A Catch-Bond Based Nanoadhesive Sensitive to Shear Stress

ABSTRACT. The bacterial adhesive protein FimH forms bonds so-called catch-bonds with the carbohydrate monomannose whose strength increases...
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A Catch-Bond Based Nanoadhesive Sensitive to Shear Stress

2004 Vol. 4, No. 9 1593-1597

Manu Forero,†,| Wendy E. Thomas,‡ Clint Bland,† Lina M. Nilsson,‡,| Evgeni V. Sokurenko,*,§ and Viola Vogel*,†,‡,| Center for Nanotechnology and Departments of Bioengineering, Physics, and Microbiology, UniVersity of Washington, Seattle, Washington 98195 and Department of Materials, Swiss Federal Institute of Technology (ETH), Zu¨rich, Switzerland Received May 5, 2004; Revised Manuscript Received July 1, 2004

ABSTRACT The bacterial adhesive protein FimH forms bonds so-called catch-bonds with the carbohydrate monomannose whose strength increases under force. We introduce FimH here as a nanoadhesive that binds strongly within a characteristic force range but poorly below and above this range. We demonstrate that a mixture of adhesive-covered beads can be sorted by size in a flow chamber or can function collectively as in situ wall shear stress sensors. We also propose applications of catch-bonds for shear-selective drug delivery and shear stress mapping in biological systems.

Natural adhesives often have sophisticated functional properties that are quite distinct from synthetic adhesives. For example, the biotin-streptavidin system has been used extensively because of its high specificity, reversibility, and ease of integration into proteins and other nanoscale components.1 Mollusks bind to rocks with an adhesive that can anneal in the presence of water and over a wide range of salinity and temperature.2 Researchers have also started to develop adhesives inspired by the tiny hairs on gecko feet that bind strongly to a wide range of surfaces via van der Waals and capillary interactions.3 Just as biological adhesion displays complex regulation by outside stimuli, there are also efforts to make smart adhesives whose properties change reversibly in response to particular inputs.4,5 In this paper we introduce an adhesive that strengthens with tensile forces and explore some applications. Until recently it was generally thought that receptorligand interactions weaken with increasing tensile force.6 “Regulation” of adhesion by mechanical force was thus limited to force-induced bond weakening. However, we discovered that the adhesion between Escherichia coli and red blood cells (RBCs) can increase with force caused by shear stress.7 Bacteria exhibiting this force-activated adhesion display type I fimbriae, which are the most common structures that mediate surface attachment between E. coli * Corresponding authors. E-mail: [email protected]. Tel: +41 1 632 08 87. E-mail: [email protected]. † Department of Physics. ‡ Department of Bioengineering. § Department of Microbiology. | Department of Materials, Swiss Federal Institute of Technology. 10.1021/nl049329z CCC: $27.50 Published on Web 08/10/2004

© 2004 American Chemical Society

and its hosts. Type I fimbriae are 6-8 nm thick rod-like filaments protruding from the surface of E. coli and are polymerized from FimA monomers to a total average length of 500 nm. The tip of this rod consists of several additional subunits and terminates in FimH, which is a lectin that binds specifically to mannose residues on proteins. In our previous study, RBCs, which have exposed mannose residues, were rolling over a carpet of bacteria attached to the bottom plate of a parallel plate flow cell. When the shear applied to RBCs was increased 10-fold, the attachment of RBCs to bacteria switched from loose to firm.7 This experiment provided the first solid evidence for a possible existence of “catchbonds”,8,9 which were proposed over 15 years ago by Dembo et al.10 and whose existence was also confirmed by Marshall et al.11 with the P-selectin system at the single-bond level. Catch-bonds are defined as bonds whose lifetimes increase when tensile force is applied, instead of decreasing as is the case in the more common “slip-bonds”. Unlike the bonds in Dembo’s model, however, both the P-selectin bonds11 and the FimH-mannose bonds7 are biphasic: their adhesive strength increases with force as in the Dembo model but then decreases again at very high forces. While we previously demonstrated that the shear-enhanced adhesion of RBCs and bacteria was at least in part due to the properties of the FimH-mannose bond and derived a structural model for how mechanical forces might switch FimH adhesion to mannose from low to high affinity,7 it remained to be determined whether other viscoelastic properties of RBCs or bacteria might also contribute to the observed shear-enhanced adhesion.

Here, we present a cell-free assay and show that fimbriae terminated by the adhesin FimH and monomannose (1M) are sufficient for reproducing shear activated adhesion. We also address how the biphasic nature of catch-bond adhesion, namely that there exists an optimal range of force for which adhesion is strongest, can be exploited for technological applications. Since the drag forces acting on a particle depend not only on shear stress but also on particle size, we discuss how catch-bonds can be used either for size-dependent particle sorting, or to measure wall shear stress in microfluidic devices, or ultimately in biological systems. To first demonstrate that force-enhanced adhesion is reproducible in a cell-free fimbriae/1M system, we performed an adhesion assay with beads in a parallel plate flow chamber. The bottom plate of the chamber was coated with fimbriae purified from a recombinant bacterial strain expressing type 1 fimbriae of E. coli strain f-18 described previously.12 Polystyrene beads, 6 µm in diameter and covered with monomannosylated bovine serum albumin (1M-BSA), were injected in the chamber, allowed to attach to the surface, and tracked at varying levels of shear (see materials and methods). Figure 1a shows the detachment rates of the 6 µm beads (circles) from the fimbriae carpet as a function of shear stress. Below 0.05 pN/µm2, the detachment rate is around 0.3 min-1. As the shear stress is increased to between 0.05 and 0.4 pN/µm2, the detachment rate is reduced by more than an order of magnitude to below 0.01 min-1, consistent with the notion that the 6 µm beads are bound to the wall via catch-bonds that break less often as force increases. Finally, at very high shear stress above 0.7 pN/ µm2, the detachment rate increases dramatically to over 20 min-1. This is consistent with the idea that even catch-bonds can transition into slip bonds that are torn apart when the force is high enough11 and results in a biphasic response to force. Similar results were found when either whole bacteria were used instead of fimbriae in conjunction with the mannose beads or when RBCs were used instead of the beads over the fimbrial carpet. When 2% alpha-methyl-mannoside, which competes with mannose for the FimH binding site, was added to the buffer solution, adhesion to the wall was inhibited. Furthermore, beads did not attach to the fimbrial carpet when they were functionalized with galactose-BSA, to which fimbriae do not normally bind. This confirms that the specific force-enhanced adhesion is mediated by 1M, supporting the notion that FimH binds to 1M via catch-bonds. That is, purified fimbriae and mannose have both the force sensor and the molecular transduction element that increase adhesion in a shear-dependent manner. Therefore, effects related to the plasticity of RBCs, active fimbriae retraction,13 or additional bacterial adhesins can be discarded. The steep change in detachment rates in this cell-free assay is consistent with our earlier suggestion of a structural mechanism by which this receptor-ligand interaction is switched from low to high affinity if pulled upon by mechanical force.7 Unlike the catch-bond P-selectin, which only mediates rolling adhesion, the above beads and also bacteria14 are found to adhere firmly and become stationary on a surface at high shear stresses. Since binding is biphasic, it should 1594

Figure 1. Force enhanced adhesion of various sizes of 1M-BSA beads over fimbriae. (a) Detachment rate of 6 µm (O) and 3 µm (2) beads over a carpet of fimbriae. For shear stresses below a particular threshold the beads detach at rates around 0.3 min-1. This threshold is 0.05 pN/µm2 for the 6 µm beads and 0.2 pN/µm2 for the 3 µm beads. Above this threshold, detachment of beads is reduced significantly to under 0.01 min-1. These data show that 1M and fimbriae are sufficient to induce shear-activated adhesion. At very high shear stresses, beads detach from the surface at rates exceeding 20 min-1, demonstrating the biphasic nature of these bonds. Error bars denote 95% confidence intervals. (b) Bead size versus range of maximum adhesion. The range for which the detachment rate is under 0.01 min-1 is denoted by a box and changes with bead size. The lines labeled τ and 4.4 τ correspond to the shears used in Figure 2. Note: 1 pN/µm2 ) 1 Pa ) 10 dynes/ cm2.

then be possible to regulate this firm adhesion by mechanical force to make a force-activated nanoadhesive. The drag force, F, exerted on a bead tethered to a wall is approximately proportional to the square of the bead radius and the shear stress imparted by the fluid. An expression of the force by Goldman15 for a static bead next to a wall within the boundary layer is given by F ) 1.7005‚6πµr2S ≈ 32r2τ where µ is the viscosity of the fluid, r is the radius of the bead, S is the shear rate, and τ ) µS is the wall shear stress. As a rough estimate, noting that the velocity of the beads is small so they can be considered to be static, we expect that the ratio of force for two beads of different sizes is approximately given by the square of the ratio of the radii. It should then take 4 times the shear stress on the 3 µm beads to generate the force experienced by a large 6 µm bead at a Nano Lett., Vol. 4, No. 9, 2004

Figure 2. Separation of 1M-BSA beads by size in a flow chamber with different channel diameters with surface-adsorbed fimbriae. (a) A mixture of 3 µm beads (red) and 6 µm beads (yellow) is seeded on the surface of a fluidic chamber under no-flow conditions. (b) The fluid flows first through a low shear (τ) region and then passes through a region of high shear (4.4τ). The low shear region is 10 mm wide while the high shear region is 2.5 mm wide. Figures (c) and (d) show areas seeded as in (a) after 5 min under a shear stress of τ ) 0.1 pN/µm2 (c), and of 4.4τ ) 0.44 pN/µm2 (d). From an initial proportion of 44% small and 56 ( 8% large beads, the low shear region (a) is depleted of small beads (only 16 ( 5% left) because there is not enough force to activate their bonds. The high shear region (c) is depleted of large beads (less than 5% left) because higher shear creates sufficient force to wash them off. Videos of the separation process in both the low and high shear regions can be found in the Supporting Information.

particular shear. Figure 1a shows the detachment rates of 3 µm and 6 µm beads prepared with equal surface receptor density moving in a parallel plate flow chamber. As expected, four times more shear stress is required to induce strong adhesion of the 3 µm beads. Figure 1b graphs the range of shear stress over which beads bind tightly (detach at a rate under 0.1 min-1) for a wider range of bead sizes: 1.5, 3, 6, and 10 µm beads. The minimum force required to keep the beads from detaching is 14 ( 5 pN for bead sizes ranging between 1.5 µm and 10 µm. This number is the average for the four sets of beads and was calculated using the above formula. Experimentally this force does not increase with increased bead size. Thus, the tensile force on the bead’s tether, not the shear stress itself, is responsible for the increased adhesive properties. As illustrated in Figure 1b, this means that the level of shear stress can be tuned to selectively bind beads of a particular size. To directly illustrate the separation of particles by size using force-enhanced adhesion, we used a flow chamber with two flow regions differing in width and thus shear stress by a factor of 4.4 (Figure 2b). A solution with 3 µm and 6 µm beads was injected and allowed to settle in both regions (Figure 2a). The flow rate was chosen such that the large but not the small beads adhered maximally in the low shear region (τ) while the small but not the large beads adhere maximally to the high shear region (4.4τ) (Figure 1b). When the flow was initiated, the large beads washed away in the Nano Lett., Vol. 4, No. 9, 2004

high shear region as presumably the drag force acting on them was enough to transition into the slip-bond mode (Figure 2d and movie 1 in Supporting Information). In this high shear region the small beads fell within the optimal range of adhesion, and remained attached. Conversely, the small beads washed away in the low shear region, because the drag force was not high enough to induce their attachment, while the large beads attached maximally (Figure 2c and movie 2 in Supporting Information). This last part of the sorting would not be possible with conventional slipbonds as the small particles would also stick in the low shear region. Since the small beads attached in the high shear region, the possibility that the small beads did not attach to the low-shear region due to a low mannose concentration on the small beads surface can be discarded. This assay illustrates how a catch-bond binding mechanism can be used to direct beads of two different sizes to adhere to separate regions in a flow chamber. Conversely, this experiment also demonstrates how a mixture of beads of various sizes that all bind via forceactivated receptor-ligand interaction can be used as shear sensors to probe unknown wall shear stresses. Small beads adhere to a high shear region in which large beads detach, and large beads adhere in a low shear region where the smaller beads detach. Here bead size determines the drag force transmitted to the bonds and thus establishes a characteristic range of shear stress for maximal adhesion as in Figure 1b. In the region shown in Figure 2c, the tight binding of the 6 µm beads indicates that the shear stress must be within the optimal adhesive range of the 6 µm beads but not the 3 µm beads, or between 0.05 and 0.15 pN/µm2 according to Figure 1b. Since this flow chamber has a known geometry, we can confirm that this is correct as it includes the known value of 0.1 pN/µm2. Similarly, the region in Figure 2d binds the 3 µm but not the 6 µm beads, and thus must experience a shear wall stress between 0.4 and 4 pN/µm2, again consistent with the calculated value of 0.44 pN/µm2. While a two-bead system can distinguish between only four shear regions, i.e., by binding one, the other, both, or neither of the beads, Figure 1b demonstrates how the range of shear stress measured can be increased by adding larger and smaller beads. Utilizing this shear-activated nanoadhesive to sort particles according to their size has considerable advantages over other methods,16 including centrifugation-based or filter sorting. It can be simply integrated into “lab on a chip” devices: it does not require moving parts, power, or complicated optics. Since adhesion is reversible, both sets of beads can be recovered for further processing by reducing or increasing shear. Sequential adjustment of the flow rates could then be used to selectively wash off larger or smaller particles, while keeping particles of interest. Because 1M-terminal proteins are already expressed on many cell types, it may be possible to sort some cells without functionalizing them. Alternatively, 1M-terminal residues can be added to proteins or particles of interest. Since the force on the particles is proportional to the square of the size, selectivity should be possible over a wide range of particle sizes. For such applications the 1595

particle sizes could be directly measured by microscopic techniques or light scattering as function of their spatial position in the device. Moreover, the particles can be labeled in a size-specific manner before the experiment to distinguish different regions of shear. Catch-bond-based assays furthermore offer promising applications in biology to probe wall shear stresses. While the physics of how to calculate or simulate shear profiles of fluid flow at low Reynolds numbers through simple channels for given flow rates and geometries is common, probing or simulating wall shear stresses in more complex fluidic systems is still hampered with difficulties: biological channels are often inhomogeneous in diameter and have irregular and often poorly understood surface topographies. Furthermore, biological fluids such as blood are non-Newtonian, meaning that their viscosity changes with shear stress, and even the size and shape of the channel changes over time. The combination of these factors makes simulations difficult and compromises their accuracy.17 Experimental measurements of wall shear stress in biology have also been limited because existing shear stress sensors are microfabricated devices that are from 80 µm18 to several millimeters long19,20 and require implantation and thermal or electrical integration into the channel.20 Moreover, their presence may perturb the flow conditions and they typically measure the wall shear stress on their own surfaces, as opposed to the surface of interest. In practice, researchers have resorted to measuring fluid flow velocities instead of wall shear stress,21 but these measurements in the important near-wall regions are the least accurate. We suggest here that a cocktail of beads with different diameters that bind through catch-bonds can serve as wall shear stress sensors in complex biological systems. To map wall shear stresses in the cardiovascular or urinary track system, for example, advantage could be taken of the fact that endothelial cells already present 1M on their cell surfaces. Consequently, beads covered with fimbriae could be used to probe wall shear stresses under in-situ conditions. Figure 3 shows the detachment rate of fimbriae-covered beads over a mannose surface, and comparison of this with Figure 1a demonstrates that whether the fimbriae are on the beads or on the channel’s surface does not affect the necessary properties for probing shear stresses. The response time of the 1M-FimH system to alterations of the flow conditions, i.e., the time it takes to switch this catch-bond from low to high binding, or vice versa, is within seconds.14 This sort of shear stress sensor also has the ability to target particular areas in a fluidic system. For example, regions of the cardiovascular system with particular shear stresses, such as vessels partially occluded by atherosclerotic plaques, could be specifically targeted by therapeutic drug-containing particles. We thus introduce the concept of shear-sensitive “smart” adhesives and propose applications in microfluidic devices and biological systems. These applications exploit the catchbond properties of the mannose-sensitive adhesins of E. coli such as firm adhesion at high shear and biphasic response. Materials and Methods. Reagents. 1M-BSA and galactosylated BSA were obtained from EY laboratories, Inc. (San 1596

Figure 3. Detachment rate of fimbriae-covered 6 µm beads over a 1M-BSA surface. Beads detach often at low shears, attach well at high shears, and detach completely at very high shears. This assay was conducted as described in Figure 1a but with the roles of 1M-BSA and fimbriae inverted. This setup can be used to measure shear stresses in biological systems where 1M is presented naturally by cells.

Mateo, CA). Polystyrene microspheres were obtained from Polysciences, Inc. (Warrington, PA). All other reagents were obtained from Sigma (St. Louis, MO). Bacterial Strains and Fimbriae. Fimbriae are sheared off by a homogenizer, followed by differential centrifugation and MgCl2 precipitation as described previously.12 The E. coli strain used was recombinant expressing F-18 FimH adhesin as described previously.7 Physisorption of Fimbriae and 1M-BSA to PS Plates. 35 mm Corning (#430165) tissue culture dishes were incubated either with purified fimbriae at 1 µg/mL in 0.02 M bicarbonate buffer or with 1M-BSA at 150 µg/mL in 0.02 M bicarbonate buffer at 37 °C for 1 h, and then washed thrice in PBS with 0.2% BSA (PBS-BSA) to prevent nonspecific adhesion by the beads to the dish. Bead Coating. Polystyrene microspheres of various sizes were prepared by rotating a solution of 50 µL of 2.6% beads mixed with 150 µL of 20 to 200 µg/mL 1M-BSA in 0.02 M bicarbonate buffer for 1 h at room temperature. A solution of fimbriae at 10 µg/mL was used instead of 1M-BSA to obtain fimbriated beads. The beads were then spun twice and resuspended in fresh PBS-BSA. They were finally diluted down to 0.1% and injected in the chamber. Parallel Plate Flow Chamber Experiments. The coated dishes above served as the bottom plate in a parallel plate flow chamber from Glycotech #31-0001 (Rockville, MD) using a silicon rubber gasket 20 mm long, 2.5 mm wide, and 250 µm thick. The fluid (PBS-BSA) was pumped through the chamber by a #975 pulse-free syringe pump from Harvard Apparatus, Inc. (Holliston, MA). The movement of the beads was recorded using an inverted Nikon TE 200 microscope with a long working distance 10× phase contrast objective by means of a Roper Scientific (Duluth, GA) Cascade CCD camera. For the separation assay a 125 µm thick rubber gasket was cut to 11 mm wide at one end and kept 2.5 mm wide on the other (Figure 2b). Data Analysis. Images of the flow chamber were recorded every half-second for 3 min at each shear rate. Bead positions Nano Lett., Vol. 4, No. 9, 2004

were tracked every frame using the point-tracking plug-in from Metamorph video imaging software by Universal Imaging Corp. (Downingtown, PA). Detached cells or beads were defined as those that flowed at over 2/3 the expected free flowing velocity at some point during the lapse of the recording. The expected free flowing velocity was calculated by multiplying the shear rate by the radius of the particle, assuming that it is barely touching the surface. Detachment rates k of beads were calculated from the fraction X remaining after t ) 3 min, assuming an exponential decay, using the formula k ) 1/t ln(1/X). Acknowledgment. We thank E. Tritchina for early technical support, H. Hess for valuable discussions, and M. McCloskey for comments. The research was supported by the NIH Bioengineering Research Partnerships (BRP) grant # 1 R01 AI 50940 (E.V.S. and V.V.); NIH Center of Excellence in Genome Sciences, Micro Life Science Center grant P50, HL002360; a Whitaker Foundation Graduate Fellowship (W.E.T.); a NSF Graduate Research Fellowship (L.N.); and the NSF, IGERT, and UIF award in Nanotechnology Fellowship at the University of Washington (M.F.). Supporting Information Available: Videos of the separation process in both the low and high shear regions. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Stayton, P. S.; Nelson, K. E.; McDevitt, T. C.; Bulmus, V.; Shimoboji, T.; Ding, Z.; Hoffman, A. S. Biomol. Eng. 1999, 16, 93-99.

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(2) Deming, T. J. Curr. Opin. Chem. Biol. 1999, 3, 100-105. (3) Geim, A. K.; Dubonos, S. V.; Grigorieva, I. V.; Novoselov, K. S.; Zhukov, A. A.; Shapoval, S. Y. Nat. Mater. 2003, 2, 461-463. (4) Khongtong, S.; Ferguson, G. S. J. Am. Chem. Soc. 2002, 124, 72547255. (5) Crevoisier, G. B.; Fabre, P.; Corpart, J. M.; Leibler, L. Science 1999, 285, 1246-1249. (6) Evans, E. Annu. ReV. Biophys. Biomol. Struct. 2001, 30, 105-128. (7) Thomas, W. E.; Trintchina, E.; Forero, M.; Vogel, V.; Sokurenko, E. V. Cell 2002, 109, 913-923. (8) Isberg, R. R.; Barnes, P. Cell 2002, 110, 1-4. (9) Konstantopoulos, K.; Hanley, W. D.; Wirtz, D. Curr. Biol. 2003, 13, R611-613. (10) Dembo, M.; Torney, D. C.; Saxman, K.; Hammer, D. Proc. R. Soc. London B Biol. Sci. 1988, 234, 55-83. (11) Marshall, B. T.; Long, M.; Piper, J. W.; Yago, T.; McEver, R. P.; Zhu, C. Nature 2003, 423, 190-193. (12) Sokurenko, E. V.; Courtney, H. S.; Ohman, D. E.; Klemm, P.; Hasty, D. L. J. Bacteriol. 1994, 176, 748-755. (13) Maier, B.; Potter, L.; So, M.; Seifert, H. S.; Sheetz, M. P. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 16012-16017. (14) Thomas, W. E.; Nilsson, L.; Forero, M.; Sokurenko, E. V.; Vogel, V. Mol. Microbiol., in press. (15) Goldman, A. J.; Cox, R. G.; Brenner, H. Chem. Eng. Sci. 1967, 22, 653-660. (16) MacDonald, M. P.; Spalding, G. C.; Dholakia, K. Nature 2003, 426, 421-424. (17) Frank, A. O.; Walsh, P. W.; Moore, J. E., Jr. Artif. Organs 2002, 26, 614-621. (18) Hsiai, T. K.; Cho, S. K.; Wong, P. K.; Ing, M.; Salazar, A.; Sevanian, A.; Navab, M.; Demer, L. L.; Ho, C. M. FASEB J. 2003, 17, 16481657. (19) Fernholz, H. H.; Janke, G.; Schober, M.; Wagner, P. M.; Warnack, D. Meas. Sci. Technol. 1996, 7, 1396-1409. (20) von Papen, T.; Steffes, H.; Ngo, H. D.; Obermeier, E. Sens. Actuators A 2002, 97-98, 264-270. (21) Wootton, D. M.; Ku, D. N. Annu. ReV. Biomed. Eng. 1999, 1, 299-329.

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