State of the Art prior to DYNAMO (2022)
Aptamer sensing of proteins using optical microscopy and nanopores for single-molecule protein analysis
Aptamers are short strands of DNA or RNA that can bind specifically to a target molecule, such as a protein, small molecule, or even a whole cell[cite: 9, 10]. The most widely used method to discover aptamers is SELEX, which stands for Systematic Evolution of Ligands by EXponential enrichment[cite: 9, 10]. SELEX is an iterative selection process that mimics natural evolution in the laboratory to identify sequences with high binding affinity and specificity[cite: 9, 10].
The SELEX process begins with a very large library of random nucleic acid sequences often containing up to 10¹³–10¹⁵ unique molecules[cite: 9, 10]. This library is exposed to a target of interest (for example, a protein)[cite: 9, 10]. A small fraction of sequences in the library will bind to the target, while the majority will not[cite: 9, 10]. The bound sequences are then separated from the unbound ones, typically through washing steps[cite: 9, 10]. The selected sequences are amplified using PCR (for DNA) or reverse transcription followed by PCR (for RNA), generating a new enriched pool[cite: 9, 10]. This cycle of binding, separation, and amplification is repeated multiple times (usually 8–15 rounds), gradually enriching the pool for high-affinity binders[cite: 9, 10]. Finally, the enriched sequences are identified using sequencing[cite: 9, 10]. Over the years, several variations of SELEX have been developed to improve specificity and applicability[cite: 9, 10]. These include cell-SELEX (targeting whole cells), in vivo SELEX, and toggle SELEX (for cross-reactivity across related targets)[cite: 9, 10]. Advances in next-generation sequencing have also enabled deeper analysis of sequence enrichment across rounds, providing insights into selection dynamics[cite: 9, 10].
Despite its widespread use, SELEX has several important limitations[cite: 9, 10]. First, it is a time-consuming and labor-intensive process, often requiring weeks to months to complete[cite: 9, 10]. Second, the iterative amplification steps can introduce biases, favoring sequences that amplify efficiently rather than those that bind best[cite: 9, 10]. Third, SELEX primarily selects for affinity (how strongly something binds) but provides limited information about kinetics (how fast binding and unbinding occur), which are equally important for many applications[cite: 9, 10]. Additionally, the selection is typically performed under specific conditions that may not reflect real biological environments, leading to aptamers that perform poorly outside the selection setup[cite: 9, 10]. Another major challenge is that SELEX does not directly provide structural or mechanistic insight into why a particular aptamer binds well[cite: 9, 10]. It identifies “winners,” but not necessarily the underlying rules that govern binding[cite: 9, 10]. This limits our ability to rationally design better aptamers[cite: 9, 10].
Before the development of high-throughput single-molecule techniques, several alternative methods were used alongside or after SELEX to characterize aptamer binding[cite: 9, 10]. Techniques such as surface plasmon resonance (SPR) and isothermal titration calorimetry (ITC) provided measurements of binding affinity and thermodynamics[cite: 9, 10]. Fluorescence-based assays, including FRET and fluorescence anisotropy, enabled studies of binding interactions and conformational changes[cite: 9, 10]. Fluorescence correlation spectroscopy (FCS) allowed analysis of molecular interactions in solution[cite: 9, 10]. However, these methods are generally low-throughput and require testing one sequence at a time, making it impractical to explore large sequence spaces comprehensively[cite: 9, 10].
As a result, while SELEX is highly effective at discovering aptamers, it offers limited scalability in understanding sequence-function relationships[cite: 9, 10]. This gap has driven the development of newer approaches that combine high-throughput screening with detailed kinetic and mechanistic insight[cite: 9, 10].
SPARXS: A State-of-the-Art Technique for High-Throughput Aptamer Screening
SPARXS (Single-molecule Parallel Analysis for Rapid eXploration of Sequence space) represents a new generation of technologies designed to overcome the limitations of traditional aptamer discovery and characterization methods[cite: 9, 10]. It combines the strengths of single-molecule fluorescence microscopy with next-generation sequencing to enable simultaneous analysis of hundreds to thousands of sequences in a single experiment[cite: 9, 10].
The key innovation of SPARXS lies in its ability to link sequence identity with real-time molecular behavior at the single-molecule level[cite: 9, 10]. In this approach, a library of nucleic acid sequences (such as aptamers) is immobilized on a sequencing chip, typically an Illumina flow cell[cite: 9, 10]. Each molecule is spatially separated and can be observed individually using total internal reflection fluorescence (TIRF) microscopy[cite: 9, 10]. This allows researchers to record fluorescence time traces that capture binding and unbinding events in real time[cite: 9, 10]. After the imaging step, the same molecules are sequenced using standard next-generation sequencing workflows[cite: 9, 10]. Because the spatial position of each molecule is preserved, the fluorescence data can be directly matched to the corresponding sequence[cite: 9, 10]. This creates a powerful dataset where each sequence is associated with its dynamic behavior, including binding frequency, dwell times, and kinetic rates[cite: 9, 10].
One of the major advantages of SPARXS is its throughput[cite: 9, 10]. Unlike traditional single-molecule experiments that study one sequence at a time, SPARXS enables parallel analysis of large libraries under identical experimental conditions[cite: 9, 10]. This not only saves time but also allows direct comparison between variants, revealing subtle sequence-dependent effects that would otherwise be missed[cite: 9, 10]. Another key strength is its ability to measure kinetics, not just affinity[cite: 9, 10]. By observing individual binding and unbinding events, SPARXS provides access to association and dissociation rates, offering a more complete understanding of molecular interactions[cite: 9, 10]. This is particularly important for applications such as biosensing and drug design, where the speed of binding can be as critical as the strength. SPARXS also enables the discovery of unexpected or rare behaviors[cite: 9, 10]. Because it does not rely on iterative selection like SELEX, it can capture a broader diversity of functional sequences, including those that might be lost during enrichment steps[cite: 9, 10]. This makes it especially powerful for uncovering new sequence motifs and understanding the underlying principles of molecular recognition[cite: 9, 10].
In summary, SPARXS represents a significant step forward in aptamer research[cite: 9, 10]. By combining high-throughput screening with single-molecule resolution, it bridges the gap between sequence and function[cite: 9, 10]. This enables not only faster discovery of aptamers but also deeper insight into the mechanisms that govern their behavior, paving the way for more rational and efficient design of molecular recognition tools[cite: 9, 10].
Current State of the Art within DYNAMO (2026)
Aptamer sensing of proteins using optical microscopy and nanopores for single-molecule protein analysis
Aptamers are short strands of DNA or RNA that can bind specifically to a target molecule, such as a protein, small molecule, or even a whole cell[cite: 9, 10]. The most widely used method to discover aptamers is SELEX, which stands for Systematic Evolution of Ligands by EXponential enrichment[cite: 9, 10]. SELEX is an iterative selection process that mimics natural evolution in the laboratory to identify sequences with high binding affinity and specificity[cite: 9, 10].
The SELEX process begins with a very large library of random nucleic acid sequences often containing up to 10¹³–10¹⁵ unique molecules[cite: 9, 10]. This library is exposed to a target of interest (for example, a protein)[cite: 9, 10]. A small fraction of sequences in the library will bind to the target, while the majority will not[cite: 9, 10]. The bound sequences are then separated from the unbound ones, typically through washing steps[cite: 9, 10]. The selected sequences are amplified using PCR (for DNA) or reverse transcription followed by PCR (for RNA), generating a new enriched pool[cite: 9, 10]. This cycle of binding, separation, and amplification is repeated multiple times (usually 8–15 rounds), gradually enriching the pool for high-affinity binders[cite: 9, 10]. Finally, the enriched sequences are identified using sequencing[cite: 9, 10]. Over the years, several variations of SELEX have been developed to improve specificity and applicability[cite: 9, 10]. These include cell-SELEX (targeting whole cells), in vivo SELEX, and toggle SELEX (for cross-reactivity across related targets)[cite: 9, 10]. Advances in next-generation sequencing have also enabled deeper analysis of sequence enrichment across rounds, providing insights into selection dynamics[cite: 9, 10].
Despite its widespread use, SELEX has several important limitations[cite: 9, 10]. First, it is a time-consuming and labor-intensive process, often requiring weeks to months to complete[cite: 9, 10]. Second, the iterative amplification steps can introduce biases, favoring sequences that amplify efficiently rather than those that bind best[cite: 9, 10]. Third, SELEX primarily selects for affinity (how strongly something binds) but provides limited information about kinetics (how fast binding and unbinding occur), which are equally important for many applications[cite: 9, 10]. Additionally, the selection is typically performed under specific conditions that may not reflect real biological environments, leading to aptamers that perform poorly outside the selection setup[cite: 9, 10]. Another major challenge is that SELEX does not directly provide structural or mechanistic insight into why a particular aptamer binds well[cite: 9, 10]. It identifies “winners,” but not necessarily the underlying rules that govern binding[cite: 9, 10]. This limits our ability to rationally design better aptamers[cite: 9, 10].
Before the development of high-throughput single-molecule techniques, several alternative methods were used alongside or after SELEX to characterize aptamer binding[cite: 9, 10]. Techniques such as surface plasmon resonance (SPR) and isothermal titration calorimetry (ITC) provided measurements of binding affinity and thermodynamics[cite: 9, 10]. Fluorescence-based assays, including FRET and fluorescence anisotropy, enabled studies of binding interactions and conformational changes[cite: 9, 10]. Fluorescence correlation spectroscopy (FCS) allowed analysis of molecular interactions in solution[cite: 9, 10]. However, these methods are generally low-throughput and require testing one sequence at a time, making it impractical to explore large sequence spaces comprehensively[cite: 9, 10].
As a result, while SELEX is highly effective at discovering aptamers, it offers limited scalability in understanding sequence-function relationships[cite: 9, 10]. This gap has driven the development of newer approaches that combine high-throughput screening with detailed kinetic and mechanistic insight[cite: 9, 10].
SPARXS: A State-of-the-Art Technique for High-Throughput Aptamer Screening
SPARXS (Single-molecule Parallel Analysis for Rapid eXploration of Sequence space) represents a new generation of technologies designed to overcome the limitations of traditional aptamer discovery and characterization methods[cite: 9, 10]. It combines the strengths of single-molecule fluorescence microscopy with next-generation sequencing to enable simultaneous analysis of hundreds to thousands of sequences in a single experiment[cite: 9, 10].
The key innovation of SPARXS lies in its ability to link sequence identity with real-time molecular behavior at the single-molecule level[cite: 9, 10]. In this approach, a library of nucleic acid sequences (such as aptamers) is immobilized on a sequencing chip, typically an Illumina flow cell[cite: 9, 10]. Each molecule is spatially separated and can be observed individually using total internal reflection fluorescence (TIRF) microscopy[cite: 9, 10]. This allows researchers to record fluorescence time traces that capture binding and unbinding events in real time[cite: 9, 10]. After the imaging step, the same molecules are sequenced using standard next-generation sequencing workflows[cite: 9, 10]. Because the spatial position of each molecule is preserved, the fluorescence data can be directly matched to the corresponding sequence[cite: 9, 10]. This creates a powerful dataset where each sequence is associated with its dynamic behavior, including binding frequency, dwell times, and kinetic rates[cite: 9, 10].
One of the major advantages of SPARXS is its throughput[cite: 9, 10]. Unlike traditional single-molecule experiments that study one sequence at a time, SPARXS enables parallel analysis of large libraries under identical experimental conditions[cite: 9, 10]. This not only saves time but also allows direct comparison between variants, revealing subtle sequence-dependent effects that would otherwise be missed[cite: 9, 10]. Another key strength is its ability to measure kinetics, not just affinity[cite: 9, 10]. By observing individual binding and unbinding events, SPARXS provides access to association and dissociation rates, offering a more complete understanding of molecular interactions[cite: 9, 10]. This is particularly important for applications such as biosensing and drug design, where the speed of binding can be as critical as the strength. SPARXS also enables the discovery of unexpected or rare behaviors[cite: 9, 10]. Because it does not rely on iterative selection like SELEX, it can capture a broader diversity of functional sequences, including those that might be lost during enrichment steps[cite: 9, 10]. This makes it especially powerful for uncovering new sequence motifs and understanding the underlying principles of molecular recognition[cite: 9, 10].
In summary, SPARXS represents a significant step forward in aptamer research[cite: 9, 10]. By combining high-throughput screening with single-molecule resolution, it bridges the gap between sequence and function[cite: 9, 10]. This enables not only faster discovery of aptamers but also deeper insight into the mechanisms that govern their behavior, paving the way for more rational and efficient design of molecular recognition tools[cite: 9, 10].
State of the art