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Langmuir 2007, 23, 8480-8484
Influence of Salt on Colloidal Lithography of Albumin D. L. Geng, Y. H. Miao, and L. E. Helseth* School of Physical and Mathematical Sciences, DiVision of Physics and Applied Physics, Nanyang Technological UniVersity, Singapore ReceiVed December 18, 2006. In Final Form: April 30, 2007 We investigate the influence of salt on colloidal lithography of biomolecular patterns. Albumin labeled with fluorescein isothiocyanate (FITC) was adsorbed on polyelectrolyte-coated glass substrates covered by negatively charged colloids using fluorescence microscopy. After removing the colloids, a well-defined albumin pattern remains, and we study how the pattern changes upon adding salt to the protein solution. The proposed method is simple and cheap and can be used to create stable one- and two-dimensional biomolecular arrays.
1. Introduction
2. Experimental Section
A number of structures can now be routinely reproduced using various self-assembly techniques.1-10 Capillary, electric, and magnetic forces have proven particularly useful for assembling structures ranging from nanometer to millimeter scale on both solid-liquid and air-water interfaces. To this end, colloidal lithography is a promising technique which has been used to pattern gold films, polymers, and protein arrays.11-14 Colloidal lithography is able to create, e.g., microscopic rings and dots and therefore has a great potential as a technique for chemical patterning.11 Here, we show how such patterns can be controlled by altering the salt concentration of the liquid solution to be deposited on the colloid-covered substrate. In particular, we investigated how albumin adsorbed to a substrate coated with the positively charged polymer poly(allylamine hydrochloride) (PAH) and covered by negatively charged colloids. Albumin adsorbed to the polyelectrolytes but was obstructed by the presence of the negatively charged colloids. Upon removing the colloids using ultrasound, it was observed that well-defined patterns could be created on otherwise uniformly coated substrates. We found that, upon regulating the salt concentration of the solution containing the albumin, we could also control the resulting fluorescence pattern. The technique presented here can be applied to different biomolecules which can be immobilized by polyelectrolyte-coated substrates, and therefore offers a simple and cost-effective method for assembling biomolecular patterns without the use of expensive lithographic techniques.
Poly(allylamine hydrochloride) (PAH, MW ∼ 70 000), FITCalbumin (prod. no. A9771; albumin of MW 66 000 labeled with fluorescein isothiocyanate of MW 389.4) and salt (NaCl, MW 58.4) were purchased from Sigma-Aldrich and used as received. Carboxylmodified paramagnetic polystyrene beads of radius a ) 1.4 µm (Dynabeads M270) were purchased from Dynal. The colloids were used as received and diluted to a density of approximately 107 beads/ mL. We used glass slides purchased from Electron Microscopy Sciences, partially covered with Teflon and with each containing three 8-mm-diameter (uncovered) glass wells as substrates; see Figure 1. The slides were first rinsed with 1 M NaOH (30 min), followed by a water wash, a bath in 1 M HCl (10 min), and finally a thorough rinse in water; see also ref 15 for more details on this procedure. The procedure for lithographic patterning is shown in Figure 2. First, the glass substrate well was covered with PAH solution for 5 min, followed by a thorough rinse in water. This left a thin layer of positively charged PAH on the glass slide (step 1). About 30 µL of Dynabeads M270 solution was deposited on the horizontally aligned substrate. In less than 5 min, the negatively charged colloids will eventually sink to the glass surface due to gravity where they are immobilized by the positively charged PAH-coated substrate. After waiting for 5 min, we washed the substrate thoroughly in water (step 2). The upper surfaces of the colloids are negatively charged, while the PAH-coated glass surface is positively charged, thus rendering a heterogeneously charged surface suitable for patterning of electrically charged molecules. Such a technique was reported in refs 16 and 17 as a method for creating colloidal structures, but we will use it here to facilitate colloidal lithography. Here, we used negatively charged FITC-albumin, because it irreversibly attaches easily to the positively charged PAH-coated substrate. We deposited 30 µL of 1 g/L FITC-albumin (10 mg FITC-albumin in 10 mL deionized water) in the glass well, waited for about 5 min, and gave the glass slide a thorough rinse with water. This procedure left a thin layer of FITC-albumin on the PAH-coated areas not covered by colloids (step 3). We repeated this step (step 3) with FITCalbumin dissolved in different NaCl concentrations ranging from 0 to 0.1 M NaCl. In the final step (step 4), we removed the paramagnetic colloids by sonication (5-10 min), thus leaving areas not covered by albumin. The colloids could not be removed by a strong magnetic field gradient due to the strong interaction with the PAH-coated
* Corresponding author. E-mail:
[email protected]. (1) Pieranski, P. Phys. ReV. Lett. 1980, 45, 569. (2) Karakurt, I.; Leiderer, P.; Boneberg, J. Langmuir 2006, 22, 2415. (3) Masuda, Y.; Itoh, T.; Koumoto, K. Langmuir 2005, 21, 4478. (4) Truskett, V. N.; Stebe, K. J. Langmuir 2003, 19, 8271. (5) Tsuji, S.; Kawaguchi, H. Langmuir 2005, 21, 2434. (6) Tsuji, S.; Kawaguchi, H. Langmuir 2005, 21, 8439. (7) Helseth, L. E. Langmuir 2005, 21, 7276. (8) Fan, F.; Stebe, K. J. Langmuir 2004, 20, 3062. (9) Abe, M.; Yamamoto, A.; Orita, M.; Ohkubo, T.; Sakai, H.; Momozawa, N. Langmuir 2004, 20, 7021. (10) Abe, M.; Orita, M.; Yamazaki, H.; Tsukamoto, S.; Teshima, Y.; Sakai, T.; Ohkubo, T.; Momozawa, N.; Sakai, H. Langmuir 2004, 20, 5046. (11) Boneberg, J.; Burmeister, F.; Schafle, C.; Leiderer, P.; Reim, D.; Fery, A.; Herminghaus, S. Langmuir 1997, 13, 7080. (12) Burmeister, F.; Schafle, C.; Matthes, T.; Bohmisch, M.; Boneberg, J.; Leiderer, P. Langmuir 1997, 13, 2983. (13) Cai, Y. G.; Ocko, B. M. Langmuir 2005, 21, 9274. (14) Snyder, C. E.; Yake, A. M.; Feick, J. D.; Velegol, D. Langmuir 2005, 21, 4813.
(15) Hau, W. L. W.; Trau, D. W.; Sucher, N. J.; Wong, M.; Zohar, Y. J. Micromech. Microeng. 2003, 13, 272. (16) Chen, K. M.; Jiang, X.; Kimerling, L. C.; Hammond, P. T. Langmuir 2000, 16, 7825. (17) Lyles, B. F.; Terrot, M. S.; Hammond, P. T.; Gast, A. P. Langmuir 2004, 20, 3028.
10.1021/la063652v CCC: $37.00 © 2007 American Chemical Society Published on Web 07/06/2007
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Figure 1. Image showing the glass slides partially covered with Teflon and with each containing three uncovered glass wells as substrates. The green light is added to illustrate the fluorescent emission. Figure 3. The fraction of colloids sticking to the substrate as a function of PAH concentration.
Figure 2. Schematic drawing of the basic steps for the patterning process. glass slide. The dye-labeled molecules were observed with an inverted fluorescence microscope (Nikon TE2000U) equipped with proper wavelength filters for detection of FITC. We used a 60× objective with numerical aperture NA ) 0.7 and a Nikon DS-5Mc-US cooled CCD camera for imaging of the fluorescent patterns. The diffractionlimited resolution of the microscope is therefore about 370 nm. The pixel size corresponds to about 320 nm, i.e., smaller than the diffraction limit.
3. Results In order to optimize step 2 in Figure 2, we investigated the fraction θ of colloids sticking to the substrate as a function of the PAH concentration. Here, θ was obtained by counting the number of colloids on the substrate before and after washing it with water, and thereafter calculating the ratio between the two. The result is shown in Figure 3, which demonstrates that a PAH solution of g10 g/L PAH will allow us to immobilize most of the colloids onto the glass slide.
Figure 4. Fluorescent patterns on a PAH-coated substrate observed after removing the colloid. The FITC-albumin was dissolved in 0 M (a) and 10-2 M NaCl (b), respectively. The contrast of the images has been enhanced for clarity.
Figure 4 shows typical images of the albumin patterns that could be observed with the fluorescence microscope when using FITC-albumin in 0 M (a) and 10-2 M NaCl (b), respectively. The areas not covered by FITC-albumin are dark, whereas the covered areas are bright green. It is seen that characteristic rings show up for 0 M NaCl, where the bright ring diameter is comparable to the diameter of the colloids with an average value (2.5 ( 0.4) µm. These rings disappear when the salt concentration is 10-2 M, thus leaving only a dark hole. In the areas not originally covered by colloids (see Figure 4), we observed that the fluorescence intensity did not change with the salt concentration, thus suggesting that the amount of albumin adsorbed here was independent of the ions in the solution. Using the software ImageJ, we recorded the intensity profiles versus position of the fluorescent patterns. Two such examples are show in Figure 5, where the black line corresponds to a pattern formed using 0 M NaCl while the red line corresponds to a pattern formed in 10-2 M NaCl (see also Figure 4a,b). An advantage of using fluorescent microscopy to detect the albumin patterns is that the observed intensity is proportional to
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Figure 5. Intensity profiles of the fluorescent patterns corresponding to FITC-albumin dissolved in 0 M (a) and 10-2 M NaCl (b) in Figure 4, respectively.
the number of FITC-albumin molecules, since the FITC molecules emit incoherent light which can be summed up accordingly.18,19 To this end, the fluorescent patterns observed in Figure 4a suggest that many albumin molecules concentrate here to form a bright ring near the edge of the charged, spherical colloid. On the other hand, there are fewer albumin molecules directly under the colloid and far away from it. Thus, it is plausible that the molecules are trapped due to an electrostatic field gradient. In order to investigate whether electrostatic forces are responsible for the bright ring, we used FITC-albumin dissolved in solutions of increasing NaCl concentrations. We observed that for salt concentrations above 5 × 10-3 M the bright ring disappeared completely (see also Figure 4b). It should be mentioned that, in order to display a clear image, we have enhanced the brightness and contrast of Figure 4b, and the gray levels seen should therefore not be compared to Figure 4a. A measure of the ratio between the number of molecules near the boundary of the colloid to the number of molecules directly under the colloids (dark region) can be obtained by defining the contrast as the difference between peak intensity (Ip) and center intensity (Ic) divided by the average of the two, i.e.,
Contrast ) 2
Ip - I c Ip + Ic
(1)
Figure 6 shows the contrast as a function of salt concentration ion concentration c. It is seen that the contrast initially decreases strongly with salt concentration, thus indicating that fewer and fewer albumin molecules are trapped in a ring formation as more salt is introduced. At a salt concentration of 5 × 10-2 M, the fluorescent patterns were seen to disappear completely, thus suggesting that the colloids no longer were able to obstruct the deposition of albumin.
4. Discussion According to refs 20-23 and references contained therein, albumin has an isoelectric point of about 5 and has a net charge (18) Herman, B.; Tanke, H. J. Fluorescence microscopy, 2nd ed., Springer: London, 1998. (19) Born, M.; Wolf, E. Principles of Optics, 6th ed.; Cambridge University Press: Cambridge, U.K., 1980. (20) Carter, D. C.; Ho, J. X. AdV. Protein Chem. 1994, 45, 153. (21) Reynolds, J. A.; Tanford, C. Proc. Natl. Acad. Sci. U.S.A. 1970, 66, 1002. (22) Shweitzer, B.; Zanette, D.; Itri, R. J. Colloid Interface Sci. 2004, 277, 285.
Figure 6. Contrast as a function of NaCl concentration. Each measurement is an average over typically 30 patterns, with the error bars denoting the corresponding standard deviation.
of about -20 electrons at pH ∼7. The interaction between albumin and polyelectrolytes has been investigated in a number of studies, where it has been shown that albumin adsorbs to both positively and negatively charged surfaces.24-28 Thus, trace amounts of albumin molecules will also adsorb to the paramagnetic colloids covered with negatively charged carboxylic acid groups as well, in agreement with what we observed in our fluorescence studies. Thus, care should be taken, since albumin has both hydrophobic and hydrophilic sites, despite its net negative charge. Here, we will only be interested in how the albumin molecules distribute spatially on the PAH-coated substrate. For pH ∼7, the molecular dimensions are 5 × 5 × 9 nm3, and it should be possible for this molecule to penetrate far under the spherical polymer spheres of diameter 2.8 µm if they were not influenced by any external forces. From the geometry of the system seen in Figure 7, we know that a spherical molecule of diameter D will be geometrically obstructed at a distance r ) xx2+y2 approximately given by (a - D)2 + r2 ) a2, which in the case of D , a gives r ≈ x2aD.13,14 A molecule of size 5 nm should therefore in principle be able to move to a position r ≈ 118 nm. It should be noted that PAH is a positively charged polymer of molecular weight of about 70 000. Each of the about 750 monomers has a molecular weight of 93 and a typical length of 1 nm, with a calculated radius of gyration of about 50 nm.28 Once the positively charged polymer collapses on the negatively charged surface, it will comprise a thin film of nanometer thickness; see ref 15. However, it is reasonable to believe that the charge density is larger for small r, since the PAH chains will be drawn toward the carboxylic groups coating the colloids, thus extending slightly above the substrate as seen in Figure 7. In the region where this happens, there is a larger density of positively charged ions, which may attract and stop the motion of negatively charged biomolecules. (23) Vilker, V. L.; Colton, C. K.; Smith, K. A. J. Colloid Interface Sci. 1981, 79, 548. (24) Ladam, G.; Schaaf, P.; Cuisinier, F. J. G.; Decher, G.; Voegel, J.-C. Langmuir 2001, 17, 878. (25) Carlsson, F.; Hyltner, E.; Arnebrant, T.; Malmsten, M.; Linse, P. J. Phys. Chem. B. 2004, 108, 9871. (26) Seyrek, E.; Dubin, P. L.; Tribet, C.; Gamble, E. A. Biomacromolecules 2003, 4, 273. (27) Salloum, D. S.; Schlenoff, J. B. Biomacromolecules 2004, 5, 1089. (28) Adamczyk, Z.; Zembala, M.; Warszynski, P.; Jachimska, B. Langmuir 2004, 20, 10517.
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Figure 7. Schematic drawing of the electrostatic trapping geometry, where an electric field is set up between the positively charged PAH-coated glass substrate and the negatively charged polymer sphere. Also drawn is the structure of PAH (on the left side of the picture).
Figure 8. Colloidal chains of paramagnetic polystyrene spheres on a substrate (a). After following the steps of Figure 1, one can obtain linear FITC-albumin patterns corresponding to the original colloidal assembly (b).
In an electrolyte, the Debye length plays a natural role and is given by
1 lD ) ) κ
x
kBT 2e2n
(2)
where is the permittivity of water, kB is Boltzmann’s constant, T is the temperature, e is the electron charge, and n is the density of ions. The Debye length tells us how effectively the charges of the albumin, PAH, and COOH- are screened and is seen to decrease with increasing salt concentration, and is smaller than 100 nm at salt concentrations exceeding 10-5 M. Let us now assume that the rings occur due to capture of negatively charged albumin molecules by positively charged PAH molecules. Then, an increase in salt concentration would screen the charges and allow the albumin to penetrate further under the colloids before being captured. We found that the bright rings disappeared at salt concentrations exceeding 5 × 10-3 M, and it is therefore
Figure 9. Two-dimensional colloidal arrays of polystyrene spheres on a substrate (a). After following the steps of Figure 1, one can obtain two-dimensional FITC-albumin patterns corresponding to the original colloidal assembly (b).
reasonable to assume that this is due to the fact that electrostatic attraction has been reduced considerably due to electrostatic screening. Since the dimension of the albumin molecule is about 5 nm, it is allowed to move to a distance r ≈ 118 nm before it is geometrically obstructed. The fact that the contrast goes to zero when the salt concentration increases above 5 × 10-2 M suggests that the colloid is not in contact with the glass slide, but is lifted a small distance above it due to the PAH coating. Thus, for larger salt concentrations the electrostatic attraction between albumin and PAH is screened out, and the albumin molecules can move more or less freely under the colloids. The technique studied here may have applications for creation of heterogeneously charged surfaces and structuring of charged molecules.Because the polyelectrolyte immobilizes negatively
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charged colloids, we found that it was rather straightforward to create one- and two-dimensional arrays. In the presence of an external magnetic field H ≈ 3 kA/m obtained by using a small permanent magnet placed next to the glass slide during deposition of the colloids, we found that chains of paramagnetic colloids could be immobilized on the glass surface, as is demonstrated in Figure 8a. By following the procedure of Figure 2, we could then create corresponding one-dimensional fluorescent patterns as seen in Figure 8b. In addition to using the paramagnetic colloids, other charged colloids could also be used for creating twodimensional arrays. One example is seen in Figure 9a, where a two-dimensional array of negatively charged polystyrene colloids has been dried on a PAH-coated substrate. After adding FITCalbumin, washing, and then sonicating the sample, we observed the two-dimensional fluorescent pattern seen in Figure 9b. We stored the substrates in darkness under dry conditions for several
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days without observing any changes in the fluorescent patterns, thus suggesting that the patterns are stable.
5. Conclusion We have demonstrated that colloidal lithography on polyelectrolyte-coated substrates can be used to create tunable patterns. We also found that one- and two-dimensional patterns can be created in a simple way. The technique can be applied to other biomolecules which can be assembled on polyelectrolyte-coated substrates, and is therefore of interest as a method for creating chemically patterned substrates for use in, e.g., detection of biomolecules or for self-assembly of colloidal structures. Acknowledgment. This work was supported by URC grant RG32/05. LA063652V
