Aptamer-Based Polyvalent Ligands for Regulated Cell Attachment

Stacy Jones , Ye Gao , Carrie Sweet , Aruna Vangara , Salma Begum , Paresh Chandra Ray ... Jonah C Rosch , Emma K Hollmann , Ethan S Lippmann...
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Aptamer-Based Polyvalent Ligands for Regulated Cell Attachment on the Hydrogel Surface Erin R. Gaddes, Gregory Gydush, Shihui Li, Niancao Chen, Cheng Dong, and Yong Wang* Department of Biomedical Engineering, Pennsylvania State University, University Park, Pennsylvania 16802-6804, United States S Supporting Information *

ABSTRACT: Natural biomolecules are often used to functionalize materials to achieve desired cell-material interactions. However, their applications can be limited owing to denaturation during the material functionalization process. Therefore, efforts have been made to develop synthetic ligands with polyvalence as alternatives to natural affinity biomolecules for the synthesis of functional materials and the control of cell-material interactions. This work was aimed at investigating the capability of a hydrogel functionalized with a novel polyvalent aptamer in inducing cell attachment in dynamic flow and releasing the attached cells in physiological conditions through a hybridization reaction. The results show that the polyvalent aptamer could induce cell attachment on the hydrogel in dynamic flow. Moreover, cell attachment on the hydrogel surface was significantly influenced by the value of shear stress. The cell density on the hydrogel was increased from 40 cells/mm2 to nearly 700 cells/mm2 when the shear stress was decreased from 0.05 to 0.005 Pa. After the attachment onto the hydrogel surface, approximately 95% of the cells could be triggered to detach within 20 min by using an oligonucleotide complementary sequence that displaced polyvalent aptamer strands from the hydrogel surface. While it was found that the cell activity was reduced, the live/dead staining results show that ≥98% of the detached cells were viable. Therefore, this work has suggested that the polyvalent aptamer is a promising synthetic ligand for the functionalization of materials for regulated cell attachment.

1. INTRODUCTION Biomolecules, such as antibodies and growth factors, are capable of binding to cell receptors with high affinities and specificities. They are often utilized in the functionalization of materials to impart strong and specific cell-material interactions1,2 for applications such as tissue engineering,3 biological separations,4,5 and biosensor development.6,7 However, natural biomolecules are generally fragile, which can lead to their denaturation during chemical processing or the material functionalization process. In an effort to create more robust affinity materials, synthetic ligands8,9 have been explored as potential substitutes for natural biomolecules. Of these synthetic ligands, nucleic acid aptamers have recently received great attention.10,11 Different from most natural biomolecules, they can recover their molecular recognition functionality12 after being processed in harsh chemical or physical conditions.13−15 However, aptamer-based molecular recognition is a monovalent binding event. Since polyvalent interactions play an important role in biological systems,16 it is desirable to synthesize polyvalent aptamers to enhance cell-material interactions. Polyvalent aptamers can be synthesized chemically via sequential conjugation of nucleotides using the standard phosphoramidite method.17−20 For instance, this method has been used to synthesize a bivalent aptamer for labeling CD62Lpositive lymphocytes.20 This bivalent aptamer demonstrated a superior binding capability compared to a monovalent control. © 2015 American Chemical Society

While the phosphoramidite method is simple and allows the incorporation of chemically modified oligonucleotides,21 the efficiency of the chemical synthesis process is limited to producing approximately 60 nucleotides in general,22,23 with the yield and quality of synthesis dramatically reduced beyond this length. Therefore, efforts have been made to seek new methods for the synthesis of polyvalent aptamers.24−28 Recently, we synthesized a novel polyvalent aptamer using short oligonucleotides and applied it to functionalize a hydrogel surface.29 The backbone of this polyvalent aptamer was synthesized via the hybridization chain reaction method.30 The backbone was further hybridized with aptamers as side units to acquire polyvalence for binding target cells.29 Polyvalent aptamers were able to enhance cell attachment on the hydrogel surface in situations where either ligand density or surface reaction sites were equal.29 However, we did not study whether the attached cells can be regulated to detach from the hydrogel surface. Additionally, we did not study whether cells can bind to the hydrogel surface in dynamic flow. These two questions are important for applications, such as cell separation, that often require cell attachment in dynamic flow for cell enrichment and cell detachment for downstream cell analysis.31−33 Received: February 6, 2015 Revised: March 16, 2015 Published: March 19, 2015 1382

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channels of the sensor chip at 10 μL/min, 37 °C, for predetermined amounts of time. Data points were recorded at 0.5 s intervals. 2.4. Hydrogel Preparation and Synthesis. For static cell capture, thin hydrogel films were formed on a silanized glass square. Glass microscope slides (VWR, Radnor, PA) were cut prior to silanization in 4 mm × 4 mm squares. These squares were sonicated in acetone for 15 min, rinsed with DI water, and then sonicated in 1 M NaOH for 10 min. After rinsing in DI water and drying completely, clean squares were immersed in silanization solution for 5 min. This solution consisted of 50 mL of ethanol, 0.5 mL of 3-(trimethoxysilyl) propyl methacrylate, and 1.5 mL of 10% glacial acetic acid in deionized water. Silanized glass squares were then rinsed with ethanol, air-dried in a ventilated chemical hood, and stored in a desiccator at room temperature until use. For dynamic cell capture, a larger 37.5 × 50 mm glass microscope slide base (Corning, Corning, NY) was silanized according to the procedure used for the glass squares. To create hydrogel films for static cell studies, 1 μL of 10% acrylamide was mixed with Acrydite-modified adapter oligonucleotides at a final concentration of 10 μM. To this solution, 0.15 μL of 10% (w/ v) ammonium persulfate and 0.15 μL of 5% (v/v) TEMED were added. Immediately, 1 μL of the solution was pipetted onto a clean glass slide and covered with a silanized glass square. To create hydrogel films for dynamic studies, 2.56 μL of 10% acrylamide was mixed with Acrydite-modified adapter oligonucleotides at a predetermined final concentration. To this solution, 0.47 μL of 10% (w/v) ammonium persulfate and 0.47 μL of 5% (v/v) TEMED were added. Immediately, 3.5 μL of the solution was pipetted onto the center of a large silanized glass slide and covered with an unsilanized 2.5 cm × 0.25 cm glass slide. In both static and dynamic hydrogel cases, polymerization occurred over a 1 h period at 37 °C, after which the silanized glass was removed from the unsilanized glass. A thin, polyacrylamide film remained the surface of the silanized glass. Hydrogels were washed in DPBS to remove any unincorporated molecules. 2.5. Surface-Based DNA Polymerization. All oligonucleotides used to create DNA polymers were adjusted to appropriate concentrations in DPBS and annealed prior to use. For the annealing procedure, a Bio-Rad T100 Thermal Cycler (Hercules, CA) was used to heat samples to 95 °C for 3 min, followed by a 1 h period for sequences to form energetically favorable secondary structures at 25 °C. Acrydite-adapter-functionalized films were incubated in a solution of 10 μM DI (with no Acrydite group) for 30 min at 37 °C. After washing the samples in DPBS, the samples were incubated in a solution of 10 μM DM1 and DM2 for 1 h at 37 °C. Following a washing step in DPBS, DNA polymer-functionalized films were incubated in a 10 μM aptamer solution for 30 min at 37 °C, forming polyvalent aptamer-functionalized films. For pH and temperature studies, hydrogels were incubated in DPBS at predetermined pH and temperatures for 1 h prior to imaging. 2.6. Imaging and Microscopy. Films with fluorescently labeled monomers and aptamers, and samples with fluorescently stained cells were imaged under an Olympus IX73 inverted microscope equipped with an Olympus U-HGLGPS fluorescence illumination source and an Olympus XM10 camera (Shinjuku, Tokyo, Japan). Cell images were acquired using phase microscopy and cells were counted using ImageJ software. For image acquisition and fluorescence pseudocoloring, CellSens Standard software was used. All error bars represent standard deviation of the mean. 2.7. Cell Culture. A CCRF-CEM lymphoblastic T-cell line (ATCC, CCL-199, Manassas, VA) was cultured in RPMI 1640 cell media (ATCC, Manassas, VA) supplemented with 10% fetal bovine serum and 100 IU/mL penicillin/streptomycin (Thermo-Fisher Scientific, Waltham, MA) and incubated at 37 °C, 5% CO2, in 95% humidity. For the Ramos B-cell line (ATCC, CRL-1596, Manassas, VA), culture conditions were the same as CCRF-CEM cells, except the media contained inactivated FBS. 2.8. Static Cell Attachment. Prior to cell capture, CCRF-CEM cells were prepared at a concentration of 5 × 105 cells/mL in binding buffer. Binding buffer was DPBS supplemented with 10 mM MgCl2, 4.5 g/L glucose, and 0.1% (w/v) BSA. For cell binding, aptamer-

The purpose of this work was to address these questions. We applied a complementary sequence (CS) to trigger the dissociation of the polyvalent aptamer and attached cells from the hydrogel surface. Triggered dissociation of polyvalent aptamers was examined by using both surface plasmon resonance and fluorescence imaging. Cell attachment on the hydrogel surface in dynamic flow was investigated by varying the values of shear stress in a parallel plate flow system and by altering the seeding density of polyvalent aptamers on the hydrogel surface. Moreover, the cell binding kinetics and specificity were examined under dynamic flow. Since cell viability is an important index to various applications, the viability of the cells after triggered detachment was further analyzed via staining techniques and activity assays.

2. MATERIALS AND METHODS 2.1. Reagents and Materials. Acetone, reagent alcohol (91% ethanol), glacial acetic acid, acrylamide/bis(acrylamide), ammonium persulfate, tetramethylethylenediamine (TEMED), Dulbecco’s phosphate buffered saline (DPBS), glycerol, tris borate ethylenediaminetetraacetic acid (TBE), hydrogen chloride, bovine serum albumin (BSA), fetal bovine serum (FBS), N-hydroxysuccinimide (NHS), 1-ethyl-3-(3(dimethylamino)propyl)carbodiimide hydrochloride (EDC), sodium acetate, and penicillin/streptomycin were purchased from ThermoFisher Scientific (Waltham, MA). Sodium hydroxide, 3-(trimethoxysilyl) propyl methacrylate, calcium chloride, HEPES, sodium chloride, Tween 20, and glucose were obtained from Sigma-Aldrich (St. Louis, MO). Magnesium chloride, RPMI 1640 media, Ramos lymphocyte B cells, and CCRF-CEM lymphocyte T cells were purchased from ATCC (Manassas, VA). All oligonucleotides were purchased from Integrated DNA Technologies (Coralville, IA). Glass microscope slides for silanization were obtained from VWR (Radnor, PA) and Corning (Corning, NY), while glass microscope slides for hydrogel preparation bases were from Thermo-Fisher Scientific (Waltham, MA). 2.2. Gel Electrophoresis. Prior to polymerization and electrophoresis, all oligonucleotides were adjusted to desired concentrations in DPBS and annealed for 3 min at 95 °C using a Bio-Rad T100 Thermal Cycler (Hercules, CA), followed by 1 h of cooling to room temperature. Adapter and DNA initiator (DI) sequences were incubated at a 1:2 DI to adapter molar ratio for 1 h at 37 °C to form DI complexes. Complexes and monomers were mixed at a 1:10 DI to monomer molar ratio for 1 h at 37 °C to enable polymerization. Samples were loaded into the wells of a 1% (w/v) agarose gel and run for 50 min at 80 V in 1× TBE running buffer. SYBR Safe DNA stain was used to visualize double-stranded DNA (green). A CRI Maestro in vivo imaging system (Woburn, MA) was used to fluorescently image gels. The subsequent images were processed with Maestro 3.0.0 software. 2.3. Surface Plasmon Resonance. The polymerization and triggered depolymerization of DNA was monitored in real time using a Reichert Technologies SR7500DC spectrometer, equipped with a SR8100 autosampler (Depew, NY). A planar polyethylene glycol/ carboxyl sensor chip (Reichert Technologies) was amine-coupled with neutravidin (Reichert Technologies) via EDC/NHS chemistry. Briefly, a 350 mM solution of NHS and EDC was prepared in deionized water and incubated on the sensor chip surface for 30 min. Next, 30 μg/mL neutravidin was prepared in 10 mM sodium acetate buffer (pH 5.6 with HCl) and incubated on the sensor chip for 2 h in humidity. The neutravidin-functionalized sensor chip was used immediately after cross-linking. For SPR experiments, running buffer consisted of 1× PBS and 0.05% (v/v) Tween 20, which was degassed for 30 min prior to use. All DNA solutions were prepared using running buffer. Biotinylated adapter sequences at 1 nM were run over the sensor chip at 10 μL/min for 30 min on the analyte channel. The DI (0.01 μM), DNA monomer 1 (DM1; 2.5 μM), DNA monomer 2 (DM2; 2.5 μM), DM1 + DM2 (2.5 μM), aptamer (1.5 μM), CS (2 μM), and scrambled complementary sequence (scrCS; 2 μM) solutions were run over both 1383

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Biomacromolecules functionalized samples were incubated with 800 μL CCRF-CEM solution for 90 min at 37 °C, 5% CO2, in 95% humidity. To remove loosely bound cells, samples were gently washed for 2 min at 90 rpm. 2.9. Triggered Cell Detachment. The trigger sequence (CS) was prepared at predetermined concentrations in DPBS and annealed using a Bio-Rad T100 Thermal Cycler (Hercules, CA) at 95 °C for 3 min and cooling the solution to 25 °C for 1 h. Samples were placed in CS solution for 20 min at 37 °C, after which they were imaged. For cell release experiments, samples were imaged using phase microscopy, while fluorescently labeled DNA was imaged using fluorescence microscopy techniques. 2.10. Regulated Cell Attachment in a Parallel Plate Flow Chamber. For dynamic cell capture, aptamer-functionalized hydrogels were used as the base of a parallel plate flow chamber. The flow chamber (Glycotech, Gaithersburg, MD) was assembled with a 0.005 in. thick gasket (Glycotech) separating the chamber from the hydrogel base. Vacuum was applied to seal the chamber to the base. Solutions were infused at predetermined flow rates using a Harvard Apparatus Pump 11 and Pump 33 syringe pump (Harvard Apparatus, Holliston, MA). For higher quality video studies, a Nikon Eclipse TE2000-U microscope equipped with a Nikon TE2 PS100W power source (Nikon, Japan) and a CCD camera (pco.1600, Cooke Corp., Romulus, MI) was used. Phase contrast images were acquired over a 3.5 min time span at a frame rate of 30 fps with NIS Elements Advanced Research imaging software and converted to Audio Video Interleave files for offline analysis. For fluorescence images of mixed Ramos and CCRF-CEM cells, CCRF-CEM cells were prestained with 25 μM CellTracker Blue CMAC dye (Thermo-Fisher Scientific, Waltham, MA) for 30 min and transferred into dye-free media with unstained Ramos cells (total concentration 5 × 105 cells/mL). Fluorescence (353 nm ex./466 nm em.) and phase images of captured cells were taken under an Olympus IX73 inverted microscope. 2.11. Cell Viability Staining and Activity Analysis. To characterize the viability and activity of released cells, two methods were used: a live/dead staining assay and an MTS assay. For both assays, released cells were counted and resuspended in media at equal cell concentrations for a final volume of 100 μL/sample. For live/dead staining, a Live/Dead Mammalian Cell Viability/Cytotoxicity kit (Thermo-Fisher Scientific, Waltham, MA) was utilized with each fluorescent dye at a 1 μM concentration. Samples were incubated with dyes at 37 °C for 15 min prior to imaging. Cells were imaged using fluorescence microscopy, where green fluorescence indicates live cells and red fluorescence identifies dead cells. For activity analysis, a CellTiter Cell Proliferation Assay (Promega, Fitchburg, WI) was utilized. Samples were seeded in a 96-well plate at 100 μL/well and 20 μL MTS reagent was added and mixed in each well. Samples were incubated at 37 °C for 3 h prior to analysis. After incubation, the absorbance of each sample at 490 nm was determined using a Tecan Infinite M200 Pro microplate reader (Tecan Group Ltd., Männedorf, Switzerland). The absorbance of each sample is directly proportional to the activity of live cells in the samples. Thus, the absorbance of released cells was compared to that of control cells (at the same seeding concentration) to determine the cell activity percentage of each sample.

Figure 1. Formation and CS-mediated dissociation of polyvalent aptamers. (A) Schematic illustrating the formation of the polymer backbone, the hybridization of aptamers to form polyvalent aptamers, and the CS-triggered molecular dissociation. (B) SPR sensogram demonstrating the polymerization and CS-triggered dissociation of polyvalent aptamers. The black arrow indicates the injection of DM1 and DM2, the blue arrow shows aptamer injection, and the red arrow marks the injection with either CS (red line) or scrambled CS (scrCS, purple line).

can hybridize with aptamers to generate polyvalence. This synthesis method is different from other methods such as the standard phosphoramidite method34,35 and the rolling circle replication method.36,37 The monomers used in both the standard phosphoramidite method and the rolling circle replication method are single nucleotides. Moreover, rolling circle replication requires enzymes that may not be able to efficiently incorporate chemically modified nucleotides. By contrast, the monomers used in this method are short oligonucleotides. The synthesis does not require conjugation chemistry or enzymes. Thus, the synthesis method presented in the current work is flexible and easy to operate on a substrate. To achieve the detachment of the polyvalent aptamer from a substrate, an overhanging toehold region was included at the base of the DNA backbone to bind a detachment-triggering CS (Figure 1A, red; Table 1). Thus, it is thermodynamically favorable for CS to detach the polyvalent aptamer from the surface (Figure 1A). Surface plasmon resonance (SPR) was first used to examine the reversible attachment and detachment of polyvalent aptamers on a substrate (Figure 1B). The first and largest signal increase shows the sequential hybridization of the monomers (black arrow); and the second, smaller signal increase shows aptamer hybridization with the DNA polymers to form polyvalent aptamers (blue arrow) at a density of approximately 40 pg/mm2. Importantly, a large decrease in signal was observed when the CS solution was passed over the polyvalent aptamers (Figure 1B, red arrow), whereas minimal signal loss was observed upon injection with a control scrambled CS (scrCS). These data clearly show that the polyvalent aptamer can be synthesized on a substrate and its display on the substrate can be effectively reversed by using CS. 3.2. Characterization of the Function of CS in Triggered Cell Detachment. Next, a study was designed to

3. RESULTS AND DISCUSSION 3.1. SPR Evaluation of Polyvalent Aptamer Synthesis and Triggered Dissociation. The synthesis of the polyvalent aptamer involves two steps: synthesis of the polymer backbone and hybridization with aptamers. The backbone of the polyvalent aptamer is formed from two DNA monomers, DM1 and DM2 (Figure 1A, orange and purple, respectively; Table 1), which sequentially hybridize to form polymers upon initiation from a short DI oligonucleotide, DI (Figure 1A, green; Table 1). The formation of DNA polymers in the presence of DI is shown in the electrophoretic gel image in Figure S1. A toehold region on DM1 provides a hybridization region for a ligand with a complementary binding region, which 1384

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Biomacromolecules Table 1. List of Oligonucleotide Sequences

Figure 2. Regulation of cell attachment using polyvalent aptamers and CS. (A) Schematic, representative fluorescence images of polyvalent aptamers, and representative phase images of cells bound to polyvalent aptamer surfaces before and after CS treatment in static conditions. Aptamers were labeled with FAM (green). scrCS indicates the scrambled CS as a control. (B) Quantitative comparison of hydrogels before and after CS treatment to show levels of fluorescent polyvalent aptamer dissociation (left graph) and cell attachment (right graph). Data were normalized to samples in buffer. (C) Effect of the toehold length on CS-triggered cell detachment. CS-5 has five nucleotides in the toehold; CS-10 has 10 nucleotides in the toehold. All error bars represent standard deviation (n = 3); *P ≤ 0.05; N.S. indicates no significant difference.

and cells remained attached to the hydrogels (Figure 2A,B). In contrast, CS treatment resulted in a statistically significant decrease in both fluorescence and the number of attached cells (Figure 2A,B). We also found that CS could trigger the dissociation of DNA polymers at different conditions when pH and temperature were varied and that DNA polymers exhibited highest stability at pH of 7.4 (Figure S2). We also varied the length of CS to investigate the effect of CS length on triggered polymer dissociation from a hydrogel surface. DI units contained CS binding toeholds consisting of either 5 or 10 nucleotides. Corresponding CS with binding regions of 5 (CS5) or 10 nucleotides (CS-10) were incubated with corresponding polymer-functionalized samples for 10 min. In both situations, CS was able to effectively detach the cells (Figure 2C). No statistically significant difference was observed for the

examine the release of the polyvalent aptamers and the cells from the hydrogel surface. The aptamer was an Sgc8c aptamer that is specific for CCRF-CEM lymphoblastic T-cells by recognizing protein tyrosine kinase 7 (PTK7) receptors.38 Polyacrylamide hydrogels were functionalized with polyvalent aptamers via the incorporation of Acrydite-modified adapter oligonucleotides into the hydrogel matrix. The adapter sequences served as seeding points for DI to form surfaceanchored DNA polymers. DNA polymers were hybridized with FAM-labeled aptamers, forming fluorescently labeled polyvalent aptamers, as depicted in the schematic in Figure 2A. The outcomes of cell attachment on the hydrogels are also shown in Figure 2A. When the hydrogel samples were treated with buffer or scrCS, strong fluorescence was observed from the hydrogels 1385

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Figure 3. Dynamic cell attachment on the polyvalent aptamer-functionalized hydrogel in a flow chamber. (A) Schematic illustration of the parallel plate flow chamber with polyvalent aptamer-functionalized hydrogel base. (B) Representative micrographs illustrating dynamic cell attachment. Frames represent 0.5 s increments, where the red arrow represents a cell that was eventually attached and the green arrow indicates a cell that passes through the viewing area without attachment at 0.01 Pa shear stress. (C) Kinetics of cell attachment on the hydrogel at 0.01 Pa shear stress. Error bars represent standard deviation, (n = 3).

Figure 4. Effects of shear stress and seeding density on dynamic cell attachment. (A) Analysis and representative images of cell attachment in response to varied shear stress. (B) Analysis and representative images of cell attachment as a function of polyvalent seeding aptamer density. All error bars represent standard deviation (n = 3).

detachment of the cells with the variation in CS binding length from 5 to 10 nucleotides. These data demonstrate that CS can detach cells through the triggered dissociation of DNA polymers. Enzymes such as proteinases and nucleases have also been used to induce cell detachment from substrates.39−41 The concerns with the use of enzymes include the potential of

damaging the cell surface or the inefficiency of digesting chemically modified oligonucleotides.42,43 The use of CS to trigger the cell detachment does not have these concerns. 3.3. Examination of Cell Attachment in a Polyvalent Aptamer-Functionalized Parallel Plate Flow Chamber. For applications such as cell isolation from a blood sample, it is 1386

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experiments were conducted at 0.01 Pa. To study the role of polyvalent aptamer seeding density on cell attachment in dynamic conditions, parallel flow chamber hydrogel bases were prepared with polyvalent aptamers seeded at concentrations ranging from 1−100 μM. Cell suspension was perfused over these substrates for equal amounts of time, allowing for the investigation of cell attachment on the substrates. As shown in Figure 4B, as polyvalent aptamer seeding density increases, the amount of cell attachment is also increased. The increase in cell attachment reaches a plateau after the seeding density of polyvalent aptamers surpasses 50 μM (Figure 4B). To demonstrate the cell attachment specificity on the aptamer-functionalized surface, aptamer and scrambled-aptamer (Scr Aptamer) surfaces were compared. As shown by the images and quantitative analysis in Figure 5A, control surfaces

often necessary to process a large volume of cell suspension in a flow system. Thus, it is important to examine whether cell attachment can occur in dynamic flow. To this end, we utilized a polyvalent aptamer hydrogel as the base of a parallel plate flow chamber, which was connected via tubing to a syringe pump feeding solutions of interest to the chamber (Figure 3A). The chamber was sealed via vacuum, and solutions exited the device through output tubing. The hydrogel base of the flow chamber could be observed in real time using a transparent viewing area, which rests on the stage of an inverted microscope (Figure 3A). As shown in the series of images (Figure 3B) and video (Movie S1), target cells could attach to the functionalized flow chamber base. The red arrow shows a representative cell that flowed across the aptamer-functionalized surface, decelerated as it interacted with the surface, and eventually halted as it became attached (Figure 3B). It was also found that many cells did not attach to the surface. A representative cell marked by a green arrow was not attached in this viewing area and was seen crossing the region without binding (Figure 3B). The data demonstrate that the hydrogel surface with the polyvalent aptamer can induce cell attachment. The data also suggest that it is important to optimize the system to enhance the opportunities for cells to interact with the hydrogel surface for cell attachment in flow conditions. In fact, several strategies have been developed to enhance the contact between affinity substrates and target cells, including the use of nanostructures or optimized chamber geometries.37,44−46 These strategies could be applied to the current polyvalent aptamer flow system for an increased surface area-to-volume ratio in an effort to increase contact between the polyvalent aptamers and cells. We further examined the kinetics of cell attachment on the polyvalent aptamer-functionalized surface. Cell suspension was perfused over the functionalized surface at 0.01 Pa shear stress for 3.5 min. As shown in Figure 3C, cell attachment proceeded at a nearly linear rate over the time range tested. Notably, flowing cells were observed to interact with attached cells. This interaction could slow their passage over the surface and promote their eventual attachment. Since cells attached to the hydrogel surface became microscale subjects, this result suggests that a hydrogel surface may be optimized with microscale protrusions to improve the cell attachment efficiency. After cell attachment on the hydrogel with the polyvalent aptamer in dynamic conditions was illustrated, experiments were pursued to identify the attachment capabilities of the surface in response to changes in flow parameters. Shear stress was altered by varying the flow rates of cell solution to elicit a shear stress range of 0.005−0.1 Pa. This range was chosen to fall within the physiological shear stress range experienced by cells in vascular circulation as well as the interstitial flow experienced in the tumor microenvironment.47 Therefore, the shear stresses imposed upon cells at the functionalized hydrogel surface mimic the physiological conditions and could be used to analyze attached cell behavior and morphology in future work. As shown in the images and analysis in Figure 4A, cell attachment was greatest at the lowest shear stress tested, 0.005 Pa, and was significantly reduced as the shear stress approached 0.1 Pa. The increase in cell attachment to a surface with decreased shear stress was also observed in other studies using P-selectin-coated tubes1 and an aptamer-coated PDMS device.37 However, while cell attachment was greatest at 0.005 Pa, the flow rate was slow. Thus, all the following

Figure 5. Examination of the specificity of cell attachment. (A) Analysis and representative phase micrographs of the attached cells on the polyvalent aptamer-functionalized hydrogel after 3.5 min of cell attachment. Scr Aptamer: scrambled aptamer. (B) CCRF-CEM target cells were stained with CMAC dye (blue) prior to mixing with unstained control Ramos cells at a 1:1 cell ratio for a final cell density of 5 × 105 cells/mm2. Attached cells were imaged under phase (CCRF-CEM + Ramos) and fluorescence (CCRF-CEM) microscopy, with merged images indicating control Ramos cells (yellow arrow). Quantitative analysis demonstrates the amount of attached cells that were identified as CCRF-CEM or Ramos. Error bars represent standard deviation (n = 3); *P ≤ 0.05.

were capable of binding ∼12 cells/mm2, whereas aptamerfunctionalized hydrogels were capable of binding ≥400 cells/ mm2. Another experiment was performed to examine the attachment of both target and control cells. Target CCRF-CEM cells were fluorescently stained and mixed with unstained control Ramos cells prior to perfusion over the substrates. Attached cells were imaged under both phase microscopy and 1387

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μM. The cell detachment efficiency of CS increased as its concentration was increased; however, it became less significant between the 5 μM and 10 μM CS solutions (Figure 6B). The effect of CS treatment time on cell detachment was also examined. The data show that cell detachment was increased with time and more than 90% could be detached at 20 min of CS treatment (Figure 6C). Since cell viability is an important index of cell detachment, CS-detached cells were retrieved from the parallel plate flow chamber and compared with equal numbers of intact cells obtained directly from a culture flask. Cell viability was observed using a live/dead staining assay, while cell activity was analyzed via an MTS assay. As shown in Figure 7A,

fluorescence microscopy, as shown in Figure 5B. All attached cells are visible under phase microscopy, but only the fluorescent CCRF-CEM target cells are visible under fluorescence microscopy. When phase and fluorescence images were merged, the distinct cell populations were visible (Figure 5B). The result shows that the attached cells were mainly CCRF-CEM cells. Overall, the substrate efficiently bound target cells, with CCRF-CEM cells making up over 99% of the total bound cells. These results show that the hydrogel with the polyvalent aptamer is highly specific for the attachment of target cells. This specificity would be important for applications such as in vivo catch of target cells for therapy, where a large amount of diverse nontarget cells would interact with the functionalized surface. However, modifications to enhance oligonucleotide stability and optimization of the functionalized surface morphology would be necessary for cell attachment in biological fluids. 3.4. Evaluation of CS-Triggered Cell Detachment from the Flow Chamber Hydrogel Base. Cell detachment is necessary in situations where isolated cells must not only be enumerated, but also analyzed or cultured to determine specific biomarkers or potential target surface receptors for drug delivery applications.10,43,48,49 Thus, a gentle and efficient detachment mechanism is required to preserve a high degree of cell integrity and viability. This requirement is particularly important to situations where target cells are present at a very low number in the sample.50,51 Thus, after demonstrating that the polyvalent aptamer surface is capable of binding target cells in dynamic conditions, we investigated the CS-mediated detachment of the cells bound to the hydrogel base of the flow chamber and detached cell viability. The release of dynamically bound cells was compared using solutions of either CS, scrCS, or buffer (Figure 6A). CS treatment led to the detachment of more than 90% cells. We also determined the effect of CS concentration on cell detachment. The CS concentration was varied from 1 to 10

Figure 7. Viability and activity of the detached cells. (A) Representative live/dead staining images of CS-detached cells in comparison to intact cells. Green fluorescence indicates viable cells, while red fluorescence marks dead cells (indicated by red arrows). (B) Analysis of cell activity after cell detachment via MTS assay. The activity of intact cells was used as 100%. Error bars represent standard deviation (n = 3); *P ≤ 0.05; N.S. indicates no significant difference.

detached cells showed similar viabilities to controls. However, live/dead staining indicates only the viability and not necessarily the activity of cells. Previous studies utilizing viability analysis after cell release have noted that viable released cells do not always express surface receptors at the same levels as control cells.40 Therefore, we performed an MTS cell activity assay to compare control and detached cells immediately after release and 12 h after release. The results show that the activity of detached cells was less than that of intact cells (Figure 7B). The decrease of cell activity may stem from the shear stress sensed by cells during dynamic flow. It was also found that the activity was recovered by more than 10% between the 0 and 12 h time points, while this increase was not statistically significant (Figure 7B).

4. CONCLUSIONS In summary, we have studied cell attachment on the polyvalent aptamer-functionalized hydrogel surface in dynamic flow conditions and further examined cell detachment in the presence of a triggering CS. The data have demonstrated that the polyvalent aptamer-functionalized surfaces can effectively induce cell attachment in dynamic flow. While it is challenging to elicit cell attachment on the hydrogel surface at high shear stress, reducing the shear stress into the physiological range can dramatically increase the efficiency of cell attachment. Notably, after attachment to the hydrogel surface in dynamic flow, the majority of the cells can be detached from the surface by the CS in physiological conditions without the involvement of any harsh conditions. As a result, the detached cells are viable. Thus, the aptamer-based polyvalent ligand is promising for the

Figure 6. CS-mediated detachment of cells from polyvalent aptamermodified flow chamber. (A) Analysis and representative phase micrographs of cell detachment via CS, scrCS, or buffer. (B) Effects of CS concentration on cell detachment. (C) Cell detachment as a function of triggering time. All error bars represent standard deviation (n = 3); *P ≤ 0.05; N.S. indicates no significant difference. 1388

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Biomacromolecules

(26) Chen, N.; Huang, Y.; Wang, Y. Biomaterials 2014, 35, 9709− 9718. (27) Borbas, K. E.; Ferreira, C. S. M.; Perkins, A.; Bruce, J. I.; Missailidis, S. Bioconjugate Chem. 2007, 18, 1205−1212. (28) Santulli-Marotto, S.; Nair, S. K.; Rusconi, C.; Sullenger, B.; Gilboa, E. Cancer Res. 2003, 63, 7483−7489. (29) Richards, E.; Li, S.; Chen, N.; Battig, M. R.; Wang, Y. Biomacromolecules 2014, 15, 4561−4569. (30) Dirks, R. M.; Pierce, N. A. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 15275−15278. (31) Wan, Y.; Liu, Y.; Allen, P. B.; Asghar, W.; Mahmood, M. A. I.; Tan, J.; Duhon, H.; Kim, Y.; Ellington, A. D.; Iqbal, S. M. Lab Chip 2012, 12, 4693−4701. (32) Hubble, J. Adv. Biochem. Eng./Biotechnol. 2007, 106, 75−99. (33) Gurkan, U. A.; Anand, T.; Tas, H.; Elkan, D.; Akay, A.; Keles, H. O.; Demirci, U. Lab Chip 2011, 11, 3979−3989. (34) Gilham, P. T.; Khorana, H. G. J. Am. Chem. Soc. 1958, 80, 6212−6222. (35) Beaucage, S. L.; Iyer, R. P. Tetrahedron 1992, 48, 2223−2311. (36) Fire, A.; Xu, S.-Q. Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 4641− 4645. (37) Zhao, W.; Cui, C. H.; Bose, S.; Guo, D.; Shen, C.; Wong, W. P.; Halvorsen, K.; Farokhzad, O. C.; Teo, G. S. L.; Phillips, J. A.; Dorfman, D. M.; Karnik, R.; Karp, J. M. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 19626−19631. (38) Shangguan, D.; Li, Y.; Tang, Z.; Cao, Z. C.; Chen, H. W.; Mallikaratchy, P.; Sefah, K.; Yang, C. J.; Tan, W. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 11838−11843. (39) Chen, L.; Liu, X.; Su, B.; Li, J.; Jiang, L.; Han, D.; Wang, S. Adv. Mater. 2011, 23, 4376−4380. (40) Li, S.; Chen, N.; Zhang, Z.; Wang, Y. Biomaterials 2013, 34, 460−469. (41) Kwong, G. A.; Radu, C. G.; Hwang, K.; Shu, C. J.; Ma, C.; Koya, R. C.; Comin-Anduix, B.; Hadrup, S. R.; Bailey, R. C.; Witte, O. N.; Schumacher, T. N.; Ribas, A.; Heath, J. R. J. Am. Chem. Soc. 2009, 131, 9695−9703. (42) Canavan, H. E.; Cheng, X.; Graham, D. J.; Ratner, B. D.; Castner, D. G. J. Biomed. Mater. Res., Part A 2005, 75, 1−13. (43) Zheng, Q.; Iqbal, S. M.; Wan, Y. Biotechnol. Adv. 2013, 31, 1664−1675. (44) Wan, Y.; Mahmood, M. A. I.; Li, N.; Allen, P. B.; Kim, Y.-T.; Bachoo, R.; Ellington, A. D.; Iqbal, S. M. Cancer 2012, 118, 1145− 1154. (45) Wang, S.; Wan, Y.; Liu, Y. Nanoscale 2014, 6, 12482−12489. (46) Stott, S. L.; Hsu, C.; Tsukrov, D. I.; Yu, M.; Miyamoto, D. T.; Waltman, B. A.; Rothenberg, S. M.; Shah, A. M.; Smas, M. E.; Korir, G. K.; Floyd, F. P.; Gilman, A. J.; Lord, J. B.; Winokur, D.; Springer, S.; Irimia, D.; Nagrath, S.; Sequist, L. V.; Lee, R. J.; Isselbacher, K. J.; Maheswaran, S.; Haber, D. A.; Toner, M. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 18392−18397. (47) Mitchell, M. J.; King, M. R. J. Mol. Diagn. 2013, 15, 1−18. (48) Baum, C. M.; Weissman, I. L.; Tsukamoto, A. S.; Buckle, A. M.; Peault, B. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 2804−2808. (49) Singh, S. K.; Clarke, I. D.; Terasaki, M.; Bonn, V. E.; Hawkins, C.; Squire, J.; Dirks, P. B. Cancer Res. 2003, 63, 5821−5828. (50) Bichsel, C. A.; Gobaa, S.; Kobel, S.; Secondini, C.; Thalmann, G. N.; Cecchini, M. G.; Lutolf, M. P. Lab Chip 2012, 12, 2313−2316. (51) Myung, J. H.; Gajjar, K. A.; Chen, J.; Molokie, R. E.; Hong, S. Anal. Chem. 2014, 86, 6088−6094.

functionalization of materials in applications that require regulated cell attachment and detachment.

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ASSOCIATED CONTENT

S Supporting Information *

Additional cell attachment video content. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Fax: (814) 863-0490. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was in part supported by the Penn State Start-up Fund and the U.S. National Science Foundation (DMR 1322332).

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REFERENCES

(1) Han, W.; Allio, B. A.; Foster, D. G.; King, M. R. ACS Nano 2009, 4, 174−180. (2) Ito, Y. Soft Matter 2008, 4, 46−56. (3) Lutolf, M. P.; Hubbell, J. A. Nat. Biotechnol. 2005, 23, 47−55. (4) Reinherz, E. L.; Kung, P. C.; Goldstein, G.; Schlossman, S. F. Proc. Natl. Acad. Sci. U.S.A. 1979, 76, 4061−4065. (5) Nam, J.-M.; Thaxton, C. S.; Mirkin, C. A. Science 2003, 301, 1884−1887. (6) Malmqvist, M. Nature 1993, 361, 186−187. (7) El-Sayed, I. H.; Huang, X.; El-Sayed, M. a. Nano Lett. 2005, 5, 829−834. (8) Kiessling, L. L.; Gestwicki, J. E.; Strong, L. E. Curr. Opin. Chem. Biol. 2000, 4, 696−703. (9) Wrenn, S. J.; Weisinger, R. M.; Halpin, D. R.; Harbury, P. B. J. Am. Chem. Soc. 2007, 305, 13137−13143. (10) Ma, H.; Liu, J.; Ali, M. M.; Mahmood, M. A. I.; Labanieh, L.; Lu, M.; Iqbal, S. M.; Zhang, Q.; Zhao, W.; Wan, Y. Chem. Soc. Rev. 2015, 44, 1240−1256. (11) Soontornworajit, B.; Wang, Y. Anal. Bioanal. Chem. 2011, 399, 1591−1599. (12) Bishop, G. R.; Ren, J.; Polander, B. C.; Jeanfreau, B. D.; Trent, J. O.; Chaires, J. B. Biophys. Chem. 2007, 126, 165−175. (13) Tuerk, C.; Gold, L. Science 1990, 249, 505−510. (14) Ellington, A. D.; Szostak, J. W. Nature 1990, 346, 818−822. (15) Jayasena, S. D. Clin. Chem. 1999, 45, 1628−1650. (16) Mammen, M.; Choi, S.-K.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 2754−2794. (17) Tan, L.; Gee Neoh, K.; Kang, E.-T.; Choe, W.-S.; Su, X. J. Pharm. Sci. 2012, 101, 1672−1677. (18) Mallikaratchy, P. R.; Ruggiero, A.; Gardner, J. R.; Kuryavyi, V.; Maguire, W. F.; Heaney, M. L.; McDevitt, M. R.; Patel, D. J.; Scheinberg, D. A. Nucleic Acids Res. 2011, 39, 2458−2469. (19) Davis, K. A.; Abrams, B.; Lin, Y.; Jayasena, S. D. Nucleic Acids Res. 1996, 24, 702−706. (20) Ringquist, S.; Parma, D. Cytometry 1998, 33, 394−405. (21) Bell, N. M.; Micklefield, J. ChemBioChem 2009, 10, 2691−2703. (22) Roy, S.; Caruthers, M. Molecules 2013, 18, 14268−14284. (23) Baldi, P.; Hatfield, G. W. In DNA Microarrays and Gene Expression: From Experiments to Data Analysis and Modeling; Halliday, K., Tranah, D., Eds.; Cambridge University Press: New York, 2002; pp 7−16. (24) Boyacioglu, O.; Stuart, C. H.; Kulik, G.; Gmeiner, W. H. Mol. Ther. Nucleic Acids 2013, 2, 1−8. (25) Ii, J. O. M.; Kolonias, D.; Pastor, F.; Mittler, R. S.; Chen, L.; Giangrande, P. H.; Sullenger, B.; Gilboa, E. J. Clin. Invest. 2008, 118, 376−386. 1389

DOI: 10.1021/acs.biomac.5b00165 Biomacromolecules 2015, 16, 1382−1389