Langmuir 2007, 23, 8015-8020
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Real-Time Quantitative Imaging of Submolecular Layers Dominique Ausserre´*,† and Refahi Abou Khachfe‡ Laboratoire de Physique de l’Etat Condense´ (UMR 6087), UniVersite´ du Maine, CNRS, Route de LaVal, 72000 Le Mans, France, and 21, rue Marcel Carne´ , 72000 Le Mans, France ReceiVed December 14, 2006. In Final Form: March 12, 2007 Using a recent optical contrast method, real-time and quantitative imaging of submolecular layers was performed with the help of a simple optical microscope. The measuring technique is exposed and documented by three examples. In particular, it allowed label-free detection of peptide-antibody binding interactions with 50 pg/mm2 sensitivity while keeping full optical lateral resolution.
Topographic images shown in the present paper were obtained not using an atomic force microscope (AFM) but using a novel optical technique that can be implemented at low cost in any laboratory. It is an extension of a new contrast technique for white-light optical microscopy which was introduced recently1 and which provides real-time imaging of ultrathin films with subnanometer sensitivity. It is based on the use of contrastamplifying substrates especially designed for having antireflecting properties when observed between cross polarizers, thus named “AR-X-POL” surfaces. Equivalently, these substrates can be described as nondepolarizing mirrors, since they do not alter the initial polarization of a light beam upon reflection. They are defined by the very simple relationship rp + rs ) 0 between the two Fresnel coefficients of their surface. This condition can be satisfied on almost any material by adding a λ/4 layer with refractive index n1 given, in a low aperture approximation, by the relationship
(
)
1 1 1 1 ) + 2 2 2 2 n1 n0 n2
where n0 and n2 hold for ambient and solid indices. When used as sample stages, these substrates provide better extinction for all extinction methods in polarized light. Then any trace deposited on the surface makes the above condition vanish so that it appears bright on a dark background. This technique already proved to be particularly powerful when combined with differential interference contrast. In this mode, it made possible to visualize for the first time a single label-free DNA molecule1 with the help of a standard wide-field commercial microscopesno laser, no scanning. Real-time imaging of molecular films and direct reading of label-free biochips with full optical lateral resolution are other promising applications. However, in addition to sensitivity, these applications need the ability of the technique to deliver quantitative measurements. The aim of the present paper is to describe a powerful instrument based on this contrast technique that delivers local thickness measurement with molecular sensitivity and that keeps the simplicity of use of a standard white-light optical microscope. Since polarization change upon reflection is the heart of ellipsometry,2 we will refer to the technique as “surface enhanced * Corresponding author. E-mail:
[email protected] † Universite ´ du Maine, CNRS. ‡ 21, rue Marcel Carne ´ , 72000 Le Mans, France. (1) Ausserre´, D.; Valignat, M.-P. Nano Lett. 2006, 6, 1384-1388. (2) Azzam, R. M. A.; Bashara, N. M. Ellipsometry and Polarized Light; NorthHolland: New York, 1977.
ellipsometric contrast” (SEEC). The possibilities of the technique will be illustrated with three prevailing examples: vicinal surfaces of supported phospholipid multilayers, crystallization of ultrathin polymer films, and label-free reading of peptide-protein biochips. The experiments reported below were carried out using a basic polarization microscope (DMR HC, Leica) equipped with a xenon white lamp and a standard video camera (SONY DXC390P). Because a microscope is imperfect and because its exact setting is difficult to reproduce, we avoided direct comparison of experiment and theory. Our route was instead very pragmatic. It is model-independent and can even be transposed to a couple of other sensitive imaging techniques, such as basic wide-field microscopy using nonpolarized light on classical antireflecting surfaces.3 It does not require proceeding with any critical tuning of the instrument. It is based on the comparison of the signal obtained with the sample under study and the signal obtained with a reference sample. In our case the reference sample is an AR-X-POL surface decorated with a set of steps with increasing thickness in the nanometer range1 (Silios, France). The AR-XPOL stage is made of a silicon support covered with a 106 nm silica layer (Vegatec, France). As explained in ref 1, the steps are made of additional silica layers, that is, simple changes in the layer thickness. They form a regular stair which was imaged between cross polarizers through a reflected microscope equipped with a standard xenon white-light source and recorded using a standard video camera. The intensity Iref in the image delivered by the camera has three color coordinates, Iref(x,y) ) [Rref(x,y),Gref(x,y),Bref(x,y)]. An image of a bare AR-X-POL surface is also recorded, and the corresponding intensity is Iback. Lightening heterogeneities over the microscope field are removed by dividing each component of Iref by the corresponding component in Iback normalized by its average value in the background image. We name IrefC the corrected image intensity. Using the same microscope setting, the same procedure, and the same background image, we obtain the corrected sample image intensity IsamC. The next step is to deduce local sample thickness from comparison of local sample image intensity IsamC with the finite set of step intensities IrefC[zi(x,y)], where “i” denotes the step index and i ) 0 corresponds to the bare surface. It supposes the existence of a mapping between intensity and thickness. Since the AR-X-POL surface corresponds to extinction of reflected light at average wavelength and average numerical aperture, the theoretical substrate intensity presents a minimum with the overlayer thickness, the position of which depends on wavelength. This is schematically illustrated in Figure (3) Blodgett, K. B. Phys. ReV. 1939, 55, 391-405.
10.1021/la0636098 CCC: $37.00 © 2007 American Chemical Society Published on Web 06/14/2007
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Figure 1. Intensity color components as a function of film thickness: (a) typical shape; (b) zoom of same in the selected thickness range and average slopes (straight lines); (c) color components weighted by their average slope and their combination (black line). This scheme illustrates how the useful gray scale is generated from the three color components.
1a. As a consequence, the relationship between thickness and intensity is not univocal at any wavelength. This difficulty can be overcome by defining a new gray-scale intensity as a linear combination of the three color components.4 However, depending on the microscope tuning, this combination would not be stable from one experiment to one another. This is why instead of a fixed linear relationship we define a nonlinear and self-adjusting combination of color components obeying the two requirements of a high sensitivity and of a monotonic variation with local sample thickness. It is set up as follows: IrefC gives the (averaged) values taken by the three color coordinates for the set of steps with thickness zi, from z1 to zn.From there, a fitting polynomial function PIrefC(z), with three color components, is determined, and its polynomial derivative (dPIrefC/dz)(z) is calculated. The thickness range is limited in such a way that a given color component has no more than one extremum. Although in general this limitation is severe, it is not important when dealing with molecular layers or ultrathin films. Then a polynomial scalar intensity PI(z) is defined as the following scalar combination:
PIrefC(z) )
∑
aXPXrefC(z)
X)R,G,B
where aX ) 〈dPXrefC/dz〉{zi}. In this expression, each color component in the image is simply weighted by its susceptibility to the film thickness averaged over the useful thickness range. Then it is checked that PIrefC(z) is a monotonic function over the thickness range covered by the steps. Then PIrefC(z) can be inverted to define a continuous function Fz(IrefC). On the other hand, the three color components of the sample image IsamC are used to generate the scalar gray-scale intensity
WIsamC[z(x,y)] )
∑
aXXsamC(x,y)
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with local sample thickness z. Figure 1b focuses on the thickness range associated to the n steps and displays the average slope aX associated with each component. Figure 1c shows the variation with z of the three weighted components aXPXrefC(z) and shows how they combine in PIrefC(z). To summarize, recording three images (sample under study, reference sample, bare substrate) with the same microscope is sufficient to obtain the sample film thickness distribution. As long as the microscope setting is not changed (lightening adjustment, aperture diaphragm, light intensity, sample orientation), new sample images can be recorded, allowing the study of thickness distribution dynamics. The possibilities offered by this technique will now be illustrated by three examples. The aim of the first example is to validate quantitative SEEC by comparison with AFM, with the help of a well-structured sample: a supported film of phospholipids which exhibits natural quantization of its thickness.5-7 Ultrathin films of commercial egg phosphatidylcholine (EPC, Aldrich) zwitterionic phospholipids were deposited on an AR-X-POL surface from chloroform solution dried under vacuum at room temperature for 2 h and then imaged though the microscope. As expected, the films presented locally the vicinal surface structure characteristic of lamellar systems, with each step corresponding to one additional bilayer. Using our optical method, the film thickness was measured and the height value of the first four steps from the solid was compared to that obtained with the AFM. On fresh samples, both instruments gave a value of 4.3 ( 0.2 nm, independent of layer rank. After some days in ambient atmosphere, this thickness slowly evolved over 5 nm due to sample hydration.8 Notice that the measurement was about 10 times faster with the optical method, and that the lateral size of a single image was not limited by the instrument as is the case with the AFM. Figure 2a presents a three-dimensional (3D) display of the stairlike reference sample. The thicknesses of the four steps shown in the image are respectively 2.0, 10.1, 18.3, and 22.3 nm, counted from the contrast silica layer. Figure 2b shows a 3D image of the phospholipids relief as obtained from the optical technique. Figure 2c shows the thickness profile obtained along a given line through the sample. After generating the gray-scale images, 3D display and topographic measurement were readily obtained with the help of a commercial 3D viewing and measuring software (Digital Surf, France). In a second example, the method was applied to probing the crystallization kinetics of a very thin polymer film. This realtime experiment would be difficult to perform with an AFM because the field of the instrument is too limited. Since most of the polymer materials crystallize, polymer crystallization is an important field of research driven by industrial needs. The basic question concerns the transition from the amorphous state of a random coil to the ordered crystalline state which consists of folded chains.9,10 Due to chain length and consequent limits in the motion and organization of individual polymer chains, only imperfect crystals may be formed, in particular for fast crystal growth. Thus, crystalline regions are typically embedded in an amorphous environment. A coexistence of flat domains, crystalline and amorphous, is obtained at a molecular scale. The field
X)R,G,B
from where we get the film thickness distribution z(x,y) ) Fz(WIsamC(x,y)). This procedure is schematically illustrated in Figure 1. Figure 1a shows a typical dependence of color-component intensities (4) Wyszecki, G.; Stiles, W.; Colour Science: Concepts and Methods, QuantitatiVe Data and Formulae; John Wiley & Sons: New York, 1982.
(5) Tokumasu, F.; Jin, A. J.; Feigenson, G. W.; Dvorak, J. A. Biophys. J. 2003, 84, 2609-2618. (6) Reviakine, I.; Brisson, A. Langmuir 2000, 16, 1806-1815. (7) Perino-Gallice, L.; Fragneto, G.; Mennicke, U.; Salditt, T.; Rieutord, F. Eur. Phys. J. 2002, 8, 275-282. (8) Seul, M.; Eisenberger, P. Phys. ReV. A 1989, 39, 4242-4253. (9) Sadler, D. M. New explanation for chain folding in polymers. Nature 1987, 326, 174-177. (10) Hoffman, J. D.; Miller, R. L. Polymer 1997, 38, 3151.
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Figure 3. Time sequence extracted from real-time observation of PEG thin film crystallization kinetics. The time is written on each image. It is counted in minutes from an arbitrary origin. Objective lens Leica HC PL Fluotar X50/0;80. The propagating crystal front passing the observed field results in a bright growing dendritic pattern surrounded by a dark depletion zone.
Figure 2. SEEC images of (a) the reference sample and (b) the supported phospholipid film. (c) Thickness profile along the dotted line through (b). Comparison with the reference sample allows one to measure and diplay the relief patterns which result from the quantization in bilayer units of the phospholipid film.
was marked in the 1970s by di Marzio,11,12 who introduced the basic model for semicrystalline polymers. In this picture, a given polymer chain contributes to both kinds of domains in a sequence of ordered partsswith several chain folds in a stemsand disordered partssrandom walks emerging from and connecting the flat stems. As a consequence, the domains are arranged in a lamellar structure with a crystal-amorphous interface normal to the stem plane. For sufficiently long polymers, the lamellar period P is smaller than the coil diameter of a molten chain, say from several nanometers to several tens of nanometers. When such a polymer crystallizes as a thin film with thickness less than or comparable to P on a solid substrate, the confined lamellae necessarily lie parallel to the surface of the solid. Flat crystalline and amorphous regions coexist side by side within the film; the interface between crystalline and amorphous regions is now located at the crystal edge. The crystalline regions are made of folded chains standing up and are fed by the still amorphous regions of the film.13 Following ref 13, polymer crystals cannot (11) di Marzio, E. A. J. Chem. Phys. 1971, 55, 4318. (12) Sanchez, I. C.; di Marzio, E. A. J. Chem. Phys. 1971, 55, 893-908. (13) Reiter, G.; Sommer, J.-U. Phys. ReV. Lett. 1998, 80, 3771-3774.
easily nucleate in ultrathin films but can nevertheless be observed there from crystal growth after nucleation in thicker parts of the film. The polymer used in this experiment is a common poly(ethylene oxide) (PEO) with molecular mass Mw ) 4000 g/mol, which crystallizes at room temperature. Here a polymer film with 6 nm thickness was spin-coated on an AR-X-POL surface and then immediately observed under the polarization microscope. The whole crystallization process lasted about 2 h. We took a series of pictures of a fixed region of the sample, which the crystallization front passed during this time. Following the procedure described above, color components were converted into thickness gray scale, which allowed us to render the relief evolution of the crystallization pattern and to study its dynamics quantitatively by measuring both the height of the flat-on crystal dendritic lamellae, the local height of the amorphous regions, and the local or averaged speed of the crystal front. Images extracted from that sequence are presented in Figure 3, showing how two-dimensional crystallization propagates through the 6 nm homogeneous film to form dendrite fingers having a 10 ()6 + 4) nm overall thickness and a typical width of 1 µm. In between these fingers, homogeneous amorphous regions are left with a thickness reduced by 0.2 nm with respect to that of molten metastable regions located far ahead of the crystal front. This depletion zone is clearly visible on images at intermediate times (t ) 41-67 min). From our experiment which was conducted at room temperature, the speed of the propagating crystal front envelope is found to be close to 1 µm/nm. The crystal height can also be used to determine the number of folds per chain in the crystal. Using published PEO characteristics,14 we found that it was only 2, indicating that the polymer chains are strongly extended.15 Our results are in good agreement with published data.16 However, in contrast to previous studies using AFM, we obtained our results in real time in a single experiment. The SEEC technique can be easily applied as well to a large variety of tasks involving soft surfaces including other phase transitions in molecular films, wetting and dewetting, or membrane swelling/ fluctuations. The third example illustrates the potential of the SEEC technique for quantitatively reading peptide-antibody biochips, (14) Kovacs, A. J.; Straupe, J. J. Cryst. Growth 1980, 48, 210-226. (15) Ungar, G.; Mandal, P. K.; Higgs, P. G.; de Siva, D. S. M.; Boda, E.; Chen, C. M. Phys. ReV. Lett. 2000, 85, 4397-4400. (16) Reiter, G.; Vidal, J. Eur. Phys. J. E 2003, 12, 497-505.
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Figure 4. SEEC images of a MYC peptide spot (left) and of a HA peptide spot (right) after incubation with HA antibody. Objective lens Leica HC PL Fluotar X20/0;50. The negative spot appears as a depression due to saturation of the outsides by BSA. The positive spot appears as an island due to the amount of captured antibody.
and more generally for probing binding interactions between biological species and receptors on surfaces. In this experiment, peptide sequences were fixed on contrast-enhancing supports and then incubated with positive and negative antibodies. Two different sequences of peptides derived from the influenza hemaglutinin (HA) sequence17 [H-S-G-(Y-P-Y-D-V-P-D-Y-A)G-(Y-P-Y-D-V-P-D-Y-A)-G-(Y-P-Y-D-V-P-D-Y-A)-S-NH2] and MYC sequence18 derived from human MYC protein [H-S-(EQ-K-L-I-S-E-E-D-L-N)-G-(E-Q-K-L-I-S-E-E-D-L-N)-A-(E-QK-L-I-S-E-E-D-L-N)-G-NH2] were spotted from 0.1 M, pH 5.5 sodium acetate buffer on amine-coated AR-X-POL surfaces. Then a standard procedure issued from ELISA tests was used. It involves two blocking agents, Tween 20 and BSA, dissolved in a pH 7.2, 1× PBS buffer (ELISA buffer). This procedure takes into account the following hierarchy of binding strength with the surface: peptide sequence-Tween 20-BSA. It basically consists in fixing peptide sequences within the spots, saturating the remaining of the spot area with Tween 20, and then saturating the remaining of the surface with BSA. As a result, spot areas were covered with peptide sequences and Tween 20, while the out-of-spot surface was covered with BSA. Since peptide and Tween 20 are much smaller molecules than BSA, a difference in height was observed between the spots and the rest of the surface. HA and MYC peptide samples were incubated with anti-HA antibodies at concentration CHA and then washed and dried and observed in air. On positive HA peptide samples, the spot altitude was increased with antibody capture, while it remained unchanged on negative MYC samples. This is illustrated in Figure 4. It shows the detailed structure of a negative spot (left) and of a positive spot (right), both obtained after incubation with HA antibody at concentration CHA ) 10 µg/mL. Prior to incubation, the negative spot was spotted with MYC peptide at molar fraction cp ) 100 µM and the positive spot was spotted with HA peptide at same molar fraction. The corresponding step heights with respect to BSA layer are respectively -300 and +1780 pm. Symmetric experiments with MYC incubation gave symmetric results. Figure 5 reports in the upper graph the variation of the spot altitude counted from the level of the BSA layer as a function of incubating anti-HA antibody concentration CHA between 0.5 and 10 µg/mL for different spotting peptide molar fractions ranging from 1 to 100 µM. A monotonic variation of the spot height with incubating concentration was found as expected. These variations are in perfect agreement with ELISA tests performed on the same systems (not reported here). Our method (17) Wilson, I. A.; Niman, H. L.; Houghten, R. A.; Cherenson, A. R.; Connolly, M. L.; Lerner, R. A. Cell 1984, 37, 767-778. (18) Evan, G. I.; Lewis, G. K.; Ramsay, G.; Bishop, J. M. Mol. Cell. Biol. 1985, 5, 3610-3616.
is therefore able to estimate amounts of captured materials. In addition, it proved to be highly reproducible since the same height was found within 50 pm on nine independent measurements (three independent experiments on three replicates) for each combination of peptide/antibody concentrations. We will now comment on the technique sensitivity. First, we focus on the lower peptide molar fraction, cp ) 1 µM, with the help of the lower graph in Figure 5, the vertical scale of which is reduced with respect to that of the upper graph. The 50 pm uncertainty of these data is imposed by sample roughness and reproducibility, with the reproducibility and sensitivity of the technique itself being better than this limit. For comparison, assuming a density of 1 g/cm3, a layer with average thickness 100 pm corresponds to a surface excess of 100 pg/mm2, or 100 RU (resonance units) in surface plasmon resonance (SPR) experiments.19 It is a reasonably good sensitivity level20 for this technique, which is the most widely used in label-free detection of binding interactions.21 However, one should keep in mind that, in contrast to SPR,22 the present technique offers the capacity of instantaneous imaging with full optical microscopy resolution. Second, we examine in Figure 6 the four situations indicated in Figure 5 by a black circle, with very close spot heights. The SEEC technique makes it possible to quantitatively estimate these heights, hence the amount of captured antibody, as well as to probe the detailed structure of each spot. The images in Figure 6 were obtained after incubating the sample with the samesthe lowestsantibody concentration CHA ) 0.5 µg/mL. From left to right and then from top to bottom, the four spots correspond respectively to spotting peptide molar fractions cp ) 1, 5, 10, and 100 µM. Thanks to lateral resolution, these images reveal localized spotting defects that were removed from the measured areas. The average level was thus found to be respectively -300, -40, +40, and +160 pm. Notice that the spots remain well visible although the range of the z-axis is smaller than the range of the x- and y-axes by a factor of 100 000. To summarize, SEEC is a quantitative imaging technique which combines high contrast enhancement by the use of nondepolarizing sample stages and self-building gray-scale intensity proportional to sample thickness. From the three examples given above, it appears as an interesting alternative or complement to AFM, imaging ellipsometry,23 and SPR for probing supported molecular and submolecular films in real time. In particular, the (19) Homola, J.; Yee, S. S.; Gauglitz, G. Sens. Actuators, B 1999, 54, 3-15. (20) Peter, J. C.; Briand, J. P.; Hoebeke, J. J. Immunol. Methods 2003, 74, 149-158. (21) Turner, A. P. Science 2000, 290, 1315-1317. (22) Su, Y. D., Chen, S. J.; Yeh, T. L. Opt. Lett. 2005, 30, 1488-1490. (23) Jin, G.; Tengvall, P.; Lundstro¨m, I.; Arwin, H. A. Anal. Biochem. 1995, 232, 69-72.
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Figure 5. Variation of the spot height (picometers) with respect to the BSA layer as a function of anti-HA incubating concentration CHA (mg/mL, log scale) for different HA spotting peptide molar fractions cp: upper curve, from top to bottom, cp ) 100, 10, 5, and 1 µM; lower curve, cp ) 1 µM with z-scale reduction. A monotonic variation of the spot height is observed as expected with both peptide and antibody concentrations.
Figure 6. Image of a spot obtained with spotting molar fraction cp ) 1 µM (upper left), 5 µM (upper right), 10 µM (lower left), and 100 µM (lower right) after incubation with HA antibody concentration CHA ) 0.5 µg/mL. Objective lens Leica HC PL Fluotar X20/0;50. The minute variations of the spot height with peptide molar fraction obtained with the lower incubating concentration are still visble on a subnanometer z-scale.
third example showed that the technique is appropriate for labelfree biochip reading and more generally for label-free detection of binding interactions. The lateral resolution of our technique is that of optical microscopy, that is, a fraction of a micron. This resolution is intermediate between that of AFM (subnanometer) and that of ellipsometry and SPR imaging (more than 10 µm). It also presents the advantages of simplicity, low cost, and high speed. Since it is a wide field microscopy technique, a major
difference from the above-mentioned other techniques is parallel acquisition of the sample image to be compared with point after point and line after line scanning. Like SPR and ellipsometry, SEEC is a refractive method. As it was used in the present work, it is mainly sensitive to the optical thickness of the supported film and only weakly sensitive to changes in the film refractive index at constant optical path (refractive index is assumed to be 1.47 in all the above experiments). Assuming a constant refractive
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index in the supported film is also a crude approximation, and it was shown by Proehl et al.24 that the effective refractive index can vary considerably in growing thin films. This very fundamental task is beyond our present scope, but we want to make very clear that the oversimplified assumption of a constant and homogeneous refractive index in the organic layer was implicitly made throughout our article. Like SPR, SEEC needs a special surface to work on. However, the flexibility to design this surface is very high,1 so the technique is not much limited in terms of surface materials and surface properties. The contrastenhancing surfaces used in this study are only approaching the index-matching condition given at the very beginning of this paper, and they can be largely improved. The entire technique is still in a first-stage level of development and has much room (24) Proehl, H.; Nitsche, R.; Dienes, T.; Leo, K.; Fritz, T. Phys. ReV. B 2005, 71, 165207.
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for progressing both in terms of performancesa subpicometer sensitivity is expected within 2 yearssand in terms of versatilitys a next step being to probe real-time kinetics under immersion. Hence combining quantitative imaging, high sensitivity, high resolution, real time, ease of use, low cost, and high evolution potential, the SEEC technique should rapidly enter the basic toolbox for biotechnology and soft surface science. Acknowledgment. We thank Gu¨nter Reiter for introducing us to polymer crystallization in thin films, for helpful discussions, and for a critical reading of our report on polymer experiments. We thank Oleg Melnyk for providing us with the biochip samples and for helpful discussions. We are indebted to Vianney Souplet and Yohann Lebel for their help with optical and AFM acquisitions. LA0636098
