- Email: [email protected]
Observation of stretched single DNA molecules by Kelvin probe force microscopy
Applied Surface Science 210 (2003) 73–78
Observation of stretched single DNA molecules by Kelvin probe force microscopy K.J. Kwak, S. Yoda, M. Fujihira* Department of Biomolecular Engineering, Tokyo Institute of Technology, 4259 Nagatsuta, Midori-ku, Yokohama 226-8501, Japan
Abstract Lambda bacteriophage deoxyribonucleic acid (DNA) molecules stained with YOYO-1 were deposited and stretched on chemically modified Si(1 0 0) wafer and cover glass substrates. The Si(1 0 0) wafer with a natural oxide layer and the cover glass surface were first chemically modified by vapor phase chemisorption with mixed two organosilanes. DNA molecules were then aligned on the substrate surfaces by chemical and physical adsorption from an aqueous solution during molecular combing. The aligned DNA molecules were observed by Kelvin probe force microscopy and intermittent contact-mode atomic force microscopy (AFM) as well as fluorescence microscopy. Interaction between the stretched DNA molecules and the chemically modified Si(1 0 0) substrate surface was examined using these AFMs. # 2002 Elsevier Science B.V. All rights reserved. PACS: 07.79.Lh; 61.16.Ch; 87.64.Dz Keywords: Stretched DNA; Molecular combing; Kelvin probe force microscopy; Atomic force microscopy
1. Introduction The study of interactions of biological molecules with a solid surface is one of the most important issues of atomic force microscopy (AFM) for its application to biological samples [1,2]. A cleaved mica surface as a flat substrate has been extensively used to prepare such a biological sample as deoxyribonucleic acid (DNA) [3–8]. When DNA molecules are bound to a substrate surface, it is desirable that DNA molecules can still hybridize with their complementary strands or interact with proteins in the bound forms. The binding of DNA molecules to a chemically treated mica surface can be *
Corresponding author. Tel.: þ81-45-924-5784; fax: þ81-45-924-5817. E-mail address: [email protected] (M. Fujihira).
strong enough to lose such biological activities [9–11]. Additionally, alignment of the immobilized DNA molecules on the flat substrate is another interesting issue of AFM [12–18]. A reproducible process called a molecular combing method has been demonstrated for alignment of DNA molecules [12,13]. The DNA strands can be uniformly stretched on the chemically modified surface with a CH2¼CH-terminated group by a hydrodynamic force of a receding meniscus. Recently, noncontact-mode AFM (nc-AFM) in ultrahigh vacuum has been demonstrated to image the DNA molecules with high accuracy and high resolution [19–21]. This technique has been extended further to the high resolution imaging DNA molecules [22–24]. However, there has been little report on interactions between DNA molecules and substrate surfaces [25].
0169-4332/02/$ – see front matter # 2002 Elsevier Science B.V. All rights reserved. doi:10.1016/S0169-4332(02)01482-4
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K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
In this work, we have demonstrated nc-AFM to image the stretched DNA molecules in air. A cantilever with an Au-coated tip was chemically modified by an organothiol compound and applied to stable observation of DNA by Kelvin probe force microscopy (KFM) [26–28]. The stretched DNA strands were deposited on mixed silane monolayers of CH2¼CH- and NH2-terminated silanes chemisorbed on oxide substrate surfaces. It is important to prepare samples having the optimized surface density of DNA strands for the AFM study.
N.A. oil immersion lens. All SPM measurements including intermittent contact (i.e. TappingTM mode) AFM and KFM were performed using an SPA-400 AFM unit and an SPI-3800 control station (Seiko Instruments). Cantilevers with an Au-coated tip having a normal spring constant of 1.6 N m1 and resonance frequency of 23–25 kHz were directly used after cleaning the tip [30] for intermittent contact AFM or chemically modified with hexadecanethiol (HDT) for KFM.
3. Results and discussion 2. Experimental 3.1. Fluorescence microscopy 2.1. Preparation of stretched DNA molecules Organosilane compounds for modification of oxide substrate surfaces were purchased from Aldrich Chemical Co. and further purified by distillation. 7-Octen1-yl (CH2¼CH-terminal) and 3-(2-aminoethyl)aminopropyl (NH2-terminal) trimethoxysilanes were used as silanization reagents. Si(1 0 0) wafers (n-type high doped; Nilaco Co.) and glass coverslips from Matsunami Glass Co. were used as the oxide substrates for AFM and/or fluorescence microscopy. Lambda bacteriophage DNA molecules (48,502 bp; Takara Bio Inc., Japan) were stained with YOYO-1 at a ratio of DNA base pairs to the YOYO-1 molecules of 5:1. The Si(1 0 0) wafers were cleaned and oxidized using a UV-ozone cleaner (Nippon Laser and Electronics Laboratory) for 1 h and/or piranha solution treatments [29]. Glass coverslips were cleaned by the piranha solution treatments. The silanized substrate surfaces were prepared by vapor phase chemisorption with a mixed two-component liquid of the organosilane compounds with a 1:10 (v/v) ratio of NH2- to CH2¼CH-terminated silanes. The silanized substrates were dipped into a TE buffer (10 mM Tris, 1 mM EDTA) containing the stained DNA molecules diluted to 0.45 mg ml1. After 5 min of incubation, the silanized substrates were lifted up with a mechanical apparatus at a constant speed of 300 mm s1. 2.2. Imaging of DNA molecules Fluorescence microscopy was performed using a Nikon microscope (Diaphot 300) with a 100/1.30
The adsorption of the lambda bacteriophage DNA molecules on the cover glass substrate by the molecular combing method [13] was easily examined using the fluorescence microscope as shown in Fig. 1. The fluorescent dye YOYO-1 showed an excitation maximum at 491 nm and an emission maximum at 509 nm when binding to DNA molecules. Fluorescence microscopy could be used to confirm the optimized surface density of the adsorbed DNA strands for AFM imaging. Here, the two-component system in the modification of substrates was carried out with the mixed gas consisting of NH2- and CH2¼CH-terminated silanes. The increase in the surface density of DNA shown in Fig. 1 in comparison with that on a CH2¼CH-terminated silane monolayer (not shown here) indicates that the DNA strands are negatively charged in aqueous solution at neutral pH, and thus the DNA molecules would be electrostatically attracted to positively charged NH2-terminals [9,11]. 3.2. AFM measurements The stretched DNA strands were observed by intermittent contact AFM with an Au-coated tip cleaned by the UV-ozone treatment. Fig. 2(a) and (b) show a topographic and a phase shift image, respectively, obtained in 45.5% relative humidity (RH) air. Both topography and phase shift on the DNA strands show higher heights and larger phase shifts (i.e. greater phase-lag) than those of the substrate surface, respectively. The greater phase-lag can be attributed to the surface water layer on the DNA strands, which
K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
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Fig. 1. A fluorescence microscopic image of the DNA strands deposited and stretched by the dynamic molecular combing method on the twocomponent NH2- and CH2¼CH-terminated substrate surface.
enhanced the adhesive force between the tip and the strands [31–33]. The higher adhesive force on the strands than on the hydrophobic substrate surface was shown previously by pulsed-force-mode AFM [34].
The present surface monolayer prepared by vapor phase chemisorption with a 1:10 (v/v) mixture seems to contain ca. 105 mole fraction of NH2-terminals from the previous results with a 1:1 (v/v) mixture [34].
Fig. 2. 3 mm 3 mm (a) topographic and (b) phase images simultaneously observed by intermittent contact AFM. Two images were acquired in 45.5% RH air.
76 K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78 Fig. 3. 1:5 mm 1:5 mm (a) topographic image observed by intermittent contact AFM and (b) topographic and (c) CPD images simultaneously observed by KFM. Three images were acquired in 48.9% RH air with an HDT-modified Au tip. The profiles of (d) and (e) along the solid lines in (a) and (b), respectively.
K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
In higher humidity than 60% RH, this enhanced adhesive force resulted in cleaving the DNA strands to their fragments (not shown here). For getting the higher spatial resolution of the contact potential difference (CPD), we used the method called KFM [28]. Fig. 3(a) shows topographic image observed by intermittent contact AFM. Fig. 3(b) and (c) show a topographic and a CPD image simultaneously observed by KFM, respectively, scanned over the same area as that in Fig. 3(a). Here, the CPD of the sample was measured against an HDTmodified Au tip. Three images were acquired in 48.9% RH air. The a.c. modulation for the KFM measurements was applied to the sample substrate with a driving a.c. frequency of 22 kHz and amplitude of 5 V (peak to peak). The CPD distributions on the silane-modified Si(1 0 0) surface ranged from 0.34 to 0.40 V as shown in Fig. 3(c). This inhomogeneity of the CPD observed on the substrate surface may be attributed to the mixed monolayer of NH2- and CH2¼CH-terminated silanes. If the mole fraction of NH2-terminated silane is ca. 105 as described above and the cross-sectional area of an alkyl chain is 0.2 nm2, the surface density of NH2-terminated silane will be ca. 50 molecules mm2. The number of the bright spots observed in Fig. 3(c) is in this order. However, origin of the increase in CPD due to NH2terminals has not been clarified yet. If the NH2-terminals are ionized, we will expect the decrease in CPD, while if the NH2-terminals are not ionized, the increase in CPD will be expected due to permanent dipoles of the NH2-terminals [35]. More interestingly, three bright lines in Fig. 3(c) corresponding well with DNA strands observed in Fig. 3(a) and (b) seem to be pinned by the several bright spots on the substrate surface. This suggests that NH2-terminal groups make ion complexes with phosphate anions on DNA. The bright lines indicate that the stretched DNA strands have the higher CPD than that on the substrate surface. The cylindrical symmetry of DNA suggests that the CPD would not be affected significantly by DNA adsorption on the substrate. The result suggests that the dipole moments formed between the surface water layer and the negatively charged DNA strands have a direction from the substrate toward air. Fig. 3(d) and (e) show the profiles along the solid lines in Fig. 3(a) and (b), respectively. The stretched DNA strands on the substrate surface have a width of
77
34 nm and a height of 0.7 nm as shown in Fig. 3(d), which are comparable to those in Fig. 3(e). This typical result indicates that the spatial resolution of the DNA strands was determined mainly by the size of the tip end [3,5]. The smaller height than 2.4 nm of van der Waals radii of DNA [23] also agrees with those measured recently by nc-AFM with the frequency modulation technique [22,23,25] and by STM [36].
4. Conclusions The stretched DNA strands on the silanized substrate surfaces were observed by KFM and intermittent contact AFM as well as fluorescence microscopy for the study of interactions between the DNA molecules and the substrate surfaces. The cantilever with the HDT-modified Au tip was applied to the stable observation of the DNA molecules by KFM.
Acknowledgements This work was supported by a Grant-in-Aid for Creative Scientific Research on ‘‘Devices on molecular and DNA levels’’ (No. 13GS0017) from the Ministry of Education, Science, Sports, and Culture of Japan.
References [1] R.J. Colton, D.R. Baselt, Y.F. Dufrene, J.B.D. Green, G.U. Lee, Curr. Opin. Chem. Biol. 1 (1997) 370. [2] H.G. Hansma, Ann. Rev. Phys. Chem. 52 (2001) 71. [3] J. Vesenka, M. Guthold, C.L. Tang, D. Keller, E. Delaine, C. Bustamante, Ultramicroscopy 42–44 (1992) 1243. [4] T. Thundat, D.P. Allison, R.J. Warmack, G.M. Brown, K.B. Jacobson, J.J. Schrick, T.L. Ferrell, Scan. Micros. 6 (1992) 911. [5] C. Bustamante, J. Vesenka, C.L. Tang, W. Rees, M. Guthold, R. Keller, Biochemistry 31 (1992) 22. [6] H.G. Hansma, J. Vesenka, C. Siegerist, G. Kelderman, H. Morrett, R.L. Sinsheimer, V. Elings, C. Bustamante, P.K. Hansma, Science 256 (1992) 1180. [7] Y.L. Lyubchenko, A.A. Gall, L.S. Shlyakhtenko, R.E. Harrington, B.L. Jacobs, P.I. Oden, S.M. Lindsay, J. Biomol. Struct. Dyn. 10 (1992) 589. [8] Y.L. Lyubchenko, L.S. Shlyakhtenko, R.E. Harrington, P.I. Oden, S.M. Lindsay, Proc. Natl. Acad. Sci. USA 90 (1993) 2137. [9] M. Bezanilla, S. Manne, D.E. Laney, Y.L. Lyubchenko, H.G. Hansma, Langmuir 11 (1995) 655.
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[10] H.G. Hansma, I. Revenko, K. Kim, D.E. Laney, Nucl. Acids Res. 24 (1996) 713. [11] V. Balladur, A. Theretz, B. Mandrand, J. Coll. Interf. Sci. 194 (1997) 408. [12] A. Bensimon, A. Simon, A. Chiffaudel, V. Croquette, F. Heslot, D. Bensimon, Science 265 (1994) 2096. [13] X. Michalet, R. Ekong, F. Fougerousse, S. Rousseaux, C. Schurra, N. Hornigold, M. van Slegtenhorst, J. Wolfe, S. Povey, J.S. Beckmann, A. Bensimon, Science 277 (1997) 1518. [14] Z.-Q. Ouyang, J. Hu, S.-F. Chen, J.-L. Sun, M.-Q. Li, J. Vac. Sci. Technol. B 15 (1997) 1385. [15] W. Wang, J. Lin, D.C. Schwartz, Biophys. J. 75 (1998) 513. [16] H. Yokota, F. Johnson, H. Lu, R.M. Robinson, A.M. Belu, M.D. Garrison, B.D. Ratner, B.J. Trask, D.L. Miller, Nucl. Acids Res. 25 (1997) 1064. [17] H. Yokota, J. Sunwoo, M. Sarikaya, G. van den Engh, R. Aebersold, Anal. Chem. 71 (1999) 4418. [18] J.Y. Ye, K. Umemura, M. Ishikawa, R. Kuroda, Anal. Biochem. 281 (2000) 21. [19] Y. Maeda, T. Matsumoto, T. Kawai, Appl. Surf. Sci. 140 (1999) 400. [20] Y. Maeda, T. Matsumoto, H. Tanaka, T. Kawai, Jpn. J. Appl. Phys. 38 (1999) L1211. [21] T. Matsumoto, Y. Maeda, Y. Naitoh, T. Kawai, J. Vac. Sci. Technol. B 17 (1999) 1941. [22] T. Uchihashi, M. Tanigawa, M. Ashino, Y. Sugawara, K. Yokoyama, S. Morita, M. Ishikawa, Langmuir 16 (2000) 1349.
[23] T. Uchihashi, N. Choi, M. Tanigawa, M. Ashino, Y. Sugawara, H. Nishijima, S. Akita, Y. Nakayama, H. Tokumoto, K. Yokoyama, S. Morita, M. Ishikawa, Jpn. J. Appl. Phys. 39 (2000) L887. [24] B. Geisler, F. Noll, N. Hampp, Scanning 22 (2000) 7. [25] T. Arai, M. Tomitori, M. Saito, E. Tamiya, Appl. Surf. Sci. 188 (2002) 474. [26] Y. Martin, D.W. Abraham, H.K. Wickramasinghe, Appl. Phys. Lett. 52 (1988) 1103. [27] M. Nonnenmacher, M.P. O’Boyle, H.K. Wickramasinghe, Appl. Phys. Lett. 58 (1991) 2921. [28] M. Yasutake, D. Aoki, M. Fujihira, Thin Solid Films 273 (1996) 279. [29] J.B. Brzoska, N. Shahidzadeh, F. Rondelez, Nature 360 (1992) 719. [30] M. Fujihira, Y. Okabe, Y. Tani, M. Furugori, U. Akiba, Ultramicroscopy 82 (2000) 181. [31] S.N. Magonov, V. Elings, M.-H. Whangbo, Surf. Sci. 375 (1997) L385. [32] A. Noy, C.H. Sanders, D.V. Vezenov, S.S. Wong, C.M. Lieber, Langmuir 14 (1998) 1508. [33] K. Sasaki, Y. Koike, H. Azehara, H. Hokari, M. Fujihira, Appl. Phys. A 66 (1998) S1275. [34] K.J. Kwak, H. Kudo, M. Fujihira, Ultramicroscopy, in press. [35] M. Fujihira, Ann. Rev. Mater. Sci. 29 (1999) 353. [36] H. Tanaka, C. Hamai, T. Kanno, T. Kawai, Surf. Sci. 432 (1999) L611.
Observation of stretched single DNA molecules by Kelvin probe force microscopy K.J. Kwak, S. Yoda, M. Fujihira* Department of Biomolecular Engineering, Tokyo Institute of Technology, 4259 Nagatsuta, Midori-ku, Yokohama 226-8501, Japan
Abstract Lambda bacteriophage deoxyribonucleic acid (DNA) molecules stained with YOYO-1 were deposited and stretched on chemically modified Si(1 0 0) wafer and cover glass substrates. The Si(1 0 0) wafer with a natural oxide layer and the cover glass surface were first chemically modified by vapor phase chemisorption with mixed two organosilanes. DNA molecules were then aligned on the substrate surfaces by chemical and physical adsorption from an aqueous solution during molecular combing. The aligned DNA molecules were observed by Kelvin probe force microscopy and intermittent contact-mode atomic force microscopy (AFM) as well as fluorescence microscopy. Interaction between the stretched DNA molecules and the chemically modified Si(1 0 0) substrate surface was examined using these AFMs. # 2002 Elsevier Science B.V. All rights reserved. PACS: 07.79.Lh; 61.16.Ch; 87.64.Dz Keywords: Stretched DNA; Molecular combing; Kelvin probe force microscopy; Atomic force microscopy
1. Introduction The study of interactions of biological molecules with a solid surface is one of the most important issues of atomic force microscopy (AFM) for its application to biological samples [1,2]. A cleaved mica surface as a flat substrate has been extensively used to prepare such a biological sample as deoxyribonucleic acid (DNA) [3–8]. When DNA molecules are bound to a substrate surface, it is desirable that DNA molecules can still hybridize with their complementary strands or interact with proteins in the bound forms. The binding of DNA molecules to a chemically treated mica surface can be *
Corresponding author. Tel.: þ81-45-924-5784; fax: þ81-45-924-5817. E-mail address: [email protected] (M. Fujihira).
strong enough to lose such biological activities [9–11]. Additionally, alignment of the immobilized DNA molecules on the flat substrate is another interesting issue of AFM [12–18]. A reproducible process called a molecular combing method has been demonstrated for alignment of DNA molecules [12,13]. The DNA strands can be uniformly stretched on the chemically modified surface with a CH2¼CH-terminated group by a hydrodynamic force of a receding meniscus. Recently, noncontact-mode AFM (nc-AFM) in ultrahigh vacuum has been demonstrated to image the DNA molecules with high accuracy and high resolution [19–21]. This technique has been extended further to the high resolution imaging DNA molecules [22–24]. However, there has been little report on interactions between DNA molecules and substrate surfaces [25].
0169-4332/02/$ – see front matter # 2002 Elsevier Science B.V. All rights reserved. doi:10.1016/S0169-4332(02)01482-4
74
K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
In this work, we have demonstrated nc-AFM to image the stretched DNA molecules in air. A cantilever with an Au-coated tip was chemically modified by an organothiol compound and applied to stable observation of DNA by Kelvin probe force microscopy (KFM) [26–28]. The stretched DNA strands were deposited on mixed silane monolayers of CH2¼CH- and NH2-terminated silanes chemisorbed on oxide substrate surfaces. It is important to prepare samples having the optimized surface density of DNA strands for the AFM study.
N.A. oil immersion lens. All SPM measurements including intermittent contact (i.e. TappingTM mode) AFM and KFM were performed using an SPA-400 AFM unit and an SPI-3800 control station (Seiko Instruments). Cantilevers with an Au-coated tip having a normal spring constant of 1.6 N m1 and resonance frequency of 23–25 kHz were directly used after cleaning the tip [30] for intermittent contact AFM or chemically modified with hexadecanethiol (HDT) for KFM.
3. Results and discussion 2. Experimental 3.1. Fluorescence microscopy 2.1. Preparation of stretched DNA molecules Organosilane compounds for modification of oxide substrate surfaces were purchased from Aldrich Chemical Co. and further purified by distillation. 7-Octen1-yl (CH2¼CH-terminal) and 3-(2-aminoethyl)aminopropyl (NH2-terminal) trimethoxysilanes were used as silanization reagents. Si(1 0 0) wafers (n-type high doped; Nilaco Co.) and glass coverslips from Matsunami Glass Co. were used as the oxide substrates for AFM and/or fluorescence microscopy. Lambda bacteriophage DNA molecules (48,502 bp; Takara Bio Inc., Japan) were stained with YOYO-1 at a ratio of DNA base pairs to the YOYO-1 molecules of 5:1. The Si(1 0 0) wafers were cleaned and oxidized using a UV-ozone cleaner (Nippon Laser and Electronics Laboratory) for 1 h and/or piranha solution treatments [29]. Glass coverslips were cleaned by the piranha solution treatments. The silanized substrate surfaces were prepared by vapor phase chemisorption with a mixed two-component liquid of the organosilane compounds with a 1:10 (v/v) ratio of NH2- to CH2¼CH-terminated silanes. The silanized substrates were dipped into a TE buffer (10 mM Tris, 1 mM EDTA) containing the stained DNA molecules diluted to 0.45 mg ml1. After 5 min of incubation, the silanized substrates were lifted up with a mechanical apparatus at a constant speed of 300 mm s1. 2.2. Imaging of DNA molecules Fluorescence microscopy was performed using a Nikon microscope (Diaphot 300) with a 100/1.30
The adsorption of the lambda bacteriophage DNA molecules on the cover glass substrate by the molecular combing method [13] was easily examined using the fluorescence microscope as shown in Fig. 1. The fluorescent dye YOYO-1 showed an excitation maximum at 491 nm and an emission maximum at 509 nm when binding to DNA molecules. Fluorescence microscopy could be used to confirm the optimized surface density of the adsorbed DNA strands for AFM imaging. Here, the two-component system in the modification of substrates was carried out with the mixed gas consisting of NH2- and CH2¼CH-terminated silanes. The increase in the surface density of DNA shown in Fig. 1 in comparison with that on a CH2¼CH-terminated silane monolayer (not shown here) indicates that the DNA strands are negatively charged in aqueous solution at neutral pH, and thus the DNA molecules would be electrostatically attracted to positively charged NH2-terminals [9,11]. 3.2. AFM measurements The stretched DNA strands were observed by intermittent contact AFM with an Au-coated tip cleaned by the UV-ozone treatment. Fig. 2(a) and (b) show a topographic and a phase shift image, respectively, obtained in 45.5% relative humidity (RH) air. Both topography and phase shift on the DNA strands show higher heights and larger phase shifts (i.e. greater phase-lag) than those of the substrate surface, respectively. The greater phase-lag can be attributed to the surface water layer on the DNA strands, which
K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
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Fig. 1. A fluorescence microscopic image of the DNA strands deposited and stretched by the dynamic molecular combing method on the twocomponent NH2- and CH2¼CH-terminated substrate surface.
enhanced the adhesive force between the tip and the strands [31–33]. The higher adhesive force on the strands than on the hydrophobic substrate surface was shown previously by pulsed-force-mode AFM [34].
The present surface monolayer prepared by vapor phase chemisorption with a 1:10 (v/v) mixture seems to contain ca. 105 mole fraction of NH2-terminals from the previous results with a 1:1 (v/v) mixture [34].
Fig. 2. 3 mm 3 mm (a) topographic and (b) phase images simultaneously observed by intermittent contact AFM. Two images were acquired in 45.5% RH air.
76 K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78 Fig. 3. 1:5 mm 1:5 mm (a) topographic image observed by intermittent contact AFM and (b) topographic and (c) CPD images simultaneously observed by KFM. Three images were acquired in 48.9% RH air with an HDT-modified Au tip. The profiles of (d) and (e) along the solid lines in (a) and (b), respectively.
K.J. Kwak et al. / Applied Surface Science 210 (2003) 73–78
In higher humidity than 60% RH, this enhanced adhesive force resulted in cleaving the DNA strands to their fragments (not shown here). For getting the higher spatial resolution of the contact potential difference (CPD), we used the method called KFM [28]. Fig. 3(a) shows topographic image observed by intermittent contact AFM. Fig. 3(b) and (c) show a topographic and a CPD image simultaneously observed by KFM, respectively, scanned over the same area as that in Fig. 3(a). Here, the CPD of the sample was measured against an HDTmodified Au tip. Three images were acquired in 48.9% RH air. The a.c. modulation for the KFM measurements was applied to the sample substrate with a driving a.c. frequency of 22 kHz and amplitude of 5 V (peak to peak). The CPD distributions on the silane-modified Si(1 0 0) surface ranged from 0.34 to 0.40 V as shown in Fig. 3(c). This inhomogeneity of the CPD observed on the substrate surface may be attributed to the mixed monolayer of NH2- and CH2¼CH-terminated silanes. If the mole fraction of NH2-terminated silane is ca. 105 as described above and the cross-sectional area of an alkyl chain is 0.2 nm2, the surface density of NH2-terminated silane will be ca. 50 molecules mm2. The number of the bright spots observed in Fig. 3(c) is in this order. However, origin of the increase in CPD due to NH2terminals has not been clarified yet. If the NH2-terminals are ionized, we will expect the decrease in CPD, while if the NH2-terminals are not ionized, the increase in CPD will be expected due to permanent dipoles of the NH2-terminals [35]. More interestingly, three bright lines in Fig. 3(c) corresponding well with DNA strands observed in Fig. 3(a) and (b) seem to be pinned by the several bright spots on the substrate surface. This suggests that NH2-terminal groups make ion complexes with phosphate anions on DNA. The bright lines indicate that the stretched DNA strands have the higher CPD than that on the substrate surface. The cylindrical symmetry of DNA suggests that the CPD would not be affected significantly by DNA adsorption on the substrate. The result suggests that the dipole moments formed between the surface water layer and the negatively charged DNA strands have a direction from the substrate toward air. Fig. 3(d) and (e) show the profiles along the solid lines in Fig. 3(a) and (b), respectively. The stretched DNA strands on the substrate surface have a width of
77
34 nm and a height of 0.7 nm as shown in Fig. 3(d), which are comparable to those in Fig. 3(e). This typical result indicates that the spatial resolution of the DNA strands was determined mainly by the size of the tip end [3,5]. The smaller height than 2.4 nm of van der Waals radii of DNA [23] also agrees with those measured recently by nc-AFM with the frequency modulation technique [22,23,25] and by STM [36].
4. Conclusions The stretched DNA strands on the silanized substrate surfaces were observed by KFM and intermittent contact AFM as well as fluorescence microscopy for the study of interactions between the DNA molecules and the substrate surfaces. The cantilever with the HDT-modified Au tip was applied to the stable observation of the DNA molecules by KFM.
Acknowledgements This work was supported by a Grant-in-Aid for Creative Scientific Research on ‘‘Devices on molecular and DNA levels’’ (No. 13GS0017) from the Ministry of Education, Science, Sports, and Culture of Japan.
References [1] R.J. Colton, D.R. Baselt, Y.F. Dufrene, J.B.D. Green, G.U. Lee, Curr. Opin. Chem. Biol. 1 (1997) 370. [2] H.G. Hansma, Ann. Rev. Phys. Chem. 52 (2001) 71. [3] J. Vesenka, M. Guthold, C.L. Tang, D. Keller, E. Delaine, C. Bustamante, Ultramicroscopy 42–44 (1992) 1243. [4] T. Thundat, D.P. Allison, R.J. Warmack, G.M. Brown, K.B. Jacobson, J.J. Schrick, T.L. Ferrell, Scan. Micros. 6 (1992) 911. [5] C. Bustamante, J. Vesenka, C.L. Tang, W. Rees, M. Guthold, R. Keller, Biochemistry 31 (1992) 22. [6] H.G. Hansma, J. Vesenka, C. Siegerist, G. Kelderman, H. Morrett, R.L. Sinsheimer, V. Elings, C. Bustamante, P.K. Hansma, Science 256 (1992) 1180. [7] Y.L. Lyubchenko, A.A. Gall, L.S. Shlyakhtenko, R.E. Harrington, B.L. Jacobs, P.I. Oden, S.M. Lindsay, J. Biomol. Struct. Dyn. 10 (1992) 589. [8] Y.L. Lyubchenko, L.S. Shlyakhtenko, R.E. Harrington, P.I. Oden, S.M. Lindsay, Proc. Natl. Acad. Sci. USA 90 (1993) 2137. [9] M. Bezanilla, S. Manne, D.E. Laney, Y.L. Lyubchenko, H.G. Hansma, Langmuir 11 (1995) 655.
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[10] H.G. Hansma, I. Revenko, K. Kim, D.E. Laney, Nucl. Acids Res. 24 (1996) 713. [11] V. Balladur, A. Theretz, B. Mandrand, J. Coll. Interf. Sci. 194 (1997) 408. [12] A. Bensimon, A. Simon, A. Chiffaudel, V. Croquette, F. Heslot, D. Bensimon, Science 265 (1994) 2096. [13] X. Michalet, R. Ekong, F. Fougerousse, S. Rousseaux, C. Schurra, N. Hornigold, M. van Slegtenhorst, J. Wolfe, S. Povey, J.S. Beckmann, A. Bensimon, Science 277 (1997) 1518. [14] Z.-Q. Ouyang, J. Hu, S.-F. Chen, J.-L. Sun, M.-Q. Li, J. Vac. Sci. Technol. B 15 (1997) 1385. [15] W. Wang, J. Lin, D.C. Schwartz, Biophys. J. 75 (1998) 513. [16] H. Yokota, F. Johnson, H. Lu, R.M. Robinson, A.M. Belu, M.D. Garrison, B.D. Ratner, B.J. Trask, D.L. Miller, Nucl. Acids Res. 25 (1997) 1064. [17] H. Yokota, J. Sunwoo, M. Sarikaya, G. van den Engh, R. Aebersold, Anal. Chem. 71 (1999) 4418. [18] J.Y. Ye, K. Umemura, M. Ishikawa, R. Kuroda, Anal. Biochem. 281 (2000) 21. [19] Y. Maeda, T. Matsumoto, T. Kawai, Appl. Surf. Sci. 140 (1999) 400. [20] Y. Maeda, T. Matsumoto, H. Tanaka, T. Kawai, Jpn. J. Appl. Phys. 38 (1999) L1211. [21] T. Matsumoto, Y. Maeda, Y. Naitoh, T. Kawai, J. Vac. Sci. Technol. B 17 (1999) 1941. [22] T. Uchihashi, M. Tanigawa, M. Ashino, Y. Sugawara, K. Yokoyama, S. Morita, M. Ishikawa, Langmuir 16 (2000) 1349.
[23] T. Uchihashi, N. Choi, M. Tanigawa, M. Ashino, Y. Sugawara, H. Nishijima, S. Akita, Y. Nakayama, H. Tokumoto, K. Yokoyama, S. Morita, M. Ishikawa, Jpn. J. Appl. Phys. 39 (2000) L887. [24] B. Geisler, F. Noll, N. Hampp, Scanning 22 (2000) 7. [25] T. Arai, M. Tomitori, M. Saito, E. Tamiya, Appl. Surf. Sci. 188 (2002) 474. [26] Y. Martin, D.W. Abraham, H.K. Wickramasinghe, Appl. Phys. Lett. 52 (1988) 1103. [27] M. Nonnenmacher, M.P. O’Boyle, H.K. Wickramasinghe, Appl. Phys. Lett. 58 (1991) 2921. [28] M. Yasutake, D. Aoki, M. Fujihira, Thin Solid Films 273 (1996) 279. [29] J.B. Brzoska, N. Shahidzadeh, F. Rondelez, Nature 360 (1992) 719. [30] M. Fujihira, Y. Okabe, Y. Tani, M. Furugori, U. Akiba, Ultramicroscopy 82 (2000) 181. [31] S.N. Magonov, V. Elings, M.-H. Whangbo, Surf. Sci. 375 (1997) L385. [32] A. Noy, C.H. Sanders, D.V. Vezenov, S.S. Wong, C.M. Lieber, Langmuir 14 (1998) 1508. [33] K. Sasaki, Y. Koike, H. Azehara, H. Hokari, M. Fujihira, Appl. Phys. A 66 (1998) S1275. [34] K.J. Kwak, H. Kudo, M. Fujihira, Ultramicroscopy, in press. [35] M. Fujihira, Ann. Rev. Mater. Sci. 29 (1999) 353. [36] H. Tanaka, C. Hamai, T. Kanno, T. Kawai, Surf. Sci. 432 (1999) L611.