How to Build an RPA-CRISPR Assay for Rapid Diagnostics

To this date, Cas12 and Cas13 systems are the primary drivers of signal generation in CRISPR-based assays due to their collateral cleavage activity. Cas12 is most commonly used for DNA detection following RPA amplification, while Cas13 enables RNA detection in both direct and amplification-coupled workflows. For DNA-targeted detection workflows, engineered Cas12 nucleases are often explored to improve signal-to-noise performance. Synthego’s hfCas12Max, a high-fidelity Cas12i-family nuclease, is designed to reduce non-specific collateral activity while retaining target-triggered activation and can be evaluated in CRISPR-based detection workflows.

Cas 12 and Cas13 systems convert target recognition into measurable outputs through fluorescent reporter cleavage. Fluorescent reporters support real-time or quantitative detection, while lateral flow formats enable rapid, instrument-free readouts. In lateral flow systems, signal interpretation depends on reporter cleavage state, where intact and cleaved reporters produce distinct visual outputs. The choice between endpoint and real-time detection ultimately depends on assay requirements such as sensitivity, quantitation, and available instrumentation, with each format offering trade-offs in speed, complexity, and data resolution. For a deeper comparison, see our endpoint versus real-time RPA guide.

In contrast, Cas9 lacks collateral activity and is typically used in engineered detection systems where signal generation is externally coupled. While this makes Synthego enzymes like SpCas9 and high-fidelity variants such as eSpOT-ON exceptionally precise and reliable for applications like gene editing—where controlled, site-specific cleavage is essential—it limits their utility in diagnostics. Without intrinsic signal amplification through collateral cleavage, Cas9-based assays often require additional components or steps, increasing complexity and potentially reducing speed and sensitivity compared to systems built around enzymes like Cas12 or Cas13.

CRISPR targeting is constrained by sequence requirements: Cas12 relies on protospacer adjacent motifs (PAMs), while Cas13 does not require a PAM but may exhibit protospacer flanking site (PFS) preferences that influence targeting efficiency. Recent advances have introduced PAM-independent detection strategies—such as engineered primer ratios (e.g., asymmetric RPA) that generate ssDNA substrates—enabling Cas12-mediated detection without reliance on canonical PAM sequences and significantly expanding the range of targetable regions. Complementary approaches that pair these PAM-free workflows with iterative crRNA design further enhance assay specificity and robustness, particularly for challenging targets.

The precision of any CRISPR-based diagnostic is entirely dependent on the design of the guide RNA (gRNA), which serves as the search engine for the Cas enzyme. While the RPA step provides the bulk of the material, the gRNA ensures that only the correct sequence triggers a signal. A primary concern in design is off-target activity, where the gRNA might bind to a sequence with 1–3 mismatches, potentially causing false positives. To mitigate this, computational tools are used to screen gRNAs against the host genome and related pathogens, ensuring the "spacer" sequence is unique to the target. Synthego synthetic guide RNAs (sgRNAs) provide a consistent and scalable source of sequence-specific targeting, enabling rapid assay optimization and streamlined workflow integration across CRISPR-based detection systems.

Furthermore, when multiplexing, gRNAs must be meticulously screened for cross-reactivity. In these high-plex scenarios, the challenge is ensuring that the gRNA for Target A does not inadvertently activate the Cas protein in the presence of Target B, while also maintaining a stable secondary structure that prevents the gRNAs from binding to one another rather than the viral template.

Before beginning assay design, it is essential to define both the starting material and the intended amplification format, as these parameters dictate the appropriate chemistry and kit selection.

Note: The "amplification readout" refers specifically to signal monitoring associated with the RPA reaction itself. This is independent of the final diagnostic readout generated by downstream detection systems—such as CRISPR-Cas cleavage, lateral flow strips, or fluorescent reporters—in either one-pot or two-pot workflows.

Establishing these parameters upfront ensures the selection of the correct enzymatic system and helps prevent workflow mismatches between the amplification and downstream detection stages.

The ideal amplicon is typically 80–200 bp. This range balances fast amplification with accessibility for CRISPR detection. Shorter regions tend to amplify quickly but may reduce flexibility in guide design, while longer regions increase the risk of secondary structure that can interfere with detection.

At this stage, it is also essential to confirm:

Once the target region is validated, it becomes the foundation for both primer and guide RNA design.

With the target defined, the next step is to build the amplification system. RPA primers are designed first—typically starting with 3–5 primer pairs as candidates, each around 30–35 nucleotides. These are screened experimentally to identify the most efficient and specific pair. Importantly, RPA is tested on its own, without CRISPR components. This isolates amplification performance from downstream detection.

A successful RPA reaction should:

At the end of this step, you should have a single optimized primer pair that reliably generates clean amplicons.

In parallel with RPA optimization, the CRISPR detection system is developed independently. Multiple guide RNAs (typically 3–6 gRNAs) are designed within the validated amplicon region. These are screened using a synthetic target under fixed reaction conditions. Again, the system is tested in isolation using a defined DNA or RNA target, Cas enzyme, and reporter.

A strong CRISPR candidate will show:

At this point, the assay has two independently validated components: one for amplification and one for detection.

Once both modules are functional, they are combined for the first time using a two-pot system.

In this setup:

This dilution step is important because it reduces carryover of RPA reagents that might inhibit Cas activity. In most cases, no additional cleanup is required. However, if the synthetic control works but the RPA product does not, this often indicates reaction inhibition. In those cases, a quick bead or column cleanup can confirm whether the issue is chemical interference rather than assay design. At this stage, different primer pairs and gRNAs are systematically combined to identify the best-performing pair.

With a working RPA–CRISPR pair established, the system is now optimized as a single biochemical unit. Unlike earlier steps, changes here affect both amplification and detection simultaneously.

Key parameters include:

The goal is not just activity, but consistent, reproducible signal generation across runs.

Once the system is stable, it is adapted for deployment. At this stage, the key decision is whether to use a two-pot or one-pot format. The two-pot format is typically used during development because it allows independent optimization of RPA and CRISPR conditions. It offers maximum flexibility and sensitivity but increases handling steps and contamination risk. The one-pot format combines everything into a single reaction. This reduces user steps and contamination risk but requires careful tuning of buffer compatibility and enzyme stability. It is generally reserved for finalized assays.

The final assay format depends on the intended use case:

Each readout imposes different constraints on reaction chemistry and reporter design.

Before deployment, the assay must be validated beyond ideal laboratory conditions.

This includes:

These tests ensure the assay is robust to biological variability and not just optimized for synthetic targets.

While adding the IVT step increases the workflow complexity—typically requiring a two-pot process—the trade-off is a massive gain in sensitivity. This is achieved through a three-tier amplification cascade:

In platforms like SHERLOCK, the IVT and Cas13 steps are combined into a single reaction mix. This allows for real-time detection: as the T7 polymerase produces RNA, Cas13 immediately recognizes it and triggers a signal. This synergy ensures that even when starting with a few copies of DNA or RNA, the system produces a robust, high-confidence result, often reaching attomolar sensitivity.

A widely used approach for in-tube multiplexing is the use of orthogonal CRISPR-associated enzymes with distinct target preferences and reporter requirements. For example, Cas12 enzymes are typically used for DNA targets and cleave single-stranded DNA reporters, while Cas13 enzymes are used for RNA targets and cleave single-stranded RNA reporters. When combined in a single reaction, this enables simultaneous detection of different target classes. To distinguish signals, each enzyme–reporter pair is assigned a unique fluorescent label (e.g., FAM for Cas12 and HEX for Cas13), allowing independent readouts with minimal spectral overlap.

For multiplexing within the same target class (e.g., multiple DNA or multiple RNA targets), assays generally rely on orthogonal guide RNAs paired with uniquely labeled or barcoded reporters, rather than differences in intrinsic cleavage behavior. This allows multiple targets to be resolved even when using a single effector protein. Because Cas12 and Cas13 also differ in their sequence recognition requirements (e.g., PAM constraints for Cas12 and PFS preferences for Cas13), they can be independently programmed to interrogate distinct targets within the same sample without cross-reactivity.

When instrumentation is limited or a visual output is preferred, multiplexing can be achieved through spatial separation of detection signals using lateral flow assays (LFAs).

In this format, amplification and CRISPR detection occur in a single reaction, but target identification is resolved downstream at the level of the test strip. Distinct capture reagents immobilized at separate test lines direct reporter accumulation to defined spatial positions, enabling each target to be read independently (e.g., Line 1 for Target A, Line 2 for Target B). This approach is particularly useful for field-deployable diagnostics, where binary or semi-quantitative interpretation is preferred over fluorescence-based readouts.

Regardless of detection strategy, the primary constraint in multiplex CRISPR assays is often upstream amplification rather than detection chemistry itself. When multiple primer sets are present in a single reaction (e.g., RPA or RT-RPA), competition for reagents can lead to uneven amplification efficiencies across targets.

To mitigate this, primer concentrations are frequently balanced to reduce amplification bias, particularly for fast-amplifying targets that might otherwise dominate reaction resources. In addition, primer design is critical to minimize dimer formation and nonspecific interactions, which can significantly reduce assay sensitivity in multiplex formats.