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Influence of Surfactants on an Evaporating Drop: Fluorescence Images and Particle Deposition Patterns Van Nguyen Truskett† Department of Chemical and Biomolecular Engineering, Johns Hopkins University, Baltimore, Maryland 21218
Kathleen J. Stebe* Department of Chemical and Biomolecular Engineering, Department of Materials Science & Engineering, Department of Mechanical Engineering, and Department of Biomedical Engineering, Johns Hopkins University, Baltimore, Maryland 21218 Received February 6, 2003. In Final Form: June 30, 2003 The insoluble surfactant pentadecanoic acid is spread at the interface of aqueous droplets, which are then deposited on a well-defined substrate and allowed to evaporate. The surface state of the surfactant is imaged as the drop evaporates using fluorescence microscopy, and the mean rate of evaporation is calculated from successive digitized silhouettes of the sessile drops. The drops contain suspended microspheres that act as tracers in the flow within the drop and deposit on the substrate in patterns that vary with the surfactant surface state. Patterns observed include circular mounds of particles at the original drop periphery termed “coffee rings”, created by an outward flow to the three-phase contact line; connected polygons formed by Marangoni-Be´nard flow; and uniform monolayers of microspheres deposited on the substrate. These results establish that surfactants can be used to alter flow fields and deposition patterns from evaporating aqueous drops. This is potentially useful in the organization of nanoparticles and in the deposition of materials functionalized with specific binding sites whose folded structures might be destroyed by other solvents. The deposition of biologically functionalized materials is demonstrated by depositing streptavidin-labeled microspheres, which retain their ability to bind biotin after deposition.
Introduction Evaporating drops have been used as vehicles for organizing small particles (microspheres to nanospheres) suspended within them.1 The particles act as tracers in the flow fields created within the drop during evaporation, which determine the distribution of the particles once the drops dry. One pattern that forms is termed a “coffeering” pattern, consisting of a mound of particles near the three-phase contact line at the drop periphery.1-3 When a partially wetting drop is placed on a solid substrate, it forms a spherical cap that intersects the substrate at its advancing contact angle. The three-phase contact line remains fixed as the drop evaporates until the contact angle reduces to its receding value. The evaporation rate is fastest in the vicinity of the three-phase contact line. This large local flux creates an outward flow that convects suspended particles toward the contact line. The particles accumulate on the substrate in the three-phase contact region, creating a locally rough surface with a very low receding contact angle. Thus, particle accumulation pins the three-phase contact line during much of the drop evaporation. The outward flow toward the contact line persists, causing particles to accumulate in a circular ring. This ring is left as residue after the drop has completely evaporated.1 The contraction of evaporating capillary bridges between the particles forces them into an ordered arrangement.3 This mechanism has also been used to * Author to whom correspondence should be addressed. † Current address: Molecular Imprints, Inc., Austin, TX 78758. (1) Deegan, R. D.; Bakajin, O.; Dupont, T. F.; Huber, G.; Nagel, S. R.; Witten, T. A. Nature 1997, 389, 827. (2) Deegan, R. D. Phys. Rev. E 2000, 61, 475. (3) Maenosono, S.; Dushkin, C. D.; Saita, S.; Yamaguchi, Y. Langmuir 1999, 15, 957.
create colloid crystals from evaporating fluid layers.4-7 A similar mechanism is responsible for the starburst pattern of DNA that is created when DNA is deposited from evaporating drops on microarrays.8 In this case, because DNA does not pin the three-phase contact line, the outward flow brings DNA to the original periphery of the drop only during the time that the contact angle is reducing from its advancing to its receding value. Once deposited, if interactions with the substrate are not too pronounced, the outward extensional flow can stretch the DNA to form a “starburst” pattern. If the distribution of cDNA (or other material to be deposited) is nonuniform within the drop, nonaxisymmetric rings can be created.9 Another pattern that has been reported from evaporating drops is created by Marangoni-Be´nard convection, a spatially periodic flow created by a thermocapillary-driven instability at the free surface.10 This flow field has been used to deposit zeolite nanocrystals on silicon wafers to form nanoporous films with characteristic feature sizes of 5-10 µm;11 to deposit a variety of nanoparticles (e.g., cadmium sulfide, silver, cobalt, and ferrite nanocrystals) from organic solvents, (4) Adachi, E.; Dimitrov, A. S.; Nagayama, K. Langmuir 1995, 11, 1057. (5) Denkov, N. D.; Velev, O. D.; Kralchevsky, P. A.; Ivanov, I. B.; Yoshimura, H. Nature 1993, 361, 26. (6) Denkov, N. D.; Velev, O. D.; Kralchevsky, P. A.; Ivanov, I. B.; Yoshimura, H. Langmuir 1992, 8, 3183. (7) Dushkin, C. D.; Yoshimura, H.; Nagayama, K. Chem. Phys. Lett. 1993, 204, 455. (8) Jing, J.; Reed, J.; Huang, J.; Hu, X.; Clarke, V.; Edington, J.; Housman, D.; Anantharaman, T. S.; Huff, E. J.; Mishra, B.; Porter, B.; Shenker, A.; Wolfson, E.; Hiort, C.; Kantor, R.; Aston, C. Proc. Natl Acad. Sci. U.S.A. 1998, 95, 8046. (9) Blossey R.; Blosio, A. Langmuir 2002, 18, 2952. (10) Pearson, J. R. A. J. Fluid Mech. 1958, 4, 489. (11) Wang, H. T.; Wang, Z. B.; Huang, L. M.; Mitra, A.; Yan, Y. S. Langmuir 2001, 17, 2572.
10.1021/la030049t CCC: $25.00 © 2003 American Chemical Society Published on Web 09/03/2003
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Figure 1. Schematic of the experimental apparatus.
with Be´nard cells as small as 0.5 µm;12 and to deposit alkanethiol-functionalized gold nanoparticles (4-10-nm diameter) to form 3-µm Be´nard cells.13 Normally, this flow is suppressed by surfactants in aqueous systems.14 However, under certain circumstances, surfactants promote this instability.15 When surfactants are spread at the interface to form coexisting liquid expanded-liquid condensed surface phases, the order-disorder transition causes the surface tension to be highly dependent on the temperature. This enhanced thermocapillary coupling and the absence of surface-tension gradients in response to fluctuations in surface concentration for a first-order phase transition drive the system into Marangoni-Be´nard flow. In this work, the effect of surfactants in changing the deposition patterns of small particles from evaporating drops is investigated. The surfactant, pentadecanoic acid (PDA), is spread on the surface of a pendant drop, whose surface tension is measured prior to depositing the drop on a solid substrate to form a sessile drop. The surface state of the surfactant at the liquid-gas interface is imaged as the sessile drop evaporates using fluorescence microscopy. Images of the silhouette of the drop are also recorded periodically, allowing the average rate of evaporation from the drops to be calculated. The mean evaporation rate of the drops is not altered by the presence of surfactants. However, the microspheres deposit in patterns that differ strongly with the surface state. Because the particles act as tracers in the flow within the drop, this suggests that strong changes in the flow field within the drop are created by the surfactants. These changes are created either by (12) Maillard, M.; Motte, L.; Ngo, A. T.; Pileni, M. P. J. Phys. Chem. B 2000, 104, 11871. Maillard, M.; Motte, L.; Pileni, M. P. Adv. Mater. 2001, 13, 200. (13) Stowell, C.; Korgel, B. A. Nano Lett. 2001, 1, 595. (14) Berg, J. C.; Acrivos, A. Chem. Eng. Sci. 1965, 20, 737. (15) Nguyen, V. X.; Stebe, K. J. Phys. Rev. Lett. 2002, 88, 164501.
Figure 2. Compression isotherm for PDA, with corresponding fluorescence images for G-LE coexistence, for the LE state, for LE-LC coexistence, and for the LC state. The bright speckles appear on the surface in the LC image because the monolayer has been compressed above the equilibrium spreading pressure of the dye, which is roughly 12 mN/m.
changing the stress conditions at the liquid-gas interface or by changing the local evaporation rate in the vicinity of the three-phase contact line. These results may be of practical as well as fundamental interest. Evaporative deposition techniques have potential in emerging technologies, including in the organization of nanoparticles for exploitation as sensors and photonic devices,16,17 and in the deposition of solutes on microarrays18 used in combinatorial chemistry studies in genom(16) Murray, C. B.; Kagan, C. R.; Bawendi, M. G. Annu. Rev. Mater. Sci. 2000, 30, 545. (17) Alivisatos, P. Pure Appl. Chem. 2000, 72, 3.
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Figure 3. Images of a surfactant-free evaporating drop containing amidine-functionalized microspheres obtained using reflected light microscopy. (a) t ) 0. (b) t ) 45 min (moments before depinning and hopping to a new location occurs). (c) t ) 55 min. Sedimented particles forming a “coffee ring” are apparent at the drop’s initial location. (d) Final pattern deposited by (dried) particles.
ics8,19 and proteomics.20 Furthermore, the ability to create these patterns of particles from aqueous media may be useful for systems containing proteins or other biomolecules whose folded structures would be disrupted in nonaqueous media. The deposition of protein-labeled particles is demonstrated using streptavidin-labeled microspheres that are deposited in a periodic array created by Be´nard convection. The streptavidin on the microspheres retains its ability to bind biotin after the patterns are deposited. Experimental Section Materials. All glassware was soaked in a H2SO4-Nochromix solution (technical grade, Fisher Scientific) overnight and rinsed with filtered and deionized water with a resistivity of 18.0 MΩ cm (Millipore Milli-Q 50). This water was used to clean the substrates and to prepare all the aqueous solutions. All chemicals were used as received without further purification. Monolayers of PDA (Fluka, g99.5%; GC grade) were spread from either chloroform or ethanol (Sigma Chemical Co., HPLC grade) doped with e1 mol % 4-hexadecylamino-7-nitrobenz-2-oxa-1,3-diazole (NBD-HDA; Molecular Probes, Inc.). The pHs of the aqueous suspensions of microspheres and of the aqueous subphase for the insoluble monolayer studies were adjusted to pH 2 by the addition of HCl (J. T. Baker, ACS grade) to prevent the dissociation of the PDA headgroup. The red-dyed surfactantfree sulfate- and amidine-functionalized polystyrene microspheres of 0.8-µm diameter (Interfacial Dynamics, Corp.) were washed with and resuspended to 0.01% solid w/v in a pH 2 aqueous solution. All colloidal suspensions were stored in acidwashed polypropylene bottles. Streptavidin-functionalized polystyrene microspheres of 0.30-µm diameter (Bangs Laboratories, Inc.) were received as a 1% solid w/v suspension in 100 mM borate, pH 8.5, + 0.1% bovine serum albumin + 0.05% Tween + 10 mM ethylenediaminetetraacetic acid buffer with 0.1% NaN3 antimicrobial agent and are red in color with a binding capacity of 3.3-µg biotin-fluorescein/mg microspheres (as was reported by the manufacturer). The streptavidin-functionalized microspheres were resuspended in a 0.015% solid w/v microspheres, pH 2.0, aqueous HCl solution. The fluorescein-biotin solution was prepared with a phosphate buffer solution (PBS). The activity of streptavidin was confirmed by exposing the particles for 2 h to a 300 nM fluorescein-biotin solution (Molecular Probes) in (18) MacBeath, G.; Schreiber, S. L. Science 2000, 289, 1760. (19) Jain, K. K. Science 2001, 294, 621. (20) Zhu, H.; Synder, M. Curr. Opin. Chem. Biol. 2001, 5, 40.
PBS and followed by a wash with PBS. The fluorescein-labeled biotin excites at a wavelength of 494 nm and fluoresces at 518 nm. Octadecyltrichlorosilane (OTS)-Modified Substrates. Silicon wafers with a native oxide layer (P type 〈111〉, prime grade; Montco Silicon Technologies, Inc.) were modified with a self-assembled monolayer (SAM) of OTS (Sigma Chemical Co., HPLC grade). OTS solutions were made with pentane (Sigma Chemical Co., HPLC grade). All glassware was silanized prior to being used by exposure to a 2 M OTS solution for 24 h. The silicon substrates were cut into 3 × 3 cm pieces, placed in a glass slide holder, and sonicated for 30 min in a fresh H2SO4Nochromix solution. Thereafter, the substrates were rinsed and sonicated for 30 min in water. These substrates were stored in water to prevent the adsorption of airborne contaminants. Prior to immersion in the OTS solution, the samples were dried in a stream of filtered air or N2. The SAM was formed by immersing the substrate in a 1.0 mM OTS solution. Solutions were used within 2 h after being prepared. For fresh reagents, an immersion time of 15 s yields a SAM with a contact angle of 50 ( 3°. Upon removal from the solution, the samples were promptly rinsed with chloroform for 2 min and dried in a stream of filtered air or N2. PDA Isotherm and Fluorescence Microscopy. The Teflon Langmuir trough (KSV Instruments) was cleaned repeatedly by wiping sequentially with ethanol, water, methanol, and water. A platinum Wilhelmy plate (KSV Instruments) was cleaned by sonication and dried in an open flame. Surface pressure data were obtained using a force transducer (KSV Instruments) with a sensitivity better than 0.5 mN/m. The PDA monolayer was spread from a 1.0 mg/mL chloroform solution onto the aqueous subphase with a 10-µL Hamilton syringe. The syringe itself was rinsed with chloroform, disassembled, and sonicated repeatedly in water. The PDA monolayer was deposited at an initial area per molecule of ∼150 Å2/molecule and compressed at a rate no faster than 0.832 Å2/(molecule min). Fluorescence images were obtained with an Axiotech Vario 100HD Carl Zeiss epi-fluorescence microscope equipped with 2.5×, 10×, and 20× objectives, a C2400-97 Hammamatsu Intensified charge coupled device (CCD) camera, EPIX PIXCI SV4 imaging board, and XCAP image software. The fluorescent probe, NBD-HDA, was excited with a Spectra Physics Stabilite 2017-05S argon ion laser tuned to a wavelength of 467 nm at a power output of less than 10 mW. Light at the emission wavelength of 538 nm was filtered through a long-pass Chroma Technology basic blue filter. The images of the surface phases formed by PDA were obtained using two Teflon 10-mL beakers containing a pH 2 aqueous subphase. One beaker was configured beneath the microscope
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Truskett and Stebe allowing the apparent contact angle and drop shape to be recorded. The substrate was located beneath a fluorescence video microscope. A laser beam impinging on the surface of the evaporating sessile drop was used to excite the fluorescence probe in the surfactant monolayer. This apparatus allows the measurement of the surface tension prior to drop deposition, the acquisition of sessile drop images, and the acquisition of fluorescence images of the surfactant surface state on the same drop. In a typical experiment, a monolayer of PDA doped with the fluorescent probe NBD-HDA was spread from ethanol on the surface of a pendant drop. The drop was expanded by injection of liquid slowly over a period of 20 min, allowing the ethanol to evaporate while reaching the experimental drop volume of 10.5 µL. The surface tension γpd 0 was determined from the pendant drop profile. The drop was deposited on the substrate by lowering the drop to meet the substrate. Two images of the sessile drop (one digitized silhouette and one fluorescence image) were acquired immediately after drop deposition (which is defined as t ) 0 in our experiments) and at 5-min intervals as the drop evaporated. The deposited drop volumes ranged from 6.5 to 8.4 µL, averaging 7.4 ( 0.5 µL. All experiments were performed at 50% relative humidity and at 23 ( 0.5 °C unless otherwise noted.
Results and Discussion
Figure 4. Data obtained from the silhouette of a surfactantfree drop containing amidine-functionalized microspheres. (a) Drop profiles shown are recorded at 5-min intervals. (b) The contact angle before and after the drop depins. (c) Radius of curvature at the apex before and after the drop depins. to image the fluorescence domains; the other beaker was used to measure the surface tension by a Wilhelmy plate technique. After aspirating the air-liquid interfaces in both beakers, PDA doped with NBD-HDA was spread from chloroform in a single shot on each interface. A cover slip was placed on top of the fluorescence beaker to quiet surface convection before imaging the domains. Drop Evaporation Imaging Apparatus and Protocol. The experimental apparatus for drop formation, deposition, and imaging is shown in Figure 1. In each experiment, the silhouette of a pendant drop formed in the path of a collimated light beam was projected onto a CCD camera configured to a frame grabber/ digitization board in a PC (shown from right to left in the figure). From the digitized drop shape, the surface tension was determined by comparing the drop profile to a numerical solution of the Young-Laplace equation (with a precision of (0.1 mN/m).21,22 The pendant drop was then placed onto a substrate to form a sessile drop. The silhouette of the sessile drop was also digitized, (21) Lin, S. Y.; McKeigue, K.; Maldarelli, C. Langmuir 1991, 7, 1055. (22) Pan, R. N.; Green, J.; Maldarelli, C. J. Colloid Interface Sci. 1998, 205, 213.
The compression isotherm for PDA is shown in Figure 2 with corresponding fluorescence images for surface gaseous-liquid expanded coexistence (denoted G-LE), in which the gaseous phase appears as dark regions in a bright liquid expanded phase; for the liquid expanded state (denoted LE), which is uniformly bright; for liquid expanded-liquid condensed coexistence (denoted LELC), in which the more ordered state excludes the fluorescent probe and appears dark in a bright LE phase; and for the liquid condensed state (denoted LC), which is dark with bright specks created by crystals of the probe. These crystals form in monolayers compressed to surface pressures above the equilibrium spreading pressure of the probe, ∼12 mN/m. These images correspond well to those obtained by prior researchers using this fluorescent probe23 and by Brewster angle microscopy.24 The G-LE and LE-LC phase transitions are first-order transitions25,26 and are strictly planar on a compression isotherm for highly purified PDA. (The slight upward slope apparent in the LE-LC coexistence region of these data is caused by residual impurities in the PDA.) The results described below were obtained using either amidine-functionalized microspheres (which are positively charged under the solution conditions) or sulfate-functionalized microspheres (which are negatively charged). Similar results were obtained for both types of microspheres. The patterns observed do not depend on electrostatic interactions between the functionalized groups and either the liquid-gas or the liquid-solid interface. The primary role of the charged groups is to create repulsive interactions among the microspheres that prevent flocculation. The microsphere system used in a particular experiment is reported in the figure captions for completeness. Illumination by light sources also do not alter deposition patterns, which remain qualitatively the same whether performed under the halogen and laser light sources that allow details of the drop to be recorded or in their absence. To confirm that the microspheres were free of surface-active impurities, the surface tension of a pendant drop containing microspheres was recorded. The (23) Moore, B. M.; Knobler, C. M.; Akamatsu, S.; Rondelez, F. J. Phys. Chem. 1990, 94, 4588-4595. (24) Honig, D.; Overbeck, G. A.; Mobius, D. Adv. Mater. 1992, 4, 419. (25) Pallas, N. R.; Pethica, B. A. Langmuir 1985, 1, 509. (26) Pallas, N. R.; Pethica, B. A. J. Chem. Soc., Faraday Trans. 1987, 1, 585.
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Figure 5. (a) Experimental flux JM as a function of the PDA surface state. Symbols: surfactant-free drops (]), G-LE (0), LE (4), LE-LC (×), or LC (O) surface states. (b) Comparison of JM (b) to JD (dashed line).
surface tension remained fixed at the value of a clean air-water interface for at least 1 h for both microsphere systems. Drops without a Surfactant Monolayer. In the absence of surfactants, it is well established that drops containing microspheres form coffee-ring patterns. Four images of the evolution of a surfactant-free evaporating drop are shown in Figure 3a-d. A topview of the drop taken immediately after deposition is shown in Figure 3a. The contact line remained fixed for 45 min (Figure 3b), after which it violently depinned and hopped to a new location, leaving a coffee-ring pattern of particles at the drop’s initial location (Figure 3c). After this first hopping event, the contact line depinned several more times, creating a series of rings and, finally, a dense patch of
particles formed by the microspheres that were suspended in the drop when it depinned for the last time (Figure 3d). These images were obtained using reflected light microscopy, where light from a halogen light source is glanced off the top of the drop or substrate. Figure 4 contains data obtained from the silhouette of the drop shown in Figure 3. The drop profiles, recorded at 5-min intervals, are shown in Figure 4a. The weak outward flow from the body of the drop to the contact line carried suspended particles with it. The periphery of the sessile drop occupied a nearly fixed location on the substrate as it flattened during the first 45 min of the evaporation. While the sessile drop profiles appear as if the contact line had receded, there was a thin film extending from the drop through a wet, sparse layer of
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Figure 6. (a) Fluorescence image of a drop at t ) 0. The interface is in G-LE coexistence (γpd 0 ) 72.4 mN/m, 2.5×). (b) A close-up view (10×). (c) Coffee ring formed by microspheres outlining the perimeter of the drop after complete evaporation.
Figure 7. (a) Fluorescence image at t ) 0. The drop interface is in the LE surface state (γpd 0 ) 72.2 mN/m). (b) A coffee ring of dried particles deposited from this drop.
particles next to the mound of particles in the “coffee ring”. The contact angle joining the drop to the thin film was far less than the apparent contact angle extracted from the drop profile. Eventually, the drop depinned. (This event has been ascribed to evaporation causing the water film that wets the deposited particles to become discontinuous.)27 The drop profile after the depinning event is indicated by the solid circles. The occurrence of these events is also evident as jumps in the data for the apparent contact angle, reported in Figure 4b, and the radius of curvature at the apex of the drop, reported in Figure 4c. Evaporative Mass Flux. For drops that remain axisymmetric (as can be ascertained from the images of the drop acquired from above), the evaporative flux from the drop JM can be calculated, using the area A and volume V extracted from successive drop profile images, according to the expression
JM )
-F dV A dt
where F denotes the density of water. The fluxes are reported for drops prior to depinning in Figure 5a. The flux JM remains fixed at roughly 2 × 10-5 g/(cm2 sec) for the first 35 min of the evaporation for all cases [except for the slightly higher value obtained for the drops covered with surfactant in the LC state, driven by the lower relative humidity in the laboratory (∼40%) during those experiments]. These results indicate that the interface does not exert a controlling role over the mean evaporation rate for drops of this curvature. The measured mass flux JM can be compared to a prediction assuming that the rate of evaporation is controlled by the rate that water vapor can diffuse from the drop across an unbounded vapor space, JD, using results from the numerical analysis of Hu and Larson.28 Their results can be expressed as a correlation in terms of the contact line radius Rc, the water vapor diffusivity D, the relative humidity H, the concentration of saturated water vapor C, and the contact angle θ: (27) Shmuylovich, L.; Shen, A. Q.; Stone, H. A. Langmuir 2000, 18, 3441. (28) Hu, H.; Larson, R. G. J. Phys. Chem. B 2002, 106, 1334.
JD )
DC (1 - H)(0.27θ2 + 1.30)(1 + cos θ) 2Rc
The parameters used to calculate JD were obtained from the sessile drop images and known laboratory conditions. As shown in Figure 5b, JM and JD (dashed line) agree to within an averaged deviation of 11% for the surfactantfree drops, which is within an averaged deviation of 30% for the drops with the LC monolayer. These deviations between JM and JD bound all of the other surfactant-laden drop cases, with no systematic variation with the surfactant concentration. For all drops studied, JM is always slightly in excess of JD, possibly because of weak convection in the laboratory. Drops with G-LE and LE Monolayers. Results for a drop coated with a monolayer in G-LE coexistence are presented in Figure 6. Fluorescence images of the surface state focused at the drop apex in Figure 6a,b were captured immediately after the drop was deposited. For this drop, the contact line remained pinned at the periphery for 55 min. Thereafter, the drop violently depinned, dragging the remaining microspheres within the drop to one side and leaving a coffee ring at the original drop perimeter (Figure 6c). Similar results were obtained for drops with an LE monolayer, as reported in Figure 7; in Figure 7a, the fluorescence image confirms that the drop interface is covered in the LE state. In Figure 7b, the residue of the particles left after the drop had dried is shown. Drops with LE-LC Monolayers. Striking changes occur in the deposited patterns when the interface is spread with a PDA monolayer in LE-LC coexistence. In Figure 8a, taken immediately after the drop is deposited, the fluorescence images confirm that the monolayer at the drop interface was in LE-LC coexistence. After 10 min, the surface area of the drop had decreased, compressing the monolayer to form more of the LC phase at the interface (not shown). During this time, a weak coffee ring was established at the drop periphery by the outward flow field. After 35 min (Figure 8b), roughly periodic hexagonal cells appeared in the drop as the microspheres, which were circulating in Be´nard convection (with particles moving upward in the center of each polygon and outward to the edge of each of the polygonal shapes),
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Figure 9. (a) Polygonal network displaying Be´nard cells with a variety of length scales and shapes, including squares, pentagons, and hexagons. (b) Irregular shapes observed with SEM for a network of Be´nard cells.
Figure 8. (a) Fluorescence images of a drop containing sulfatefunctionalized microspheres with a monolayer at its interface in LE-LC coexistence (γpd 0 ) 69.1 mN/m) at t ) 0. (b) Image of the particle distribution in an evaporating drop taken by reflected light microscopy at t ) 35 min. Periodic hexagonal cells are apparent in the drop. (c) Image of the residue of dried particles forming a connected network of polygons with a coffee ring encircling the perimeter taken by reflected light microscopy.
deposited on the substrate. These images were captured using reflected light microscopy, so only the particles were visible, not the fluorescent monolayer. The final pattern created by the particles is a connected network of polygons with a coffee ring encircling the perimeter (Figure 8c). In this example, the drop did not depin and the pattern was created everywhere under the original drop footprint. These connected polygonal networks are created by Marangoni-Be´nard flow, a flow created by a surface tension-driven instability. (These results are discussed in greater detail in ref 15). As the solvent evaporates, the drop cools at the interface to establish a temperature gradient as a function of the depth. Small perturbations in the surface temperature create gradients in the surface tension, or thermal Marangoni stresses. The interface contracts toward the high-tension regions, pulling hot fluid to the interface from below the warm regions, driving the system into a spatially periodic flow. The characteristic wavelength dictates the dimensions of the periodic flow patterns (known as Be´nard cells). It is well-established that the size of the Be´nard cells depends on the thickness of the film or drop height and on the magnitude of the Marangoni number for Be´nard flow B:
B)-
∂γ β d2 ∂T η κ
where κ is the thermal diffusivity and d is the depth of the fluid, ∂γ/∂T is the derivative of the surface tension γ with respect to temperature T, β is the base state temperature gradient established by the evaporative flux, and η is the viscosity. The critical Marangoni number, Bc ) 80, and the characteristic wavelength λ, giving the length scale of the Be´nard cells for linearly driven systems, are well-
established for liquid films in a planar geometry.10,29,30 Surfactants are usually thought to quell this instability by creating compositional Marangoni stresses that resist the surface flow.14 However, for surfactant monolayers undergoing first-order phase transitions, the surface tension is decoupled from the surface composition, so compositional Marangoni stresses are absent. Furthermore, for surfactant-laden interfaces in LE-LC coexistence, ∂γ/∂T ) -1.3 mN/(m K) (an order of magnitude greater than the derivative of the surface tension with temperature for surfactant-free aqueous drops).31 Thus, the mechanism that ordinarily resists this flow is absent, and the mechanism that drives this flow is enhanced. The corresponding value for B ) 650 for drops evaporating with interfaces in LE-LC coexistence is well in excess of the critical value for the onset of flow. The connected polygonal networks, mostly hexagonal in shape, were observed in several of our experiments, although other shapes can be identified, including squares and pentagons. In Figure 9, Be´nard cells with a variety of length scales and shapes are shown, with squares, pentagons, hexagons, and some irregular shapes. Mixtures of cell shapes and length scales have been reported in the literature for Marangoni-Be´nard flow in other solvents. The mixed polygonal state has been observed as an intermediate state for a nonlinearly driven instability undergoing a transition from a steady formation of hexagons to squares.32 Coarsening of the polygonal shapes has been reported for highly nonlinearly driven systems.33 For our system, B is far in excess of the critical value predicted by the linear analyses, and it is not unreasonable to expect nonlinear effects to manifest in these experiments. Drops with LC Monolayers. Images of a drop coated with a monolayer in the LC state are shown in Figure 10a. The interface is dark, with the exception of the crystals of the fluorescent probe. This drop remained in the LC state with a pinned contact line throughout its evaporation. The residue left by the particles is shown in Figure 10b. A dense layer of particles was deposited beneath the drop. The small bright regions in the figure were created by small local accumulations of particles stacked in multilayers. Scanning electron microscopy (SEM) images (not shown) reveal unevenly distributed, disordered microspheres everywhere under the drop, with no coffee ring at the periphery. Contact angle data are presented in Figure 10c. The initial contact angle for the LC case for (29) Scriven, L. E.; Sternling, C. V. J. Fluid Mech. 1964, 19, 321. (30) Nield, D. A. J. Fluid Mech. 1964, 19, 341. (31) Akamatsu, S.; Rondelez, F. J. Phys. II (France) 1991, 1, 1309. (32) Schatz, M. F.; VanHook, S. J.; McCormick, D. W.; Swift, J. B.; Swinney, H. L. Phys. Fluids 1999, 11, 2577. (33) Colinet, P.; Legros, J. C.; Velarde, M. G. Nonlinear Dynamics of Surface Tension Driven Instabilities; Wiley-VCH Verlag: Berlin, 2001; pp 6-9.
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Figure 10. (a) Fluorescence image at t ) 0 of a drop containing amidine-functionalized microspheres with a LC monolayer at its interface (γpd 0 ) 54.4 mN/m). (b) Reflected light image of the residue of dried particles left beneath the origial drop location. (c) Apparent contact angle.
all of the drops studied was slightly higher than the other surface states studied. This can be attributed to surfactant being trapped beneath the droplet when it is placed on the substrate. Experiments were performed to confirm that these results cannot be attributed to the high contact angle that prevailed during the LC experiments. Highcontact-angle OTS SAMs (105° with clean water) were used as substrates. The clean water evaporation experiments yielded coffee-ring patterns. The deposition patterns created beneath a LC-covered drop were similar to those in Figure 10a. The pattern formed beneath the LC-covered drops indicates that the outward flow toward the threephase contact line was absent or at least very weak. This can be ascribed to surfactant accumulation near the contact line, which was probably sufficiently pronounced to retard evaporation near the contact line relative to the rest of the drop surface. This would eliminate the outward flow that forms the coffee-ring pattern. (Compositional Marangoni stresses could alter the flow field, but because these stresses have a similar magnitude in the LE and in the LC states, they cannot explain the differences in the deposition patterns.)34 Patterning with Streptavidin-Functionalized Microspheres. The results described previously indicate that evaporating drops with surfactant-laden interfaces can be used to direct particle deposition from aqueous solutions. These results are of particular interest when proteins or other biologically based materials are to be assembled, for which an aqueous environment is favored over organic solvents to prevent conformational changes of the molecules. To demonstrate this concept, streptavidin-functionalized microspheres are deposited using the Marangoni-Be´nard instability. Prior to performing the deposition experiments, the surface tension of a pendant drop containing the streptavidin-functionalized microspheres was measured over a 2-h time span. For a sufficiently dilute suspension (0.015% solid w/v), the dynamic surface tension remained fixed at the value for that of a clean air-water interface up to 2 h, suggesting that streptavidin adsorption is sufficiently weak that a (34) If evaporation created a flow that collected surfactants locally at the three-phase contact line, the surface tension near the drop periphery would be reduced relative to elsewhere on the surface. This would create a stress that pulls from the contact line to the drop apex, resisting the outward flow. The characteristic magnitude of the Marangoni stresses is given by the Marangoni number Ma: Ma ) -[(∂γ/ ∂Γ)Γ]/µU. In this expression, µ ) 1 cp is the viscosity of the aqueous solution, U is the characteristic velocity (which is set by the evaporation rate so that U ) JM/F ) 10-7 m/s), Γ is the characteristic surface concentration, and ∂γ/∂Γ is the slope of the surface tension as a function of the surface concentration. The slope ∂γ/∂Γ is estimated by recasting the compression isotherm in Figure 2 in terms of the surface tension as a function of the inverse area/molecule, or surface concentration. Estimates obtained for the Marangoni number for the LE state (Ma ) 8.0 × 107) and for the LC state (Ma ) 2.0 × 108) are comparable.
Figure 11. (a) Fluorescence image of the LE-LC monolayer at the apex of a drop containing streptavidin-functionalized microspheres (γpd 0 ) 56.4 mN/m) at t ) 0. (b) Images of dried particles deposited on the surface after the drop had completely evaporated.
PDA monolayer can be used to control the behavior at the liquid-gas interface during the first hour the drop is formed. However, even weak adsorption of streptavidin into the PDA monolayer alters the surface pressures at which the phase transitions occur. We observed LE-LC coexistence using fluorescence images on streptavidin containing drops with γpd 0 values ranging from 56.4 to 68.7 mN/m. In the absence of a spread monolayer, the usual pattern of coffee rings and dense particle patches formed by depinning is observed (not shown). Results for drops covered in LE-LC coexistence are shown in Figure 11. In Figure 11, a uniformly distributed polygonal pattern is deposited with the length scale of the Be´nard cells being approximately 20-90 µm. The activity of the streptavidin bound to the deposited microspheres was determined by exposing the substrate to a solution of fluorescein-biotin. Prior to immersing the patterned substrate into the fluorescein-biotin solution, the sample was exposed to the excitation wavelength of the probe. No fluorescence was observed. The microspheres themselves are red in color (Figure 12a). After immersion in the fluoresceinbiotin solution (and subsequent rinsing), the polygonal network fluoresces green when excited at the appropriate wavelength (494 nm), confirming that streptavidin maintained its active binding sites (Figure 12b).
Surfactants, Drop Evaporation, Pattern Deposition
Langmuir, Vol. 19, No. 20, 2003 8279
Figure 12. (a) Particle distribution created by the evaporation of a drop in LE-LC coexistence containing streptavidin-labeled microspheres imaged using reflected light microscopy. (In a true-color image, the red color of the microspheres is apparent.) (b) A fluorescence image obtained by exciting fluorescein-labeled biotin bound to the streptavidin-labeled microspheres in part a. (In a true-color image, the green color of the excited fluorescein is apparent.)
Conclusions Evaporating aqueous drops containing suspended particles were studied as a function of the state of a surfactant monolayer at the aqueous-gas interface. The mean evaporation rate of the droplets was not influenced by the surfactant. However, the patterns formed by microspheres deposited from the evaporating drop changed strongly. The coffee-ring pattern formed from drops with monolayers in either the surface gaseous or in the liquid expanded states. A periodic array of microspheres arranged in polygonal cells were deposited by Marangoni-Be´nard flow created by the enhanced thermocapillary coupling for surfactants in liquid expanded-liquid condensed coexist-
ence. A layer of disordered microspheres settled everywhere beneath the original location of droplets covered with monolayers in the liquid condensed state. These results confirm that surfactants can provide a means of tailoring the fluid interface response in evaporating aqueous layers and can be used to direct the deposition of particles from evaporating aqueous systems. Acknowledgment. We thank NASA’s Office of Biological and Physical Research (Grant NAG3-1923 from NASA Glenn Research Center) and NASA’s Graduate Student Researchers Program (Grant GNT5-50268). LA030049T
