regulated uniform attachment of a solid matrix at the extremities of the long DNA chain with minimal entanglement or wrapping around the matrix. Materials and Methods 1. Preparation of Phenylene Mono-isothiocyanate (MITC)-Polystyrene (PS) Microspheres for Oligomer Attachment. The latex particles used for the experiments were monodisperse aminated PS microspheres with a diameter of 0.31 µm. These were obtained from Bangs Laboratories Inc., Fishers, IN, in 5% weight per volume solution. Before use, they were washed extensively in Nanopure MilliQ water (18.2 MΩ resistance) by the described procedure28 to remove soluble surfactants or chemicals absorbing at 260 nm and interfering in the quantitation of primer or DNA attachment. Usually, most uniform polystyrene microspheres are made by emulsion polymerization using surfactants. These surfactants are anionic in nature and may get adsorbed on the microsphere surface and can interfere with the coupling chemistry. Thus, repeated washing is an important step to ensure a clean coupling surface. An alternative protocol for washing was also used in which the beads were made surfactant/chemical free by washing with methanol and then water. A final suspension of 4-5% solids (w/v) was used for all attachments. Optical microscopic observations at 100× magnification showed evidence of clustering among the beads (an aggregate of greater than a thousand beads) after washing, but they were redispersed easily by sonication. All solutions were prepared and all reactions were done in Nanopure MilliQ water unless otherwise specified. Upon sonication, in the further modification steps, many single beads and a maximum of 2-10 bead aggregates were observed. (24) Flavell, R. A.; Birefelder, E. J.; Sanders, J. P. M.; Borst, P. Eur. J. Biochem. 1974, 47, 535. (25) Bunemann, H. Nucleic Acids Res. 1982, 10, 7181. (26) Peterlinz, K. A.; Georgiadis, R.; Herne, T. M.; Tarlov, M. J. J. Am. Chem. Soc. 1997, 119, 3402. (27) Okahata, Y.; Kawase, M.; Niikura, K.; Ohtake, F.; Furusawa, H.; Ebara, Y. Anal. Chem. 1998, 70, 1288. (28) Bangs, L. B. Technote No. 201: The Latex Course; Bangs Laboratories, Inc.: Fishers, IN, 1999.

The strategy used for the attachment of the oligonucleotides to the latex particle and the subsequent hybridization of the long ds-DNA chains to the bound oligonucleotides is shown in Figure 1. DITC (1,4-phenylene di-isothiocyanate) was obtained from Fluka, USA, and was used as a homodifunctional linker. One of the thiocyanate groups can react covalently with the surface amine groups, whereas the other remains free and available for a further reaction. A 677 µL portion of a 5% aminated bead solution (34 µg, corresponding to 2.07 × 1012 beads, containing a total of 7.5 × 1017 amine groups) was mixed with 0.007 g of DITC (2.2 × 1019 molecules) at room temperature for 2 h. Since the DITC to amine group ratio is large, typically 29:1, we can assume that essentially all amine groups present at the surface of the beads reacted with DITC. The modified beads were then washed to remove unattached DITC. Washing was done by centrifuging the beads at 8000g (10 000 rpm); the supernatant was discarded, and precipitated beads were resuspended in water to make the total volume 500 µL. Hereafter, we refer to these modified polystyrene microspheres as MITC-beads (Figure 1). 2. Attachment of the Primers to the MITC-PS Microspheres. Three primers, named P1, P2, and P3, respectively (Figure 2), were custom-prepared by Microsynth GmBH, Switzerland. They are all 20-mers but with different sequences. Primers P1 and P2 have a sequence complementary to position -80 to -100 of the T7A1 DNA or ∆DIIIT7 DNA, respectively. ∆DIIIT7 DNA is a derivative of T7 DNA, containing only one (T7A1) promoter specific for Escherichia coli RNA polymerase.29 P3 is identical to P1 except for a single base mismatch in the 10th position, where an A has replaced a C. The -105 to +14 sequences of T7A1 DNA and ∆DIIIT7 DNA are also shown in Figure 2. The position +1 represents the base from which the transcription starts. The promoter sequence is between positions -49 and +14 and is displayed in bold italics. The 5′ ends of all three primers were also modified with an amino group to allow their tethering to the free thiocyanate group of the MITC beads. The coupling reaction was performed as done by others too.30 A 500 µL portion of a 5% aqueous solution containing 2.07 × 1012 molecules (3.44 pmol) of MITC-beads was mixed with 2 µL (1.0 nmol or 6.0 × 1014 molecules) of the desired primer solution in TE buffer (10 mM Tris-HCl, 1 mM EDTA) at pH 9.0. Under such conditions, the ratio of thiocyanate groups on the MITC-beads to primer molecules is very large, typically 1000:1, and this ensures that all primer molecules would react. On the other hand, the ratio of primer molecules to MITC-beads is also sufficiently large, 250-300:1, so a significant number of primers will be attached to each bead. The reaction was allowed to proceed at room temperature for 6 h. The beads were then washed repeatedly by centrifugation, as above, until no more UV absorption at 260 nm due to unattached (29) Dunn, J. J.; Studier, F. W. J. Mol. Biol. 1983, 166, 477. (30) Elaissari, A.; Chevalier, Y.; Ganachaud, F.; Delair, T.; Pichot, C. Langmuir 2000, 16, 1261.

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Figure 2. (a) Sequences of the primers, P1, P2, or P3; (b) partial sequence of the pBR322 plasmid in which the A1 promoter is inserted to obtain the pAR1435 plasmid; (c) the partial sequences of T7A1 DNA and ∆DIIIT7 DNA showing the T7A1 promoter sequence (in bold italics) and the region of overlap to the 20 bp oligonucleotides P1 or P2, respectively. primer molecules could be detected in the supernatant. The primer-modified beads were finally resuspended in 500 µL of water. In the following, we refer to these modified polystyrene microspheres as P-beads, P being P1, P2, or P3 depending on the primer attached. 3. Preparation of T7A1 DNA and ∆DIIIT7 DNA. For the attachment to the beads, we employed two kinds of DNA here, a small linearized plasmid DNA (pAR1435, 4482 base pair (bp)) or a long double-stranded ∆DIIIT7 DNA (39 336 bp).30 T7, a naturally occurring bacteriophage, has three strong promoters for E. coli RNA polymerase: A1, A2, and A3. It is a long doublestranded DNA (∼40 kilobase pair (kbp)). pAR1435 was prepared by subcloning only the T7A1 promoter in a small E. coli plasmid, and ∆DIIIT7 DNA keeps all the sequences of T7DNA except the A2 and A3 promoters. Thus, both of these DNAs drive RNA synthesis with E. coli RNA polymerase exclusively from the A1 promoter. Figure 2 shows a schematic diagram of T7A1 in pBR322 plasmid generating pAR1435. Henceforth we shall refer to this as T7A1 DNA. It was prepared in supercoiled form according to the standard protocol of Sambrook et al.31 and linearized by the EcoRV restriction enzyme that has a unique site in the plasmid. The DNA was then electrophoresed at 100 V in a 0.8% agarose gel and visualized using ethidium bromide to check for completion of the digestion reaction. ∆DIIIT7 DNA was prepared using a protocol adapted from Nierman and Chamberlin.32 Both T7A1 DNA and ∆DIIIT7 DNA are double stranded, and their respective molecular weights are 2.958 × 106 and 25.962 × 106 Da. 4. Hybridization Reaction of T7A1 DNA and ∆DIIIT7 DNA to the Primer-Modified Polystyrene Spheres. Linearized T7A1 DNA (10 µg, 3.38 pmol, 2.03 × 1012 molecules) was dissolved in 500 µL of annealing buffer (2 mM Tris-HCl, pH 7.4, 5 mM NaCl, 0.2 mM MgCl2) in an eppendorf tube. The solution (31) Sambrook, J.; Fritsch, E. F.; Maniatis, T. Molecular Cloning: A Laboratory Manual; Cold Spring Harbor Laboratory Press: Cold Spring Harbor, New York, 1989. (32) Nierman, W. C.; Chamberlin, M. J. J. Biol. Chem. 1979, 254, 7921.

was boiled for 2 min to denature the ds-DNA and quickly frozen by dipping the tube in an ice bath. The separated DNA strands were then added quickly to a 500 µL solution (3.44 pmol) of P1modified beads at a temperature of 70 °C in a water bath. The ratio of DNA to bead was kept at 1:1. After thorough mixing, the solution containing the P1-beads and the denatured DNA chains was slowly cooled until the temperature of the bath dropped to 30 °C. It is during this stage that the DNA chains have a chance to hybridize with the complementary oligomers tethered on the beads. To test the effect of a noncomplementary primer, experiments where the P1-beads were replaced by P2-beads were also performed. The same entire protocol was followed for ∆DIIIT7 DNA, the only difference being in the amount of DNA used, 100 µg instead of 10 µg, to keep the DNA to bead ratio equal to unity despite the 10 times higher molecular weight of ∆DIIIT7 DNA. Following the hybridization procedure, the beads were electrodialyzed at 50 V for 2 h to detach the physically adsorbed DNA chains from the beads. The beads were then examined by electrophoresis using 0.8% agarose gels (Figure 3). The ∼0.2 µm pore size of such gels is small enough to block the migration of the beads but not that of individual DNA chains.33 The absence of detectable bands in the gel was taken as the proof of complete removal of physisorbed DNA during the electrodialysis step. The beads were then washed, resuspended in 500 µL of water, and studied both optically and spectrophotometrically (UV and fluorescence). 5. Fluorimetric Studies on Bead-Bound DNA. Fluorescence measurements were carried out using a SPEX Fluoromax spectrofluorimeter, at excitation and emission wavelengths of 485 and 515 nm, respectively, and with slits set at 2.5 nm bandpass. Data points were taken every 5 s. Titrations were done in TE buffer (10 mM Tris-HCl, pH 8.2, 1 mM EDTA). Trizma Base and EDTA were purchased from Sigma-Aldrich. YOYO-1 (the common name for the monomer YO-PRO1 (YO) is oxazole yellow (33) Tietz, D.; Chrambach, A. Electrophoresis 1992, 13, 286.

DNA Immobilized on PS Microspheres

Figure 3. (a) 0.8% agarose gel showing T7A1 DNA preparation and its subsequent attachment to modified PS microspheres (gel run in 1X TAE buffer (40 mM Tris acetate, pH 8.0, 1 mM EDTA) at 100 V): lane 1, T7A1 DNA preparation; lane 2, linearized T7A1 DNA obtained by digesting overnight with EcoRV; lane 3, P1-beads hybridized with linearized T7A1 DNA; lane 4, P1-beads hybridized with linearized T7A1 DNA and electrodialyzed. (b) 0.6% agarose gel showing ∆DIIIT7 DNA preparation and its subsequent attachment to modified PS microspheres (gel run in 1X TAE buffer (40 mM Tris acetate, pH 8.0, 1 mM EDTA) at 100 V): lane 1, ∆DIIIT7 DNA preparation; lane 2, P2-beads hybridized with ∆DIIIT7 DNA; lane 3, P2-beads hybridized with ∆DIIIT7 DNA and electrodialyzed. and YOYO is its dimeric form) and quinolinium, 1,1′-[1,3propanediyl bis[(dimethylimino)-3,1-propanediyl]] bis[4-[3-methyl-2(3H)-benzo-oxazolylidene) methyl]]-tetraiodide)) were purchased from Molecular Probes, Illinois. YOYO-1 stock (provided in DMSO) was stored at -24 °C. A dilution of the stock was made in TE buffer to ∼1 µM concentration before each series of experiments. The YOYO-1 solution was kept in the dark at room temperature and vortexed prior to each experiment. The stability of the YOYO-1 turned out to be very critical for experimental reproducibility, so careful handling is required. Preparation of the solution and bead-bound DNA used for the experiments has been described in detail earlier. DNA and YOYO concentrations were measured by spectrophotometry using a Perkin-Elmer Lambda 800 UV/vis spectrophotometer. The concentrations were deduced from the absorbances using the following values: (457 nm) ) 96 100 M-1‚cm-1 for YOYO, and 1 OD unit at 260 nm is equivalent to 50 µg/mL of DNA. The intercalation of YOYO into DNA increased its fluorescence intensity as expected. Thus YOYO was added to the DNA solution progressively until the intensity increased and reached a maximum (YOYO was added up to a ratio of 1 dye molecule for 2 bp), and no more DNA was available

Langmuir, Vol. 19, No. 14, 2003 5833 for intercalation. During the fluorescence titrations, YOYO was added progressively to the DNA or bead suspension, one injection being done every 40 s. In every experiment, the DNA amount was set at 0.116 µg (0.005 pmol), and 4.02 pmol YOYO was added per injection. 6. Microscopy. 6.1. Fluorescence Microscopy. The setup used for all fluorescence microscopy experiments was a Leica DM IRB microscope, with a Leica PL FLUOTAR 100× oil immersion objective. Picture acquisition was carried out using a Lhesa LH 509 ULL intensified camera connected to a computer. Samples were dyed with YOYO at least 1 h prior to each experiment, the DNA base pairs to YOYO ratio being set at 5:1. The YOYO/DNA complex fluoresces 3200 times as much as that of free YOYO. The intercalation of the YOYO into the DNA has no major influence on its physicochemical properties, considering the amount of dye added (1 dye molecule for 5 bp), which permits detection of the presence of DNA and estimation of its local density. 6.1.1. Preparation of Amine Slides. Amine-modified microscope slides were prepared starting from cover glasses (Marienfeld no. 1, 24 × 60 mm). The cover glasses were first cleaned in “piranha” solution (30% hydrogen peroxide/70% sulfuric acid) at room temperature for 15 min. The glasses were then rinsed thoroughly with Nanopure water and stored in water for no more than a few hours. After rinsing in methanol, the slides were immersed in a reactor containing 40 mL of methanol, 344 µL of acetic acid, and 1.7 mL of water. N-[3-(Trimethoxysilyl)propyl]-ethylenediamine (850 µL) (purchased from SigmaAldrich) was then injected in the reactor and was kept at room temperature for 2 h. The slides were then rinsed with methanol and dried in a 70 °C oven for 10 min. 6.1.2. Microscopy. For microscopy experiments, the 5% beadbound DNA suspension was diluted 100-fold and incubated with the required amount of YOYO. A 7 µL drop of that sample was put on a bare cover glass and was covered by an amine-modified slide. During spreading of the sample between the two slides, hydrodynamic forces stretched the DNA molecules. Moreover, DNA molecules that get in contact with the amine-modified surface remained adsorbed on it due to electrostatic interactions. A few seconds after placing the amine slide on the drop, all of the DNA molecules were immobilized on its surface, some of them being stretched. DNA attached to beads can also be adsorbed, permitting an observation of the DNA-bead complex. 6.2. Bright-Field Microscopy. The setup used for brightfield microscopy was a Leica DMRM microscope with either a Leica PL FLUOTAR 100× oil immersion objective or a Leica PLAN L 40× air objective. Picture acquisition was carried out using a Roper Scientific Coolsnap camera connected to a computer.

Results and Discussion 1. Hybridization of ∆DIIIT7 DNA (39.34 kbp) or T7A1 DNA (4.48 kbp) ds-DNA on Primer-Beads. Quantitation of DNA Attachment to Beads by UV-Vis Spectroscopy. The amount of primers and DNA binding to the MITC-beads was determined from a comparison between the residual amount of primer/DNA in each supernatant and the amount of primer/DNA added into the sample, adapted from previously published methods.34 The values were estimated by UV analysis from the maximum absorbance of the primer or DNA at 260 nm. The details of the quantitation are given in Table 1. The primer attachment reactions were found to be very efficient, and the final attachment of the primers to the MITC-beads was found to be of molar ratio 1:180 for P1 and 1:225 for P2. In the case of DNA, the longer DNA attachment by hybridization was more efficient than that of the shorter DNA. Thus, approximately 35% of the added T7A1 DNA remained bound to the beads, making a final ratio of bead/DNA of 1:0.35, whereas in the case of ∆DIIIT7 DNA, 60-80% attachment of the DNA was achieved, thus (34) Charreyre, M.-T.; Tcherkasskaya, O.; Winnik, M. A.; Hiver, A.; Delair, T.; Cros, P.; Pichot, C.; Mandrand, B. Langmuir 1997, 13, 3103.

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Table 1. Quantitation of DNA Attachment to MITC-Beads from absorbance at 260 nma sample initial ratiob MITC-B + P1 (B/P1 ) 1:250) P1-B + T7A1 (B/DNA ) 1:1) MITC-B + P2 (B/P2 ) 1:300) P2-B + ∆DIIIT7 (B/DNA ) 1:1)

average supernatant attached attachment DNA (%) DNA (%) (%)

final ratio B/P1 ) 1:179.50

30.8

69.2

30.8 23.1 70.9

69.2 76.9 29.1

71.8

61.3 61.3 22.7

38.7 38.7 77.3

35.5

26.3 26.3 32.2

73.7 73.7 67.8

74.9

25.1 35.5

74.9 64.5

69.1

B/DNA ) 1:0.35

B/P2 ) 1:225

B/DNA ) 1:0.69

a After each reaction, beads were washed by centrifugation and the supernatants were stored. Values were obtained by measuring the absorbance of the supernatant at 260 nm wavelength. b B, beads; P1, primer 1; P2, primer 2; T7A1, T7A1 DNA; ∆DIIIT7, ∆DIIIT7 DNA.

Figure 5. Fluorescence micrographs of (a) single ∆DIIIT7 DNA molecules attached to single P2-beads and extended on aminemodified glass slides and (b) ∆DIIIT7 DNA molecules extended on an amine-modified glass slide.

Figure 4. Fluorescence micrograph of bead-bound ∆DIIIT7 DNA combed and extended on a glass slide.

getting a final ratio of bead/DNA of 1:0.6-0.8. We think the shorter DNA has a coiled conformation disallowing overlapping of the hybridization sequence, but this remains to be tested. Optical Fluorescence and Bright-Field Microscopy. The primer-beads hybridized with T7A1 DNA (4.48 kbp) or ∆DIIIT7 DNA (39.34 kbp) and subsequently washed with water by centrifugation when viewed under the brightfield microscope showed well-dispersed beads (60-70%) with a few small aggregates (∼30%). These bead-bound DNAs after subsequent staining with YOYO were seen between two glass slides under the fluorescence microscope as described above. As the dye does not show any fluorescence alone or with the beads, the only fluorescence emission that can take place is from the dye-DNA complex bound to the beads. In the case of the T7A1 DNA, it appears that the DNA is attached with the bead and perhaps is completely wrapped around (not shown). However, the length of the DNA being short (4.48 kbp), no distinct shape was observable. Figure 4 shows a fluorescence micrograph of a few aggregates of beads (2-10) attached to ∆DIIIT7 DNA stretched on the surface of a glass slide due to combing by the receding meniscus. The estimate of the aggregation was made from the diameter of the aggregate

as observed under the bright-field microscope. This image showed that DNA is present on the bead and that it is dangling. Next, the bead-bound ∆DIIIT7 DNA was deliberately allowed to stretch on an amine-modified glass surface, and photographs were taken. Figure 5a shows representative pictures in which the DNA is seen to be dangling from the bead and stretched on the amine slides. The photographs show that the DNA is covalently endgrafted on the polystyrene microspheres. In addition, no unbound free DNA was seen, indicating that no DNA was released after the washing step. Bright-field microscopy confirmed that a single DNA was attached to one bead. Fluorescence intensity and comparison with a normalized DNA solution provided an average estimate of one DNA per bead and also showed that the systems were quite homogeneous; 60-70% of the population of beads contain DNA consistent with the results obtained from UV quantitation studies. As a control, the solution DNA was also stretched on amine slides (Figure 5b) and shows a profile similar to that of the bead-bound DNA. Different batches of hundreds of beads were scanned throughout the slide, and in each batch, 60-70% of the beads were calculated to have DNA as seen under the microscope. We observed that only one DNA was stretched (Figure 5a) and the other DNA molecules were not stretched, giving rise to bright fluorescence on the spherical beads. However, in the control (Figure 5b) the DNA molecule is attached to the aminated surface without beads and does get coiled up, in some cases giving the appearance of a small sphere. As described earlier, an alternate protocol was also used for DNA attachment in which beads were first made free of any chemical or surfactant by washing with methanol. The beads were redispersed by sonication and subsequently hybridized with the ∆DIIIT7 DNA. Here also endgrafting was seen as in Figure 5a (figure not shown), but the number of bead-bound DNA molecules stretched was much less in this case. This gave us an indication that

DNA Immobilized on PS Microspheres

DNA was more tightly held by the beads washed with methanol compared with the ones washed with water. 2. Hybridization as against Nonspecific Physisorption. Even if the hybridization had taken place between the primers present on the beads and the added DNA, the possibility of nonspecific physisorption was always present. Figure 3 shows the results of gel electrophoresis of the beads before and after electrodialysis was performed to remove the physisorbed DNA chains. A migrating band was observed in lane 3 but not in lane 4 of Figure 3a and in lane 2 but not in lane 3 of Figure 3b as the physisorbed DNA is loosely bound and would detach from beads under the electric field and migrate into the gel. This indicates the presence of physisorbed chains on the beads before electrodialysis. We estimated from the densitometric analysis of gels that about 10% of the chains were physisorbed rather than hybridized. The chemically bound DNA was evident from the fluorescence in the well before or after electrodialysis. 3. Hybridization Kinetics of the Linear T7A1 DNA with P1-Beads. To investigate the influence of temperature on the hybridization rate, the hybridization between the P1-beads and T7A1 DNA was repeated at several temperatures between 25 and 60 °C. At any particular temperature, aliquots of the mixture were taken out over a period of 1 h at different time intervals of 1, 5, and 10 min, loaded on 0.8% agarose gels, and electrophoresed. EtBr-stained wells of the gels were then scanned for densitometric analyses carried out with a BioRad Imaging system using QuantityOne software, version 4.1.1, Build003. The analyses gave the values of intensity of EtBr fluorescence per unit area. These values of intensity were plotted against time to give the saturation kinetics (Figure 6a) under a pseudo-first-order condition since [P1] . [T7A1 DNA]. Unattached DNA was included as an internal standard during analysis. Since the precise initial and final concentrations of the reactants were not known, a Gugenheim plot35 was used to obtain first-order rate constants from each of the kinetics experiments (Table 2). The rate of hybridization of T7A1 DNA with the primer P1 in the heterogeneous phase was directly proportional to the temperature, which is, however, expected (Table 2). The values of the rate constants were in the same range as those obtained earlier18 where one of the partners was immobilized. However, the rate was extremely slow when compared with the bimolecular rate constant for selfassociation of complementary strands of DNA20 which was in the millisecond range. The natural logarithms of rate constants were plotted against inverse of temperature to ascertain the activation energy of the hybridization reaction in the heterogeneous phase, following the relationship

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Figure 6. (a) Illustrative data for hybridization kinetics of T7A1 DNA with primer P1 attached to polystyrene microspheres at 328 K. The lower curve shows a similar hybridization reaction of T7A1 DNA with primer P2 with no change through the entire time range. (b) Plot of ln k (k ) rate constant) versus 1/T (T ) temperature in kelvin), giving the activation energy for the hybridization kinetics of T7A1 DNA with primer P1 in the heterogeneous phase (P1 attached to the polystyrene microspheres (O)). The range of error associated with the experimental data is also shown. The dotted lines show the maximum range to which the activation energy values can vary.

where ln k is the natural logarithm of the rate constant at a particular temperature, Ea is the activation energy, R is the universal gas constant, and T is the temperature in kelvin. A plot of ln k versus 1/T gave the value of Arrhenius activation energy, Ea (Figure 6b, Table 2). The best-fit line with linear regression analysis yielded a value of ∼5.3 kcal/mol. However, the two extreme cases are shown in Figure 6b, which were drawn considering the four closest points in linear regression, and the values were 10.4 and 4.2 kcal/mol. The theoretical limit of the

energy of diffusion (ω) in a particular solvent from the temperature dependence of the viscosity equation η ) η0 exp ω/KT, where K is Boltzmann’s constant, generates ω for water36 to be ∼3.5 kcal/mol at room temperature. Thus it appears that the reaction rate in our experiments is non-diffusion-controlled. This suggests that the rate of the reaction is governed by the rate at which the linearized T7A1 DNA can diffuse toward the bead-bound P1 and form hydrogen bonds with it and the presence of beads poses a barrier to the reaction by creating a heterogeneous phase. Presence of Base Mismatches Abrogates Hybridization. For P2 (which is complementary to ∆DIIIT7 DNA and is a 14 out of 20 bases mismatch to P1, sequence given in Figure 1), hybridization was done with T7A1 DNA only at 55 °C (328 K). Since it was a mismatch oligomer, we did not expect any binding kinetics. Though there was background absorption of the T7A1 DNA on P2-beads, the hybridization did not show any kinetics (Figure 6a). Thus, in all our rate constant calculations, these back-

(35) Frost, A. A.; Pearson, R. G. Kinetics and Mechanisms; Wiley Eastern Pvt. Ltd.: New Delhi, 1970; p 49.

(36) Weast, R. C. CRC Handbook of Chemistry and Physics; CRC Press: Boca Raton, FL, 1969; p F36.

ln k ) -Ea/RT + constant

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Ghosh et al.

Table 2. First-Order Rate Constants and Activation Energy Data for the Hybridization of Bead-Bound Primers to T7A1 DNA

primer P1

P2 P3

temp (K)

kb (s-1) × 103 (solid)a

error

333 328 323 318 313 298 328 328 318 298

1.16 1.42 1.17 0.80 0.69 0.55

0.0740 -0.0290 -8.8000 × 10-3 -0.2400 -0.1043 -7.1000 × 10-3

0.95

activation energy (kJ) (solid)

}

22.0 (5.25 kcal)

range of activation energy (kJ) (solid) higherb

lowerc

43.7 (10.4 kcal)

17.6 (4.2 kcal)

0.0124

a k are the rate constants of hybridization when the primer is attached to the bead. b Calculated based on the rate constants for the b temperatures 328, 323, 318, and 313 K only. c Calculated based on the rate constants for the temperatures 333, 318, 313, and 298 K only.

grounds were subtracted from the actual data. For P3, with one base mismatch, hybridization was done at 55 °C (328 K), 45 °C (318 K), and 25 °C (298 K). At higher temperature (55 °C), the hybridization reaction with P3 showed similar saturation kinetics as in the case of P1. However, at lower temperatures (45 and 25 °C), the hybridization reaction did not proceed and showed background fluorescence as in the case of P2. Comparing the rate constants for P1 and P3 at 55 °C, it was seen that the rate of the reaction was faster for P1 than for P3 as expected (Table 2). The half-life of a reaction of hybridization with P1, t1/2, at 55 °C was found to be 8 min, and that for the reaction of the template DNA with P3 was found to be 12 min. This appears to be an important observation and is in accordance with those obtained earlier18 on similar hybridization reactions, which showed that by manipulation of the number of mismatches in the bases, the rate of a reaction could be varied. 4. Interaction of ds-DNA with Primer-Beads before or after Hybridization. Fluorescence experiments were performed in order to gather additional information about the nature of interaction of ds-DNA with beads. Our strategy was to find out how the probe YOYO intercalates DNA when they were hybridized to the beadbound primer or just present in the reaction mixture without any covalent attachment with the beads. As we have mentioned earlier, free YOYO shows appreciable enhancement of fluorescence upon binding to DNA. However, prior to this experiment we have prepared beads in two different ways, that is, by washing them with water or with methanol. Figure 7 shows the typical enhancement of fluorescence of YOYO leading to saturation as a function of increasing concentration when added to a fixed concentration of DNA. It is clear from the figure that the free DNA or water-washed bead-bound DNA showed similar titration profiles, indicating thereby that the DNA was available for YOYO binding. On the other hand, when bead-bound DNA was prepared by methanol, the increase in fluorescence intensity was much less in the same time scale and we inferred that less DNA was accessible for YOYO. This result in a way supports the conclusion drawn from microscopic studies reported in an earlier section where the DNA bound to the methanol-washed beads was more difficult to stretch by normal stretching conditions of DNA. It appears as though upon methanol washing of the bead-DNA adduct, DNA was not free and collapsed on the beads by adsorption. This conclusion is in accordance with the results obtained earlier.30 It was observed that in the presence of surfactants on the beads, the beads would not be able to adsorb DNA. Upon washing with methanol, beads were progressively getting devoid

Figure 7. Fluorescence titration curves of ∆DIIIT7 DNA in solution and bead-bound form (bound to water-washed or methanol-washed P2-beads), upon the addition of YOYO.

of surfactants, resulting in physical adsorption of DNA by simple charge interaction. Our control experiments showed that when primerbeads were added to the mixture in the presence of DNA without hybridization, DNA was available for intercalation with YOYO (Figure 8a). Washing with methanol or water did not make any difference here in comparison to the free DNA. However, MITC-beads, water- or methanolwashed, adsorb DNA to some extent, and thus the resulting fluorescence intensity upon YOYO binding was less when compared with that of free DNA (Figure 8b). This explains why interaction was observed between the DNA and the beads after hybridization, since under that condition, the DNA interacts with the free MITC on the beads and makes itself less available for YOYO intercalation. Conclusion Through several experiments, we have clearly been able to establish the success of hybridization for grafting dsDNAs to oligonucleotides immobilized on the surface of a polystyrene microsphere. This reaction takes place irrespective of the size of the DNA and thus establishes an easy and convenient method for grafting of long DNAs to surfaces. Through our experiments, we could also optimize the best conditions for cleaning the beads so as to get an free, accessible immobilized DNA. This is very important since now this well-characterized system can be used further to perform various biological reactions. From optical fluorescence microscopy, it is evident that a long DNA (∼40 kbp) is covalently end-grafted on a latex

DNA Immobilized on PS Microspheres

Figure 8. (a) Fluorescence titration curves of ∆DIIIT7 DNA in solution and in the presence of P2-beads (water-washed or methanol-washed) without hybridization, upon the addition of YOYO. (b) Fluorescence titration curves of ∆DIIIT7 DNA in solution and in the presence of MITC-beads (water-washed or methanol-washed) without hybridization, upon the addition of YOYO.

particle by hybridization, and this, to the best of our knowledge, has not been documented before in the literature. The end-grafted DNA is free and close to the stoichiometric ratio of the beads present in the system. According to Poisson statistics, the proportion of beads with 0, 1, and 2 DNA per bead should be 33%, 50%, and 15%, respectively. This supposes that the probability of hybridization is very large and that there is no restriction on the number of chains per bead. Actually, the size of the beads (0.31 µm) is much lower than the radius of gyration, Rg, of the DNA. One can calculate Rg to be 0.37 and 1.16 µm for double-strand chains with 4000 or 40 000 base pairs, respectively. Steric constraints will therefore limit the maximum number of DNA per bead to 1. Physisorption of the tethered DNA strand is prevented by the unhybridized primer P2; electrostatic repulsion is playing an important role to keep the DNA free and dangling from the bead. If P2 is not present, physisorption is observed between the DNA and the MITC on the beads, mostly by charge interaction. This criterion is however overcome by the absence of surfactants on the beads, where physical adsorption is favored and the DNA is more tightly bound to the beads making it less available or dangling even in the presence of excess unhybridized primer.

Langmuir, Vol. 19, No. 14, 2003 5837

Hybridization kinetics was followed for a ds-DNA attachment to a primer DNA in the heterogeneous phase. Hybridization to the mismatch primer showed no kinetics of binding but a background fluorescence suggesting that there was nonspecific attachment of DNA to the beads, but hybridization was possible only when the sequence was exactly complementary. Introduction of one mismatch in the complementary sequence also retarded the rate of hybridization as expected. The activation energy for the hybridization of the T7A1 DNA to the immobilized primer is high, suggesting a barrier in the reaction due to the introduction of the heterogeneous phase correlating with a non-diffusion-limited reaction. This suggested that the position for hybridization was not properly accessible when one of the partners was immobilized. It has been shown recently18 that the distance of the hybridizing sequence from the support point is also a predominant factor for hybridization to occur. In this case, hybridization of DNA complementary strands was attempted keeping one strand immobile, and the immobile surface was a planar gold plate18 on which thiol-based oligonucleotides were immobilized and the hybridization to complementary strands was studied. Here, we have shown a hybridization reaction through hydrogen bonding on a surface, which in its own merit is unique, since a double-stranded DNA is attached to a latex particle through a linker which is covalently bound. The present system is also more versatile than that reported earlier,18 since here the DNA is attached to a microsphere and is present as a suspension, and so the reactions on the DNA can be performed in bulk, at the air-water interface, or even at the solid-air interface by transferring the solid on a metal surface in the form of a monolayer or film. The efficiency of hybridization is high, 60-70%, and comparable to the efficiency in solution.23 In principle, there is a competition between the two strands of the dsDNA hybridization and single-strand hybridization to a smaller complementary oligomer. In solution, the second reaction is favored due to the high diffusion, since the smaller unit will diffuse faster in space than the long single strand of the DNA. When the oligomer is immobilized, we have to compare the diffusion coefficient of the bead itself to that of the long single-stranded DNA. Depending on the size of the bead, it can be smaller or larger than the DNA. In the present case, the size of the bead is smaller than the Rg of the DNA. Therefore, it is not surprising to find a high efficiency similar to that for the oligomer in solution. Acknowledgment. This work was carried out under Indo-French Grant Number 2303-1 from IFCPAR (CEFIPRA). We also thank Professor R. Varadarajan for letting us use the BioRad Imaging system. List of Abbreviations ds PS DITC MITC P1 P2 P3

double stranded polystyrene 1,4-phenylene di-isothiocyanate phenylene mono-isothiocyanate primer 1 primer 2 primer 3

LA0341963