Micromotors Powered by Enzyme Catalysis

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1 Supporting Information Micromotors Powered by Enzyme Catalysis Krishna K. Dey, 1 Xi Zhao, 1, Benjamin M. Tansi, 1, Wilfredo J. Méndez-Ortiz, 2 Ubaldo M. Córdova-Figueroa 2, * Ramin Golestanian, 3, * and Ayusman Sen, 1, * 1 Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania 16802, USA. 2 Department of Chemical Engineering, University of Puerto Rico-Mayagüez, Mayagüez, PR 00681, Puerto Rico, USA. 3 Rudolf Peierls Centre for Theoretical Physics, University of Oxford, Oxford OX1 3NP, UK. * asen@psu.edu (AS), ramin.golestanian@physics.ox.ac.uk (RG), ubaldom.cordova@upr.edu (UMCF) These authors contributed equally S1

2 Coating of Streptavidin-functionalized Polystyrene Microspheres with Biotinylated Enzymes EZ-Link-Maleimide-PEG2-Biotin (Life Technologies) was used to tag thiol groups of urease and thereby binding the enzymes in specific and unobtrusive manner. This reaction with EZ-Link- Maleimide-PEG2-Biotin occurs at a ph between 6.5 and 7.5. To tag catalase, EZ-Link-Sulfo- NHS-Biotin (Life Technologies) was used. This reacts with free primary amines in the enzyme structure. The tagging was performed by mixing enzyme and biotin solutions, both diluted in 100 mm phosphate buffer. A 4:1 enzyme-biotin ratio was maintained. Prior to synthesis, we estimated the concentration of enzyme required to achieve complete monolayer coverage of the microparticles. An excess of enzyme was typically employed. The enzyme-biotin solution was left for 2 hours in a mechanical shaker (speed 500 rpm) in order to allow the reaction to be completed. A calculated volume of streptavidin-functionalized polystyrene beads suspension was mixed with the enzyme-biotin solution and placed at 4 C for 30 min followed by 30 min in mechanical shaker (speed 500 rpm) at room temperature. To remove excess enzymes, the suspension of functionalized particles was centrifuged at 4500 rpm for 7 min for four successive cycles, re-dispersing the particles in 10 mm phosphate buffer after each cycle. The final solution was adjusted to approximately 500 μl by adding fresh 10 mm buffer. For chemotaxis experiments, fluorescent streptavidin-functionalized polystyrene particles of diameter 2 μm (Fluoresbrite YG Microspheres, Polysciences, ex/em = 441/486) were coated with catalase and urease following the same protocol. Larger beads however required higher concentration of enzymes. S2

3 A B Figure S1. Fluorescence and optical microscopy images of (A) 2 µm streptavidin-functionalized polystyrene particles coated with tagged biotinylated catalase and (B) uncoated 2 µm streptavidin-functionalized polystyrene particles. Optical Microscopy Studies For catalase and urease coated beads, we used buffered solutions of 100 mm H2O2 and 1 M urea as stocks. To investigate the motion of enzyme-coated micromotors, a suspension of enzymeattached polystyrene particles was added to an appropriate mixture of buffer and substrate stock solutions which was then used to fill a secure seal hybridization chamber (20 mm diameter, 1.3 mm height, Grace, Bio Labs) forming a quasi-two dimensional layer of liquid. The chamber was placed on top of a micro glass slide. This created uniform layer of solution with no convective flows. The diffusion of the enzyme coated particles was measured using a Zeiss-Axiovet 200 S3

4 inverted optical microscope with Flea 3 USB video camera (FL3-U3-32S2C-CS) from Point Grey. The recorded videos of the particles were analyzed using codes developed in LabVIEW and Vision Development Module (National Instruments). The videos were recorded at a height of approximately 50 µm from the glass surface, only when the motion of the particles was found to be free from convection. Videos were all recorded with 20X objective and the mean square displacements with time were fitted linearly to extract diffusion coefficients of the micromotors in two dimensions. The average these diffusion coefficients was calculated and the entire procedure was repeated six times to estimate the standard deviations in the measurements. For fixed amount of enzyme coated particles, the composition of the experimental solutions was changed by varying the volume of 10 mm phosphate buffer and amount of buffered substrate solution, keeping the total volume always the same. Diffusion Coefficient Calculation using LabVIEW Experimental videos were analyzed for mean square displacements and particle diffusion using codes developed in LabVIEW and Vision Development Module (National Instruments). Videos were analyzed for x and y displacements of particles at various times t. Mean square displacement for each particle was then plotted against time and fitted linearly to calculate the slope. For motion in 2D, the slope of the mean square displacement vs. time plot was equal to four times the diffusion coefficient of the particle. Average of diffusion coefficients calculated for all the particles in a video gave the result for a single measurement. Diffusion experiments were repeated six independent times to calculate the mean and standard deviations reported in Figure 1 in the manuscript. S4

5 Representative Mean Square Displacement Plots for Catalase and Urease Coated Particles Figure S2. Representative mean square displacement plots for (A) catalase and (B) urease coated beads in 10 mm phosphate buffer and 4.4 mm H2O2/170mM urea. To generate these plots we averaged the MSDs calculated for all the particles in a video and plotted the average value against time. The plots clearly show the enhancement in the (effective) diffusion coefficients. Enhanced Diffusion of 2 µm Streptavidin-functionalized Polystyrene Particles Coated with Catalase S5

6 Figure S3. Diffusion of 2 µm streptavidin-functionalized polystyrene particles coated with catalase in 10 mm PBS and 10 mm buffered solution of H2O2. In the presence of H2O2, we observed a 5% increase in diffusion of the particles. Dynamic Light Scattering Measurements The DLS measurements were carried out in a Malvern Zetasizer ZS instrument, which used noninvasive backscatter optics (NIBS) to measure particle size of suspensions. Lower concentrations of enzyme coated particles were used in DLS measurements to reduce uncertainties in measurements due to aggregation. The measurements were all carried out at 25 0 C, considering the particle refractive index and absorbance to be 1.59 and 0.01 respectively. Calculation of Enzyme Concentration in the Experimental Solution We consider 125 μl of 1.36% w/v microsphere stock solution, which contains approximately 1.7 mg of solid. Now, radius of one bead is 6 ~ m. Therefore, volume ( ) = πr = π m = m 3. Considering the density of the polystyrene to be 6 3 = gm/m, S mass of one bead is = ( ) ( ) gm = gm. Thus, the number of beads in 125 μl microsphere solution = g g 9 13 = Assuming no significant loss of beads during enzyme functionalization, the final solution contains beads dispersed in approximately 500 ul phosphate buffer. Now, in optical microscopy experiments, 10 µl of the coated bead solution is dilute to ~500 µl (by mixing with substrate and buffer) and is used as the final experimental solution. Therefore, ( ) approximate number of bead in the final solution = = S6

7 Now, surface area of each bead ( ) = =. Cross sectional 4 πr = 4π m m area of one streptavidin molecule ( ) = =. S2 Therefore, πr = π 2 10 m m m 5 number of streptavidin molecules on one bead = = Two biotinylated m enzymes can be attached to each streptavidin molecule. S3 Therefore, total number of enzyme molecules needed to completely coat one bead is 5 = Therefore, in the experimental chamber, approximate number of enzyme molecules = = Therefore, approximate concentration of enzymes in the experimental solution = = 0.13 μm Microfluidic Channel Fabrication Microchannel masters were fabricated over silicon wafers in the Nanofabrication Laboratory of Materials Research Institute at Penn State. Before photolithography, wafers were cleaned with acetone and then air-dried properly. These were then spin-coated with 5 ml of SPR-220 photoresist (Microposit) at 200 rpm for 10 s and then at 1000 rpm for 30 s. The coated wafers were then soft-baked over a hot plate at C for 5 min. The microchannel geometry was modelled in CAD and printed over a chrome-on-glass mask (Nanofabrication Laboratory, Materials Research Institute, Penn State). The mask was placed in contact with the photoresist over the wafers and the resist was exposed to UV radiation for 40 s in a Karl Suss MA/BA6 Contact Aligner. This was then followed by post-baking the wafer for 5 min over a hot plate at C to cross link the exposed film. To remove unexposed SPR 220 from the wafers, the mold S7

8 was developed in CD26 developer for 2 min while being agitated, followed by washing thoroughly with deionized water. The wafers were then dried with a nitrogen blower followed by creation of a 100 µm deep master pattern over them using deep reactive ion etching facilities (Alcatel). Polydimethylsiloxane (PDMS, SylgardTM 184, Dow Corning) elastomer solution was prepared by mixing pre-polymer with a cross-linking agent in weight ratio of 10:1. To minimize adhesion of PDMS to the surface, the wafers were exposed to trichloro (1H, 1H, 2H, 2H- perfluorooctyl) silane (Sigma Aldrich) before the PDMS solution was poured over them. The PDMS solution was poured on top of the wafers to the desired thickness. This was then degassed in vacuum for about an hour to remover air bubbles from the PDMS mixture. The masters were then heated in an oven at 60 C for 2 h. After curing, the PDMS channels were peeled from the mold and inlets/outlets were made using a stainless steel puncher. The devices were sealed to glass coverslips (VWR) by exposing them to oxygen plasma, followed by bonding them together manually and baking them on a hot plate at C for 2 min. The PDMS channels were connected to polyethylene tubes (SPC Technology, Internal diameter 0.66 mm) and fluid flow through them was controlled using syringe pumps. References S1. Huang, R.; Chavez, I.; Taute, K. M.; Lukić, B.; Jeney, S.; Raizen, M. G.; Florin, E-L. Nat. Phys. 2011, 7, S2. Jurchenko, C.; Chang, Y.; Narui, Y.; Zhang, Y.; Salaita, K. S. Biophys. J. 2014, 106, S3. Sabanayagam, C. R.; Smith, C. L.; Cantor, C. R. Nucl. Acid. Res. 2000, 28, e33. S8