Assessing Aptamer-Analyte Binding Kinetics by Microfluidic Fluorescence Microscopy

Abstract

Kinetic measurement plays a crucial role in understanding aptamer binding mechanisms and identifying appropriate aptamers for clinical and research applications. Current techniques, while well established, generally require large sample volumes, bulky and expensive instruments operated by trained personnel, and are hence not readily accessible to resource-limited research laboratories. This paper presents a fluorescence microscopy-based microfluidic assay for measuring aptamer-analyte binding kinetics in a simple and cost-effective manner. Kinetic measurements are achieved by monitoring time-course fluorescence of fluorescently labeled aptamers as they bind to the targets trapped in a microfluidic chip. Fluorescence measurements are performed on a standard fluorescence microscope and are accessible to laboratories with only modest resources. Moreover, microfluidic technology allows efficient and cost-effective immobilization of small amounts of target molecules or live cells as well as flow-based manipulation of aptamers for the measurements. Kinetic measurements of aptamer binding to immunoglobulin E protein and CCRF-CEM cells have yielded results consistent with those obtained from established methods, demonstrating the potential utility of our method for exploring aptamer-target interactions and identifying aptamers that best suit specific given biomedical applications.

1. Introduction

Aptamers are short, single-stranded oligonucleotides (DNA or RNA) that bind to a diverse set of biological targets such as small molecules, heavy metal ions, proteins, virus, and cells [1, 2]. Isolated in vitro from randomized oligonucleotide libraries through an evolutionary process known as SELEX, aptamers have seen applications in a broad range of fields including biotechnology, medicine, and diagnostics [3]. Systematic characterization of target binding properties of aptamers, including affinity, specificity, and kinetics, is essential for such applications [4, 5]. The affinity between an aptamer and its intended target can be assessed by equilibrium binding measurements via methods such as enzyme-linked oligonucleotide assay (ELONA) [6], electrophoretic gel shift assay, and flow cytometry [7, 8]. However, kinetic information is often desired or even required to gain insights into aptamer-target binding mechanisms and choose the most suitable aptamers for various applications. For instance, aptamers with fast association and dissociation kinetics may be desirable for sensing applications, while histological staining may require aptamer-target interactions with very long half-lives. Similarly, therapeutic aptamers with long half-lives may be administered in lower dosages, reducing therapeutic costs and potential side effects.

Kinetic properties of aptamer-target binding are currently measured with techniques such as surface plasmon resonance (SPR) [9, 10] and optical interferometry [11, 12]. In these techniques, time-dependent changes in refractive indices and shifts in interference patterns are measured to quantify the association and dissociation rates of the aptamer-target complex formation. While well established, they generally require large volumes or high concentrations of samples, involve bulky and expensive instruments operated by trained personnel, and are hence not easily accessible to research laboratories with limited resources or expertise. Furthermore, the common approach for studying aptamer interactions with suspension cells is flow cytometry assay, but this method only provides end-point data and lacks kinetic information [7, 8].

There is therefore an urgent need for simple and low-cost methodologies for studying aptamer-target binding kinetics. Aiming to address this need, we present a microfluidic fluorescence assay for kinetic measurements, utilizing time-resolved imaging of fluorescence from the binding between aptamers and biological targets, such as proteins and live cells, in a microfluidic chip. The synergistic combination of microfluidic technology and fluorescence microscopy, whose utility has been established in kinetic measurements of enzymatic reactions [13, 14], antibody-antigen interactions [15, 16], and biotin-streptavidin interactions [16, 17], represents a new opportunity with several unique advantages for measuring kinetics of aptamer-target analyte binding.

First, microfluidic technology allows efficient and cost-effective immobilization of small amounts of biological targets such as proteins and live cells within a microchamber through using microbeads and trapping microstructures. Microfluidic allows the easy operation of a minimal number of microbeads (tested in 20 beads coated with ~0.1 picomole protein molecules) or live cells (~1000 cells) for interaction with aptamers within fluid stream, resulting in a significant reduction in sample consumption by over two orders of magnitude compared to SPR techniques (Table S1). These trapped beads or cells can also be easily removed by reverse flow to reuse the microfluidic device, resulting in reduced chip fabrication costs. Second, the use of bead-based immobilization for protein targets offers compatibility with post-SELEX aptamer characterization. This approach streamlines the workflow, especially considering that many aptamers are isolated through bead-based SELEX [18–20]. The direct utilization of target molecule-coated beads obtained from the SELEX process eliminates the need for additional bead preparation, saving valuable time, resources, and effort. In addition, microfluidics offers a novel, simple and inexpensive platform for live cell trapping and manipulation that remains challenging for SPR, thereby enabling highly efficient interactions between aptamers and live cells and facilitating precise kinetic measurements. These trapped beads or cells can also be easily removed by reverse flow to reuse the microfluidic device, resulting in reduced chip fabrication costs. Third, the use of microbeads or single-cell trapping microstructures provides spatially separated microenvironments allowing multiplexed measurements of proteins or cells with multispectral imaging. The assay thus allows for simultaneous kinetic measurements of a large number of aptamer-target interactions with improved throughput and drastically reduced measurement time. Finally, while a standard fluorescence microscope is used for demonstration of principle in this work, the current non-optimized instrumentation setup still allows low-cost aptamer kinetics measurements in non-specialized laboratories where standard fluorescence microscopes are already available. This standard fluorescence microscope could ultimately be replaced with a portable digital fluorescence microscope or a smartphone-based fluorescence microscope [21, 22] to produce an optimized, truly low-cost measurement system.

The microfluidic assay has been demonstrated by measuring the kinetics of aptamers binding to human immunoglobulin E (IgE) as a representative target protein, and to a human acute lymphoblastic leukemia cell (CCRF-CEM) as a target cell. The constants of association rate and dissociation rate of aptamer-IgE binding are determined to be 3.7 × 10⁵ M⁻¹ s⁻¹ and 2.2 × 10⁻³ s⁻¹, respectively, which are consistent with the previously measured values using SPR. For aptamer-cell binding, the constants of association rate and dissociation rate are determined to be 1.6 × 10⁵ M⁻¹ s⁻¹ and 6.8 × 10⁻⁴ s⁻¹, respectively. The equilibrium constant extracted from our method (4.3 nM) is consistent with the value obtained from the flow cytometry assays (~1 nM). Furthermore, we have demonstrated that the assay can be used not only for measuring aptamer-target binding kinetics at different ionic strengths and temperatures, but also for measuring kinetics of aptamers against IgE and His tag simultaneously.

2. Experimental Section

2.1. Principal and Device Fabrication

To measure aptamer-protein or aptamer-cell binding kinetics, we monitor the fluorescence of fluorescently labeled aptamers over time as they interact with live cells or proteins immobilized on beads inside a microfluidic chip. The microfluidic chip contains a hexagonal chamber measuring 7 mm in length and 3 mm in width. This chamber has two inlets, an aptamer inlet for association measurement and a wash buffer inlet for dissociation measurement. As fluorescently labelled aptamers flow through the cells or bead-immobilized proteins, they interact with their targets and the amount of bound aptamer increases until it reaches an equilibrium state. To evaluate the dissociation rate, buffer is injected to wash away dissociated aptamers from the beads and the fluorescence intensity on the beads is measured over time (Figure 1a). For aptamer-protein binding kinetic measurement, we use a weir-like structure to trap the beads in the microfluidic chip. The weir structure made the height (20 μm) of outlet channel smaller than the diameter of microbeads (45~165 μm), which allows the beads can to be uniformly packed in the microscale chamber for efficient interaction between aptamer and targets and accurate determination of fluorescence intensity. To decrease the background fluorescence of free aptamer in solution, we use a thin chamber height (180 μm), enabling the fluorescence intensity of bound aptamer to be easily distinguished from that of free aptamers in solution (Figures 1b and 1c, S2). For measuring aptamer-cell binding kinetics, a cell-trapping section are incorporated inside the microchamber. Cells introduced through the inlet channel and enter the trapping section, which consists of an array of microscale traps. The traps, each comprised of a pair of microposts resembling a cup with a slit at the downstream end. Cells are directed into the individual traps by design of the trap geometry and placement. The slit is sized to be smaller than cells; hence, as the medium flows through the slit, cells become trapped (Figures 1d-1e). The trap structure can accommodate cells within the 10–20 μm diameter range. Given that the majority of cultured suspension cancer cell lines fall within the 15–20 μm range, our chip exhibits versatility, making it suitable for trapping and conducting kinetic analyses on other suspension cancer cells. The device is fabricated from polydimethylsiloxane (PDMS) using standard multilayer soft lithography techniques as described previously by our group [23–25]. Briefly, SU8 and silicon wafers are used to create master molds bearing the device design. PDMS is then poured onto the master molds, cured, peeled, cut, and bonded onto the glass slide (Figure S1).

Figure 1.

Microfluidic-based measurement of aptamer-protein and aptamer-cell binding kinetics. (a) Schematic illustration of the microfluidic device with two inlets for aptamer and buffer injection. Schematic illustration of (b) protein-coated beads trapped in a microchamber using a weir-like structure, and (c) kinetic reaction of aptamer-protein binding. Schematic illustration of (d) the cell traps used for cell trapping, and (e) interaction with fluorescently labeled aptamers.

2.2. Immobilization of Proteins on Microbeads

Human IgE protein was immobilized covalently on NHS-activated Sepharose beads. An aliquot of 10 μL of NHS-activated Sepharose^™ 4 Fast Flow slurry (BioFront Technologies) was washed three times with 100 μL of phosphate-buffered saline (PBS) buffer (ThermoFisher Pierce). Then, IgE protein was incubated with the NHS-activated beads on a homemade rotator at room temperature (RT) for 30 min. Following washing three times with 100 μL of PBS, the unoccupied sites on agarose beads were passivated with 100 mM of ethanolamine with slow rotation for 30 min at RT. The unbound ethanolamine was washed 3 times with PBS. Finally, the IgE-coated beads were suspended in 50 μL of 1× PBS buffer and stored at 4 °C for use. His-tagged human BCL2-associated X (BAX) protein (OriGene) was immobilized on the NHS-activated beads using the same protocol as IgE beads above.

2.3. Cell Culture

CCRF-CEM cell lines (ATCC) were cultured in RPMI 1640 medium (ATCC) with 10% fetal calf serum (ATCC). Prior to being loaded into microfluidic devices, cells were washed three times by centrifugation at 125 × g for 5 min and then resuspended in PBS buffer.

2.4. Kinetic Measurements

At the start of the experiment, microchamber was injected with a PBS/BSA/Tween 20 solution from the inlets to precoat channel walls and reduce nonspecific binding. The fluorophore-conjugated aptamers (Integrated DNA Technologies) shown in Table S2 were pretreated at 95 °C for 10 min and immediately cooled at −20 °C for 4 min, followed by incubation at room temperature for 15 min to fold the ssDNA molecules. Protein-coated beads (5 μL) or cells (100 μL, 1×10⁴ cell/mL) were injected into the microchamber and washed with the same buffer as the one aptamer dissolved. Then, the prepared aptamer was introduced by a syringe at a fixed velocity to interact with the protein-coated beads or cells. The florescence images on beads or cells were acquired by an inverted fluorescence microscopy (Carl Zeiss Microscopy, LLC) equipped with a X-Cite 120 LED Boost System, FL Filter Set 43 HE Cy3, and FL Filter Set 38 HE GFP at time intervals. The average fluorescence intensity was quantified with ImageJ software by selecting line profiles through the beads and recording the intensity at the bead surface.

The measured fluorescence intensities were assumed to be proportional to the concentration of aptamer-protein/cell complex and were fit to the standard first-order interactions [26, 27]. Briefly, the association rate constant $\left(k_{\text{on}}\right)$ is given by Equation (1),

$$F_{\text{t}}=F_{\text{eq}}\left(1-e^{-\left(k_{\text{on}}c+k_{\text{off}}\right)t}\right)$$ (1)

and the dissociation rate constant $\left(k_{\text{off}}\right)$ is given by Equation (2),

$$F_{\text{t}}=\left(F_{\text{eq}}-F_{\infty }\right)e^{-k_{\text{off}}\left(t-t_{\text{d}}\right)}+F_{\infty }$$ (2)

where $F_{\text{t}}$ represents the measured bead fluorescence intensity at time $t$, $F_{\text{eq}}$ is the background bead fluorescence intensity at the equilibrium state, $F_{\infty }$ is the fluorescence intensity at infinite time, $c$ is the solution aptamer concentration, and $t_{\text{d}}$ is the time when the dissociation process was initiated. The $k_{\text{off}}$ was first obtained by fitting the data of dissociation profile $\left(t\ge t_{\text{d}}\right)$ with Equation (2), and $k_{\text{on}}$ was obtained then by fitting association profile $\left(0\le t\le t_{\text{d}}\right)$ with Equation (2). The equilibrium dissociation constant $\left(K_{\text{D}}\right)$ can be obtained from $k_{\text{on}}$ and $k_{\text{off}}$ by Equation (3).

$$K_{\text{D}}=k_{\text{off}}/k_{\text{on}}$$ (3)

In addition, the $K_{\text{D}}$ can be calculated by Equation (4), using the corresponding equilibrium fluorescence intensities at varying concentrations of aptamer [28],

$$F_{\text{eq}}=\frac{F_{\max }c}{K_{\text{D}}+c}$$ (4)

where $F_{\max }$ represents the maximum fluorescence intensity as all target binding sites on beads are occupied by aptamer. The $K_{\text{D}}$ calculated from the equilibrium reaction should be comparable with the one measured from the kinetic experiments.

3. Results and Discussion

3.1. Measurement of Aptamer-IgE Binding Kinetics

Binding kinetics of a well-characterized IgE aptamer, D17.4, was first measured to validate the microfluidic fluorescence assay. FAM-labelled aptamer solution was introduced continuously at a flow rate of 40 μL/min to interact with IgE protein immobilized on beads and trapped in the microchamber. Then, PBSM buffer was introduced to dissociate the aptamers. Fluorescence images were acquired and fluorescence intensities were obtained at defined time intervals to monitor kinetic process of the binding event (Figures 2a and 2b). By fitting with Equations (1) and (2), we extracted the constants of association rate $\left(k_{\text{on}}\right)$ and dissociation rate $\left(k_{\text{off}}\right)$ as (3.7 ± 0.9) × 10⁵ M⁻¹ s⁻¹ and (2.2 ± 0.6) × 10⁻³ s⁻¹, respectively. These results are in a good agreement with the values of 4.3 × 10⁵ M⁻¹ s⁻¹ and 2.0 × 10⁻³ s⁻¹ previously measured by SPR [29]. The $K_{\text{D}}$ was determined as 5.9 ± 2.2 nM based on Equation (3), which is consistent with values of 4.7 nM measured by SPR [29], 6.1~15 nM by fluorescence anisotropy [30, 31], 12 nM by graphene field effect transistors [26]. In addition, we evaluated the D17.4-IgE equilibrium binding affinity by measuring the equilibrium bead fluorescence at different concentrations of fluorescently labeled aptamer (Figure 2c). By fitting the equilibrium bead fluorescence measurements with Equation (4), the $K_{\text{D}}$ was determined to be 9.3 ± 3.4 nM, which is consistent with the value from kinetic measurement, confirming the validity of the microfluidic fluorescence microscopy assay for measuring the aptamer-protein binding kinetics. It is essential to note that the addition of a fluorophore label to aptamers could theoretically influence the binding kinetics between the aptamer and the analyte. However, the fluorophore labeling is typically performed at non-essential regions of the aptamer, often at the 5' end or primer regions, which are not typically involved in the core binding interaction. As a result, the impact of fluorophore labeling on the aptamer's binding affinity and kinetics is usually minimal. This is supported by the consistent results obtained using the SPR, where the aptamer was not modified with a fluorophore. In addition, the coating of proteins on beads may alter the spatial structure of the protein and potentially affect the kinetics between the aptamer and the protein. This is a known limitation of many immobilization-based techniques for kinetic measurements. In such cases, advanced techniques that minimize or account for structural changes during immobilization are essential. Future developments in this area may lead to improved methods that mitigate these limitations.

Figure 2.

Measurements of aptamer-protein binding kinetics. (a) Fluorescence image snapshots taken during association and dissociation processes of FAM-labelled aptamer D17.4 (25 nM) interacting with IgE-coated beads. Scale bar: 200 μm. (b) Fluorescence measurements of association and dissociation kinetics of (b) bead-immobilized IgE protein interacting with D17.4 with varying concentrations. (c) Corresponding plots of aptamer-IgE binding at steady equilibrium states from the end of the association phases against aptamer concentration were used to calculate the equilibrium dissociation constant. Error bars represent the standard deviation from three repetitive measurements.

To minimize nonspecific binding, a PBS/BSA/Tween 20 solution was used to precoat PDMS chamber walls, and ethanolamine was utilized to block the beads. No adsorption of aptamers to the PDMS-based microfluidic walls or the surfaces of agarose microbeads was observed (Figure S2). While our results are promising, we acknowledge the potential challenges associated with strongly adherent molecules or intensely labeled samples, which might contribute to background fluorescence. To address this concern, we recommend the incorporation of robust surface blocking reagents like polyethylene glycol (PEG) and casein to effectively treat surfaces and prevent undesired adsorption. Additionally, for kinetic measurements, optimizing labeling concentrations can mitigate background fluorescence in both the solution and on the walls or beads. Adjusting the microchamber thickness is another strategy to further minimize background fluorescence in the solution. These measures can be selectively employed to address specific cases where background interference becomes a critical factor.

To further ensure accurate determination of aptamer-target binding kinetics, the stability of the fluorophores was characterized to assess the impact of photobleaching on the fluorescence measurements. After the aptamer binding step on the beads, we measured the fluorescence intensity on the beads without continuous washing. The fluorescence intensity only exhibited a minimal decrease of approximately 5% after subjecting the fluorophores to 10 successive measurements, each with a 150 ms exposure time (Figure S3). It is important to note that the decrease in fluorescence intensity observed includes contributions from both photobleaching of the fluorophores and the dissociation of the aptamer from the beads, as we performed a one-time quick wash after the aptamer binding step. Considering this, we can conclude that the effects of fluorophore photobleaching on the determination of aptamer-target binding kinetics are negligible, ensuring the reliability of our measurements. Furthermore, to mitigate potential issues related to mass transport limitation, we conducted a comprehensive analysis of the effects of different flow rates on aptamer-target binding kinetics. This analysis aimed to ensure that our measurements accurately reflected the binding behavior. Association and dissociation curves were obtained at various flow rates ranging from 5 to 60 μL/min (Figure S4). No significant changes in aptamer association and dissociation behavior were observed across the flow rate range of 20 to 60 μL/min. This finding suggests that the impact of diffusion limitation on the measured kinetics is negligible. Thus, our measurements accurately represent the true aptamer-target binding kinetics, as they are not significantly affected by variations in flow rates within this range. To provide further confirmation regarding the absence of diffusion limitation in our measurements of aptamer-target binding kinetics, we conducted an investigation on the effect of the number of target-coated beads. Specifically, we examined the association and dissociation behavior at a flow rate of 40 μL/min, while varying the number of beads. The association and dissociation curves obtained for different numbers of beads, ranging from 20 to ~500, were nearly identical (Figure S5). This finding suggests that the varying number of target-coated beads did not induce diffusion limitation in our experimental system. Therefore, the measured aptamer-target binding kinetics remain unaffected by the number of beads, further affirming the reliability of our aptamer-target binding kinetics measurements.

3.2. Effects of Environmental Conditions on Aptamer-Target Binding Kinetics

Aptamer-analyte binding kinetics are highly affected by environmental conditions. In order to demonstrate the effectiveness of the microfluidic fluorescence assay in assessing the impact of these conditions on aptamer binding kinetics, we specifically investigated the effects of two important environmental factors, Mg²⁺ concentration and temperature.

Mg²⁺ plays a crucial role in facilitating the formation and maintenance of specific aptamer conformations required for target binding. To meet the specific requirements of various application scenarios, it is essential to optimize the Mg²⁺concentration to modulate aptamer binding kinetics [32]. Here, we examined the effect of increasing Mg²⁺ concentration from 1 to 20 mM on aptamer-IgE binding kinetics. We found that this increase led to an increase in the dissociation rate constant $k_{\text{off}}$ from (2.2 ± 0.6) × 10⁻³ s⁻¹ to (17.6 ± 4.6) × 10⁻³ s⁻¹, and an increase in the association rate $k_{\text{on}}$ from (3.7 ± 0.9) × 10⁵ M⁻¹ s⁻¹ to (6.2 ± 1.6) × 10⁵ M⁻¹ s⁻¹, respectively (Figure 3a), consistent with previously reported results [32]. In addition, the fluorescence intensity at equilibrium state decreased with increasing Mg²⁺ concentration from 1 mM to 20 mM. These results indicated that the aptamer-IgE binding affinity decreased with increasing Mg²⁺ concentration. This is because Mg²⁺ can promote the folding and structuring of nucleic acids via interacting with negatively charged phosphate backbones, resulting in a conformation less capable of recognizing the target [32]. Since the aptamer was dissociated completely from the IgE beads within 3 minutes at 20 mM Mg²⁺ concentration, the PBSM buffer (PBS + 20 mM Mg²⁺) can be used as a regeneration reagent during our measurements of binding kinetics to regenerate the IgE beads without affecting the IgE protein (Figure 3b). This allows for reuse of the IgE beads in the chamber, saving cost and improving the accuracy of fluorescence intensity measurements when repeating the experiment.

Figure 3.

Effects of Mg²⁺ concentration and temperature on aptamer–IgE binding kinetics. (a) Association and dissociation profiles of the aptamer–IgE interaction at different Mg²⁺ concentrations. (b) High Mg²⁺ concentration (20 mM) buffer can be used as a regeneration reagent to regenerate IgE-coated beads. (c) Schematic illustration of the on-chip close-loop temperature control system, consisting of resistive heaters and temperature sensors. (d) Association and dissociation profiles of the aptamer–IgE interaction at different temperatures. Error bars represent the standard deviation from three repetitive measurements.

Temperature can influence the secondary and tertiary structure of aptamers. Changes in temperature may cause conformational changes that can affect the aptamer's ability to bind to its target. To prove that the assay is applicable to investigate the influence of temperature on aptamer binding kinetics, two temperatures, 24 °C (room temperature) and 37 °C (human body temperature), were studied because they are commonly used for aptamer isolation and applications. An on-chip closed-loop temperature control system, which was described previously by our group [33, 34], was integrated with the microfluidic fluorescence assay o control the temperature (Figure 3c and S6). The dissociation rate $k_{\text{off}}$ increased from (2.2 ± 0.6) × 10⁻³ s⁻¹ at 24 °C to (15.5 ± 4.9) × 10⁻³ s⁻¹ at 37 °C, which was in a good agreement with the $k_{\text{off}}$ value of 20.3 × 10⁻³ s⁻¹ previously reported [26]. The association rate $k_{\text{on}}$ increased from (3.7 ± 0.9) × 10⁵ M⁻¹ s⁻¹ at 24 °C to (6.3 ± 1.8) × 10⁵ M⁻¹ s⁻¹ at 37 °C (Figure 3d). Furthermore, the binding affinity of aptamer to IgE decreased, as evidenced by the increase in the $K_{\text{D}}$ value from 5.9 nM to 24.6 nM. This can be attributed to the progressive destabilization of the aptamer structure resulting from the elevated temperature of 37 °C [31], despite the fact that the aptamer D17.4 was isolated using SELEX performed at 37 °C [35].

3.3. Toward Multiplexed Measurement of Aptamer-Target Binding Kinetics

A high-throughput measurement of aptamer kinetics is particularly valuable when characterizing a large number of aptamer-analyte binding interactions. The microfluidic fluorescence assay offers the potential for high-throughput measurement of kinetics by enabling simultaneous measurement of the binding kinetics of multiple aptamer-protein interactions through optical and spatial multiplexing. To illustrate this capability, each target protein was immobilized on a distinct population of beads, which were sequentially mixed and injected into the microchamber to interact with a mixture of aptamers labelled with a spectrally distinct fluorophore. The beads were imaged using different fluorescence filter sets designed to coincide with each fluorescent aptamer, enabling the simultaneous measurement of the binding kinetics of two different aptamers (D17.4 and HisA1-T63) to two proteins (IgE and His-tagged BAX), respectively. HisA1-T63 is a DNA aptamer against His tag [24]. The binding of FAM-conjugated aptamer D17.4 to IgE beads and Cy3-conjugated aptamer HisA1-T63 to His-BAX beads were spectrally distinguished (Figure 4a). The $k_{\text{on}}$ and $k_{\text{off}}$ of aptamer HisA1-T63 were determined to be (1.5 ± 0.4) × 10⁵ M⁻¹ s⁻¹ and (3.2 ± 0.6) × 10⁻³ s⁻¹, respectively (Figure 4b). The $K_{\text{D}}$ of aptamer HisA1-T63 was calculated to be 21.3 ± 7.0 nM, which is in a good agreement with the 26.1 nM reported previously [24]. In addition, the $k_{\text{on}}$ and $k_{\text{off}}$ of aptamer D17.4 were determined to be (3.9 ± 0.5) × 10⁵ M⁻¹ s⁻¹ and (2.5 ± 0.3) × 10⁻³ s⁻¹, respectively, consistent with the values measured above in a single channel. These results demonstrate that multiplexed bead measurements can simultaneously analyze the binding kinetics of a panel of aptamers to multiple different target molecules, improving the efficiency of kinetics measurements when characterizing a large number of aptamers.

Figure 4.

Simultaneous measurement of binding kinetics from multiple aptamer-target interactions using optical and spatial multiplexing. (a) Fluoresce images taken with distinct fluorescence filter cubes to identify the FAM-conjugated D17.4 and the Cy3-conjugated HisA1-T63 bound to their respective target proteins. (b) Measured association and dissociation kinetics of two distinct aptamers interacting with two different target molecules. Error bars represent the standard deviation from three repetitive measurements.

3.4. Measurement of Aptamer-Antibody Binding Kinetics

To demonstrate the versatility of the microfluidic fluorescence assay, we measured binding kinetics for an aptamer (named TX7-Apt3), isolated by our group recently [36], which is specifically binding to the idiotype region of a human monoclonal IgG antibody, TNX7, with a high $K_{\text{D}}$ value. To ensure that the antibody remained immobilized on the beads during the experiment, protein A/G PLUS agarose beads was used for immobilizing the antibody TNX7 covalently. The kinetic measurements were performed by introducing FAM-labelled aptamer with varying concentrations from 50 nM to 800 nM (Figure 5a). The values of $k_{\text{on}}$ and $k_{\text{off}}$ were determined to be (1.7 ± 0.2) × 10⁴ M⁻¹ s⁻¹ and (7.5 ± 0.7) × 10⁻³ s⁻¹, respectively. The $K_{\text{D}}$ was calculated to be of 441.2 ± 66.3 nM based on the Equation (3), which is consistent with the $K_{\text{D}}$ of 378.8 ± 43.3 nM obtained by measuring equilibrium bead fluorescence intensity (Figure 5b). These results confirm that the microfluidic bead-based assay can be used for measuring the binding kinetics of aptamers with high $K_{\text{D}}$ values and that the assay is compatible with other types of beads, in addition to demonstrating its applicability to different types of aptamers. That is, the microfluidic bead-based fluorescence assay can be used for kinetic measurement of aptamers with a subnanomolar range of $K_{\text{D}}$ values, spanning from approximately 1 nM to 500 nM. This range covers the majority of $K_{\text{D}}$ values that aptamers typically have, indicating that the assay is applicable to a broad range of aptamer.

Figure 5.

(a) Fluorescence measurements of association and dissociation kinetics for bead-immobilized antibody TNX7 interacting with its aptamer with varying concentrations. (b) Corresponding plots of aptamer-TNX7 binding at steady equilibrium states from the end of the association phases against aptamer concentration were used to calculate the equilibrium dissociation constant. Error bars represent the standard deviation from three repetitive measurements.

3.5. Kinetics of Aptamer-Cell Binding

While a large number of aptamers against live cells have been discovered by cell SELEX, the affinity characterization has been limited to the end-point flow cytometry assays that cannot provide kinetic information [37–39]. Here, we measured the aptamer-cell binding kinetics using the microfluidic fluorescence assay. An well-known aptamer, Sgc8, binding to cell surface of human acute lymphoblastic leukemia cell line, CCRF-CEM, was chosen as the model aptamer [40]. By incorporating cup-shape microstructures [25, 41], our microfluidic device enabled cell trapping and fluorescence measurements on live cells. The efficiency of cell trapping for CCRF-CEM cells are 89.4 ± 5.4%, which is good enough for kinetic analysis since the number of cells does not affect the kinetic analysis. Using FAM-conjugated Sgc8, we monitored its flow through the cells trapped in the chamber and determined the $k_{\text{on}}$ and $k_{\text{off}}$ values to be (1.6 ± 0.3) × 10⁵ M⁻¹ s⁻¹ and (6.8 ± 2.3) × 10⁻⁴ s⁻¹, respectively. The $K_{\text{D}}$ value was calculated to be 4.3 ± 1.6 nM (Figure 6). As our method represented the first kinetic measurement for the aptamer Sgc8 binding to CCRF-CEM cells, we could not make a direct comparison with other techniques. However, the equilibrium constant extracted from our method (4.3 nM) was consistent with the value obtained from the flow cytometry assays (~1 nM) [40]. It is worth noting that the aptamer may be internalized into the cells without external assistance [42], which may lead to an underestimation of dissociation rate. To minimize the influence of internalization, we conducted the kinetic measurements within 20 minutes. While techniques for adherent cells have been well-established, studying the kinetics of binding with suspension cells presents a significant challenge. Our method fills this critical gap by offering a novel and effective approach for measuring aptamer-cell binding kinetics, specifically tailored to suspension cells. Overall, our microfluidic fluorescence assay provides a powerful tool for the kinetic characterization of aptamer-cell binding, which can aid in the development and optimization of aptamer-based therapeutics.

Figure 6.

Measurement of binding kinetics of FAM-labelled aptamer Scg8 to live CCRF-CEM cells. Error bars represent the standard deviation from three repetitive measurements.

4. Conclusion

We have presented a simple and cost-effective microfluidic-based fluorescence assay for monitoring aptamer binding kinetics to both proteins and live cells. The biding kinetics was characterized by measuring the fluorescence intensity, in a time course manner, on protein-immobilized beads or live cells that are trapped in a microfluidic chamber when the fluorescently labelled aptamers flow through. Our assay was successfully demonstrated by measuring the binding kinetics of protein IgE to aptamer D17.4. Additionally, we were able to measure the aptamer-live cell binding kinetics by incorporating cell trap features in the microchamber to trap live cells. The binding kinetics of aptamer Scg8 to CCRF-CEM cells were determined to be 1.6 × 10⁵ M⁻¹ s⁻¹ and 6.8 × 10⁻⁴ s⁻¹ for $k_{\text{on}}$ and $k_{\text{off}}$, respectively. Furthermore, we have demonstrated that the assay can be used not only for measuring aptamer-target binding kinetics at different Mg²⁺ ionic strengths and temperatures, but also for measuring kinetics of multiple aptamers simultaneously. The microfluidic fluorescence assay is simple and cost-effective since it can be easily combined with the routine fluorescence microscopy that is available in most laboratories. Thus, this assay can be potentially used as a routine and alternative tool for characterizing aptamer binding kinetics, facilitating the selection of appropriate aptamers in research and clinical applications based on their kinetic properties.

Highlights.

• Fluorescence microscopy-based microfluidic assay for measuring aptamer-analyte binding kinetics.

• Kinetics are measured for aptamer D17.4 binding to IgE and for aptamer Sgc8 binding to CCRF-CEM cell.

• Offers a cost-effective approach for studying aptamer interactions.

Biographies

Kechun Wen is a PhD candidate in Mechanical Engineering at Columbia University. He received his BS and MS degrees in Biotechnology and Material Engineering from University of Electronic Science and Technology of China, Chengdu, China, in 2016 and 2018, respectively. He received his MPhil degree in Mechanical Engineering at Columbia University, New York, USA, in 2020. His research interests are microfluidic-based isolation of aptamers for biomedical applications.

Xin Meng is a PhD candidate in Mechanical Engineering at Columbia University. He received his BS degree in Chemistry from Zhejiang University, Hangzhou, China, in 2012. He received his MS degree in Chemistry from Indiana University Bloomington, USA, in 2016. He received his MPhil degree in Mechanical Engineering at Columbia University, New York, USA, in 2019. His research interests are on microfluidic-based isolation of aptamers for biomedical applications.

Chengxi Wang received his BS degree in Mechanical Engineering at Columbia University. His research interests are in microfluidic aptamer isolation for biomedical applications.

Jingyang Zhao received his MS degree in Mechanical Engineering from Columbia University. His research interests are in microfluidic aptamer isolation for biomedical applications.

Samantha Botros received her BS degree in Biomedical Engineering from Los Andes University, Colombia. Currently, she is pursuing her MS degree in Biomedical Engineering from Columbia University, New York. Her research interests are in microfluidic aptamer isolation for biomedical applications and biosensors for DNA detection.

Qiao Lin received his PhD in Mechanical Engineering from the California Institute of Technology in 1998 with thesis research in robotics. He conducted postdoctoral research in microelectromechanical systems (MEMS) at the Caltech Micromachining Laboratory from 1998 to 2000, and was an assistant professor of Mechanical Engineering at Carnegie Mellon University from 2000 to 2005. He has been an associate professor of Mechanical Engineering at Columbia University since 2005. His research interests are in designing and creating integrated micro/nanosystems, in particular MEMS and microfluidic systems, for biomedical applications.