Discover our step-by-step development process of RPA, DNA hybridization, and CRISPR-Cas experiments, as well as the purification of LbCas12a. This page documents the entire process - from amplification to detection and ultimately towards the creation of a paper-based test. It also explains the reasoning behind our design decisions, the adjustments we made after each test, and the lessons we learned. We followed the engineering design cycle Design → Build → Test → Learn across multiple rounds of experiments. Each iteration refined our methods and informed the next stage of development. Through this systematic progression, we moved from basic amplification methods to reliable detection, leading to a functional diagnostic system.
Even though primers and DNA sequences for all six pathogens were designed on Benchling (see the page “Experiments and Protocols”) and checked on BLAST for specificity, we decided to first experimentally focus on one pathogen. In this way we aimed to establish the whole circuit working for one pathogen and then apply to the rest. We selected HPV16, specifically its L1 gene, because a clinical study [1] demonstrated that HPV16 can be reliably detected in first-void urine at sufficient concentrations, supporting its suitability for non-invasive diagnostic testing. It is worth mentioning that in this study HPV DNA was extracted from the urine samples prior to detection. Among our three HPV16 target sequences, we selected the L1 gene because it showed the highest amplification yield in our Nanodrop measurement (225.2 ng/μL) and is frequently used in diagnostic assays, supporting both assay reliability and clinical relevance.(see more on our ”Notebook” page). Our first goal was to test the three RPA primer pairs designed for RPA and identify the most efficient one. A second goal was to determine whether there was a difference between using DNA at a concentration of 1 ng/μL and 0.0001 ng/μL (= 0.1 pg/μL), since patient samples may contain very low concentrations of pathogen DNA, and the sensitivity of our test depends on detecting these low levels.
The RPA primers were all designed following the instructions provided by TWIST, as we used the TwistAmp basic kit for our RPA reactions. The designed primers were then run in BLAST to be sure that they were unique for HPV16 and wouldn’t bind to other pathogens, thereby preventing the chances of false-positives results. Find the primers in the Experiments and Protocolspage. We ordered the primers from Microsynth and DNA sequences from Twist Bioscience. The kit for RPA was bought from TWIST.
We conducted the RPA following the protocols detailed on our Experiments and Protocols page. A description of the experiments can be found on the Notebook page.
Legend:
The results are very dim because we added too little of our sample, namely only 1 μL out of 50 μL, mistakenly following the same loading volume as for PCR gels. Our instructor Cheng-Han advised us to load at least 10% of the product in the gel, which here would have been 5 μL. We followed his advice in the next experiments.
Based on the gel results, no clear difference was observed between amplifying 1 ng/μL or 0.1 pg/μL, so we decided to perform all subsequent experiments using the lower concentration of 0.1 pg/μL, as this better reflects real-world conditions. According to Table 4 from a study by Baud et al. [2], HPV DNA concentrations in clinical urine samples can range from only a few dozen to a few hundred copies per microliter, depending on sample processing. Since our longest RPA amplicon is approximately 400 bp, 0.1 pg/µL corresponds to roughly 200–300 copies/µL, making this dilution a realistic approximation of the low DNA concentrations found in clinical urine samples. Additionally, the differences between the amplicons in the gel were minor, so we arbitrarily selected the first amplicon for all subsequent experiments.
One mistake we made was not measuring the Nanodrop concentration after purifying the product. Doing so would have allowed us to detect finer differences in concentration. We did not perform this measurement due to some confusion about the validity of Nanodrop readings for RPA, although we later learned that this concern mainly applies to asymmetrical RPA and can be addressed (see “Learn” part of Cycle 2 - Asymmetrical RPA for DNA-hybridization method).
This experiment served as the foundation for our future work and signifies that the first part of the CRISPR detection method is complete.
The CRISPR method can directly work with double-stranded RPA amplicons, which were already achieved in the first cycle. However, the DNA-hybridization method requires the amplicons to be single-stranded as we learned in a conversation with our instructor Cheng-Han (see the Attribution page). Double-stranded DNA would prevent proper hybridization and reduce signal efficiency [3]. That is why we performed asymmetrical RPA, to get single-stranded RPA amplicons. Our goal here was to set up a parameter screening with varying primer concentrations to understand whether the gene sequence had a preference for the forward or reverse primers, and select the optimal ratio. Literature [4] and discussions with our instructor, led us to test three different ratios, namely 1:10, 1:20 and 1:50 (forward:reverse). We had also considered using only one primer and none of the other, but our instructor then explained that we needed an exponential phase of growth at least initially, which is only possible with both primers working together to elongate the strands. Our hypothesis was that, as stated in the paper [4], the concentration of 1:20 would work best and there would not be a preference between the forward and reverse DNA strands. An important consideration to note is that asymmetrical RPA drops the yield compared to regular RPA [4], which is why after every asymmetrical RPA we performed, we purified the product and loaded 10 μL of it in the gels.
We conducted the asymmetrical RPA following the protocols detailed on our Experiments and Protocols page. A description of the experiments can be found on the Notebook page. Results were checked with agarose gel and NanoDrop.
Legend:
| Well numbers | Content 1 | Concentration (ng/μL) |
|---|---|---|
| 1 | 10 reverse : 1 forward , 0.1 pg/μL | N/A due to technical error |
| 2 | 1 reverse : 10 forward , 0.1 pg/μL | 1265.17 |
| 3 | 20 reverse : 1 forward , 0.1 pg/μL | 1917.39 |
| 4 | 1 reverse : 20 forward ,0.1 pg/μL | 2015.26 |
| 5 | 50 reverse : 1 forward ,0.1 pg/μL | 2050.51 |
| 6 | 1 reverse : 50 forward , 0.1 pg/μL | 1732.99 |
| 7 | Positive control (symmetrical RPA with normal conditions) | 3111.1 |
The results were successful for all asymmetrical products. However, there is indeed a difference in the gel between the DNA strands outputs and between the different primer ratios. Diluted forward primers show only one line at the expected location of approximately 400bp but are dimmer than the positive control. While, diluted reverse primers show three bands, which could indicate either there was the formation of DNA secondary structures or dimerization of single-stranded RPA products. To test that we would have to denature them, but due to time constraints we didn’t do that, so we cannot conclude whether one strand performs better than the other. To be safe, we proceeded using only diluted forward primers, which also guided the design of DNA-hybridization primers to target the antisense strand.
Regarding primer ratios, agarose gel brightness and NanoDrop results suggested that 1:20 and 1:50 (forward:reverse) worked better than 1:10, with 1:20 appearing brightest. The first well may have been dimmer due to a technical error, so we could not conclusively determine the best ratio.
It is worth mentioning that the NanoDrop results for asymmetrical RPA amplicons had to be corrected later due to two reasons. First, when using NanoDrop, it was forgotten to switch from the DNA-50 mode to the DNA-33 mode, thus we were measuring ssDNA with the dsDNA function, inflating our results. To correct this, we multiplied the results obtained using DNA-50 by 0.66.Second, excess primers added for amplification were co-purified, further inflating concentrations. Our instructor Cheng-Han, advised us to do an experiment to calculate this overestimation factor by comparing purified asymmetrical RPA amplicons with purified asymmetrical RPA reactions without the enzyme pellet. The results of this experiment showed overestimation factors of 9.57 for 1:10, 15.97 for 1:20 and 26.66 for 1:50 forward:reverse primer ratio (see the page „Notebook: Nanodrop overestimation calculation (22.09.2025)“). To obtain more accurate results, we divided the measured concentrations by these factors. However, this experiment could be further refined by accounting for the actual volume recovered during purification, as a lower recovery volume would result in a more concentrated product. Such an implementation was not carried out due to time constraints.
In conclusion, this was our first step toward establishing DNA-hybridization as a detection strategy.
The goal is to run a parameter screening for different temperatures because this is crucial information for us, as we want our self-test to operate at room temperature (RT). RPA was accomplished by first testing a range of four temperatures (RT = 23.8°C, 25°C, 32°C, 37°C) and two incubation times (20min and 30min). A master mix was prepared so that the only parameters that changed were indeed the temperature and the incubation time. We conducted the RPA following the protocols detailed on our Experiments and Protocols: Isothermal RPA page. A description of the experiments can be found on the Notebook: Isothermal RPA page.
Legend:
The results on the gel don’t look clear, that is why we decided to attempt it again, this time refining our parameters. We included a positive control and another incubation time variable, namely 60 minutes. This time around, the gel results were much clearer, except for sample 14, which was the only one prepared using a different stock of DNA. We thus assumed that the DNA aliquot used, had an issue.
What we learned from the agarose gel and NanoDrop results is that: RPA is robust across a wide temperature range (25-37 °C), but when performed at room temperature (~23 °C) it requires a longer incubation time to achieve amplification. At 20 minutes, yields at room temperature were very low, but after 30 minutes they reached levels similar to those obtained at 25-37 °C. This indicates that for our project, RPA can be reliably performed without equipment at ambient conditions, provided the reaction time is adjusted to around 30 minutes.
| Temperature [°C] | Incubation time [min] | Order of gel | DNA amount [ng/μL] |
|---|---|---|---|
| 39 | 20 | 1 (positive control) | 106 |
| 39 | 20 | 2 (negative control) | 33.4 |
| RT / ~23.8°C | 20 | 3 | 12.9 |
| 25 | 20 | 4 | 38.7 |
| 32 | 20 | 5 | 34.1 |
| 37 | 20 | 6 | 83.5 |
| RT / ~23.8°C | 30 | 7 | 87.3 |
| 25 | 30 | 8 | 75.2 |
| 32 | 30 | 9 | 103.5 |
| 37 | 30 | 10 | 121.4 |
| RT / ~23.8°C | 60 | 11 | 108.3 |
| 25 | 60 | 12 | 96.1 |
| 32 | 60 | 13 | N/A |
| 37 | 60 | 14 | 51.1 |
After having achieved the first steps for our two detection methods and experimented with different temperatures and incubation times, the next big step was to test RPA on the strip. We are happy to say that for this, we used the protocol “RPA on Paper“, written by the iGEM 2017 Munich team, “CascAID” (see the Experiments and Protocols page). Our goal was to make this work for symmetrical and asymmetrical RPA, which is necessary for our two detection methods, even though their protocol was only designed for the former. A description of the experiments can be found on the Notebook page.
Legend left:
Legend left:
| Concentration of DNA | DNA amount [ng/μL] |
|---|---|
| 1 ng/μL | 3.7 |
| 0.1 pg/μL | -0.8 |
Symmetrical RPA on paper strips yielded only a 3-fold increase, which is technically amplification but represents a very low efficiency compared to expectations. This highlights the need for protocol optimization. Upon discussion with our instructor Cheng-Han, we identified several potential issues and troubleshooting strategies.
First, to minimize enzyme degradation, he recommended avoiding repeated lyophilization of the enzyme pellet. Instead, only the primers and rehydration buffer should be freeze-dried directly on the strip, while the enzymes and probe are added afterwards.
Second, harvesting efficiency may be a limiting factor, since our current workflow involves soaking the paper in water and purifying the eluate. To evaluate this, we could compare NanoDrop results from liquid-phase RPA reactions with those from liquid-phase RPA reactions subsequently applied to the strip and then purified.
Finally, to ensure complete wetting of the strip and reduce evaporation losses, it was suggested to load twice the amount of reaction mixture onto the strip and optimize the reaction volumes.
While these insights provide a clear roadmap for optimization in the next Design → Build → Test → Learn cycle, we were not able to fully implement them within the available timeframe.
One challenge for our prototype is the lysis of pathogens, ensuring that our methods can access their DNA and reliably detect even very small amounts. To address this, we tested a broader range of diluted DNA concentrations beyond the already tested 0.1 pg/µL. Specifically, we extended our experiments to 0.01 pg/µL and 1 fg/µL for both symmetrical and asymmetrical RPA. These reactions were performed with diluted DNA templates using our existing stocks of primers and DNA. We conducted the RPA following the protocols detailed on our Experiments and Protocols: Diluted RPA page. A description of the experiments can be found on the Notebook: Failed diluted RPA (28-29.07) page..
When we ran the reactions, they stopped working altogether. On agarose gels, we observed either smears or no bands at all. In some cases, even the DNA ladder appeared unreliable, which suggests that both the reaction itself and our gel conditions could be contributing to the unclear results.
As a first troubleshooting step, we suspected a degraded DNA aliquot and prepared fresh aliquots from the original purchased tube, but the problem persisted (Notebook: diluted RPA troubleshooting template (30.07)). Since ordering a new DNA stock would have delayed our project by at least a week, we instead prepared fresh primer aliquots, but results did not improve (Notebook: diluted RPA troubleshooting primers (31.07)). Moreover, opening a new RPA kit (Notebook: normal RPA (8.8.25)) and testing alternative templates (Notebook: failed RPA Syphilis (31.07) and RPA (11.08.25)) also failed, suggesting that the issue was not specific to the target DNA.
We further learned that gel quality control played a role: inconsistent ladders were linked to buffer replacement frequency, so we adopted our instructor’s advice to refresh the gel buffer every three days when running many gels, instead of only once a week. Another lesson was to prepare individual aliquots of nuclease-free water for each experiment to reduce the risk of contamination. As an alternative way to get RPA amplicons for our next applications, namely our two detection methods, we performed a PCR using RPA primers to generate RPA amplicons (Notebook: PCR to get RPA amplicons (14.08)), which the first time produced successful NanoDrop and agarose gel results. However, this method did not work again afterwards.
Legend:
Finally, after following the advice to mix and tap the aliquots after freeze-thawing, we obtained a smeared but correctly sized band with HPV16, showing that proper template handling was critical for RPA success (Notebook: RPA (14.08.25)). We later realized that a more effective approach would have been to sequence the DNA stock and to purchase a replacement tube as soon as RPA experiments stopped working.
Due to time constraints, we didn’t return to testing highly diluted RPA templates.
At the beginning, our aim was simple: demonstrate that DNA hybridization works as a detection method for our target pathogen sequence. We designed complementary DNA oligonucleotides, with one serving as the template (RPA or PCR amplicons of the HPV16 L1 gene) and the other carrying a fluorescent label. To explore how sequence length and stability influence sensitivity and specificity in DNA hybridization, we tested three variants: a 15-mer probe with LNA modifications, a 20-mer DNA probe, and a 25-mer DNA probe, each labeled with the fluorophore Dy681.To provide templates, we prepared products from symmetrical RPA (dsDNA), asymmetrical RPA (ssDNA), and PCR. All probes and primers were checked with BLAST to ensure specificity to HPV16.
Each probe was rehydrated to 100 µM stock and used at 1:1000 dilution (working ~100 nM). For every reaction we prepared a 10 µL hybridization mix: 1 µL probe, 1 µL 0.5 M NaCl, 7 µL nuclease-free water, and 1 µL template. Templates came from symmetrical RPA (dsDNA; 1 ng input; 1:1 primers), asymmetrical RPA (ssDNA-enriched; 1:50 forward:reverse), and PCR (≈200–210 ng product). Negative controls contained water instead of template.
In our first attempt, we combined the probes with symmetrical RPA, asymmetrical RPA, and PCR products. We denatured at 95 °C for 5 min, cooled to 25 °C, and incubated ≥30 min. Products were run on 1.5% agarose with ROTI® Gel Stain and were read on the fluorescence imager.
Legend:
Single-probe hybridization was successful across all template types. Symmetrical RPA (lanes 5-7) gave the brightest signals, while asymmetrical RPA (lanes 2-4) and PCR (lanes 8-10) showed weaker but still visible bands. Among the probes, the 15-mer with LNA (lanes 2,5 and 8) performed best. Based on these results, we decided to adopt the LNA probe as our standard and move on to testing more challenging conditions such as at room-temperature hybridization and specificity against other pathogens.
For our test to be practical on a paper strip, hybridization needed to work at room temperature without a denaturation step. In addition, we had to prove that our probe was specific to HPV16 and would not cross-react with DNA from other pathogens. We therefore designed an experiment to test asymmetrical RPA products at room temperature and to run hybridization reactions against PCR products of several non-target pathogens (Chlamydia, Gonorrhoea, Syphilis, Trichomonas).
For asymmetrical RPA at RT we used 1:50 forward:reverse primers, incubated at ~23 °C for ≥35 min. For specificity, we used PCR products (~200 ng) from the non-targets. Hybridization mixes remained 10 µL with the 15-mer LNA probe at 1:1000, but for the asymmetrical RPA tube we sometimes increased the volume of the DNA to 2 µL and reduced water accordingly (6 µL) to compensate for lower amplification yield.
For the pathogen PCR products, we used the standard profile of 95 °C denaturation for 5 min followed by 25 °C annealing for ≥30 min. For the asymmetrical RPA, we ran the reaction at room temperature only, without denaturation. Products were analyzed on 1.5% agarose gels with fluorescence readout and were read on the fluorescence imager.
Legend:
Two important outcomes came from this cycle:
We hypothesized that the ratio of forward to reverse primers in asymmetrical RPA influences the production of ssDNA and thus the success of hybridization. To increase ssDNA yield, we compared two asymmetrical RPA primer ratios: 1:10 (forward : reverse) and 1:20 (forward : reverse), combined with different template inputs (1 ng/µL and 0.1 pg/µL ).
We produced fresh asymmetrical RPA reactions at 1:10 and 1:20. Hybridization kept the 15-mer LNA probe at 1:1000 in a 10 µL mix: 1 µL probe, 1 µL 0.5 M NaCl, 7 µL water, 1 µL template (in some repeats we increased template to 3 µL and reduced water to 5 µL to improve sensitivity). Normal RPA and a no-template control were run in parallel.
Thermal profile remained 95 °C 5 min → 25 °C ≥30 min (unless explicitly RT only). Gels were 1.5%, read by fluorescence.
Legend:
The 1:10 (forward:reverse) asymmetrical RPA (lanes 2-3) gave robust hybridization with 1 ng and with 0.1 pg DNA template. The 1:20 (forward:reverse) asymmetrical RPA (lanes 4-5) worked with 1 ng but failed with 0.1 pg DNA template, indicating reduced ssDNA availability. We standardized on asymmetrical RPA 1:10 (forward:reverse) for downstream work.
We introduced a second Cy7-labeled probe with a distinct emission wavelength to enable two-site hybridization on the same target, facilitating multiplexing and providing stronger confirmation of true positives. In addition, this probe design allows one site to function as an anchoring probe, potentially biotin-labeled for attachment to the lateral flow strip, while the other can be visualized through a gold nanoparticle-labeled detection probe. To reduce photobleaching and maintain signal intensity, the high-λ channel was read first.
Both probes were prepared at 100 µM stocks and used at 1:1000 dilution. For single-template double binding, we used a 10 µL hybridization mix: 1 µL Probe A (low-λ), 1 µL Probe B (high-λ), 1 µL 0.5 M NaCl, 6 µL water, 1 µL template. For low-yield asymmetrical RPA we sometimes used 2 µL template and 5 µL water. Templates were symmetrical RPA 1:1 (forward:reverse) (1 ng/µL) and asymmetrical RPA 1:10 (forward:reverse) (both 1 ng/µL and 0.1 pg/µL). Early runs used working aliquots; after suspected dye degradation we switched to fresh aliquots directly from the original stock and handled them under light protection.
We followed the denaturation profile for hybridization (95 °C 5 min → 25 °C ≥30 min) when template allowed. Fluorescence was recorded high-λ first, then low-λ, to minimize bleaching. Gels were 1.5% agarose.
Legend:
Legend:
In the first three double DNA hybridization experiments, only one channel (Dy681) consistently showed signal. The high-λ fluorophore (Cy7) had likely been over-exposed during previous handling. After switching to fresh, light-protected aliquots and adjusting the readout order, double hybridization was successful. On symmetrical RPA, fluorescence was clearly visible in both channels (Figures 12 & 13, lane 4). The same result was then achieved with asymmetrical RPA at a 1:10 forward:reverse ratio using both 1 ng and 0.1 pg inputs (Figures 14 & 15, lanes 2-3). These results demonstrated that two independent probes can bind simultaneously to dsDNA (symmetrical RPA) as well as ssDNA-enriched (asymmetrical RPA) templates under our experimental conditions.
After establishing reliable single- and double-probe DNA hybridization in gels, the next step was to transfer the method onto a paper strip to evaluate whether hybridization could also be detected in a lateral flow test. We used the Milenia Biotec HybriDetect kit and designed the experiment with two different probes, one labeled with FAM and the other with Biotin, so that double binding could be read out directly on the strip. In this test, anti-FAM antibodies conjugated with gold nanoparticles bind the FAM label and, together with capture of the biotinylated partner at the test line, generate the visible red signal. To systematically test sensitivity, we combined two probe concentrations (100 µM and 0.1 pM) with two symmetrical RPA product concentrations (1 ng/ µL and 0.1 pg/µL).
Both FAM- and Biotin-labeled oligonucleotides were rehydrated to 100 µM stock. For annealing, we prepared 20 µL mixes containing 2 µL FAM probe, 2 µL Biotin probe, 2 µL 0.5 M NaCl, 12 µL nuclease-free water, and 2 µL template (symmetrical RPA product). The mixes were denatured at 95 °C for 5 min, cooled to 25 °C, and incubated for ≥30 minutes. Afterwards, the 20 µL hybridization products were pipetted directly onto the sample application pad of the HybriDetect strips. Strips were then placed upright into 100 µL assay buffer and left at room temperature until the bands developed.
Within 2-5 minutes, control and test lines appeared depending on the probe and template input.
Legend:
This experiment showed that DNA hybridization can successfully be detected on a paper strip, marking an important milestone toward a paper-based diagnostic. At 100 µM probe concentration, valid test results were achieved for both undiluted (1 ng) and diluted (0.1 pg) symmetrical RPA products. At 0.1 pM probe concentration, however, no signal was visible, indicating that the input was too low to be detected on the strip under our current conditions. These results highlight the importance of sufficient probe concentration for reliable readout and provide a clear direction for optimization in future work.
We decided to express and purify the Cas protein used in our experiments ourselves. The vectors, materials and necessary equipment were kindly provided to us by the Jinek Laboratory at the University of Zurich. Christelle Chanez, member of the Jinek Lab, assisted us in the process.
Two Cas12a vectors were available, pDS113 and pDS115. Since pDS115 contained a GFP addition which was not necessary for our experiments, we went ahead with the pDS113 vector. The vector was sent for sequencing before use to make sure it was intact. From the lab's experience with these specific vectors, we knew that the expression worked well in Rosetta2 competent cells. We therefore chose to conduct the pDS113 expression in Rosetta2 cells.
We transformed the pDS113 vector into Rosetta2 cells according to the heat shock protocol. The following day we observed a number of healthy colonies on the plate, indicating that the transformation was successful and the bacteria had acquired the antibiotic resistance encoded in the vector.
We picked a colony and inoculated our starter culture overnight with 50 µg/mL kanamycin and 33 µg/mL chloramphenicol. The starter culture was then used to inoculate 6L of LB medium with 50 µg/mL kanamycin the next day. We observed the OD600 of our culture with regular measurements and cooled for 30min at 18°C with shaking once OD600 reached 0.61. To be able to test successful expression later, we took a sample of the uninduced culture. Finally we induced expression with 0.4 mM IPTG (isopropyl β-D-1-thiogalactopyranoside) for overnight expression. The following day we were able to harvest our protein. After spinning down the culture we resuspended the pellet in our lysis buffer.
To verify the correct expression of our protein we ran a SDS-PAGE with the uninduced and induced probe. We expected a band at 148 kDa in the induced sample, but not in the uninduced. However, only an extremely faint band was visible in the induced sample. Instead it seemed as if something was retained in the wells of the induced samples.
These test results could have different explanations. We first went through our protocol to exclude human error. We then concluded that the expression was unexpectedly low. We also considered the issue to be the lysis of the cells. In this case, the additional lysis in the Maximator Cell homogenizer might resolve the problem. Low protein expression might be explained if the chosen bacterial strain, Rosetta2, was not well suited to express this specific vector. We therefore decided to conduct a test expression with different bacterial strains, to investigate how well they express our vector.
To determine the best-suited bacterial strain for expressing the pDS113 vector carrying LbCas12a, we conducted a test expression. The following strains were chosen: BL21 Star, BL21 RIL, Rosetta2, T7 Express, and BL21 Arabinose-Induced. All of these strains are compatible with T7-driven expression systems, but differ in codon usage compatibility, protein folding support, and overall expression efficiency.
Chemically competent cells of each strain were transformed with the pDS113 vector and plated on LB agar containing the appropriate antibiotic. Single colonies were picked and grown overnight in LB. The following day, cultures were inoculated into fresh medium and grown until mid-log phase. Protein expression was induced with 0.5 mM IPTG, and the cultures were incubated for an additional 3 hours at 37°C. Samples were collected for both induced and uninduced conditions.
After expression, cells were harvested by centrifugation and lysed using a standard lysis buffer. The lysates were then incubated with Ni-NTA magnetic beads to test for His-tagged protein binding. Elution fractions were analyzed by SDS-PAGE to evaluate the level of LbCas12a expression. We compared band intensities between strains and between induced and uninduced conditions.
All strains expressed LbCas12a to varying degrees, but Rosetta2 showed the clearest difference between uninduced and induced samples. In Rosetta2 lysates, the LbCas12a band was more intense and more protein was successfully pulled down using the nickel beads, indicating better expression and solubility. This confirmed the initial choice for Rosetta2 as the most suitable host strain for the large-scale protein purification of LbCas12a.
Following the results of the test expression we proceeded with large-scale cultures of Rosetta2 cells.
However, since we were unsure of what caused the unexpected pattern in the initial gel, we wanted to prepare for the case that the large-scale expression with the pDS113 vector would not be successful and we would not be able to obtain sufficient protein for our future experiments. We therefore decided to start the transformation and expression of the alternative vector pDS115 in Mach1 cells, while working in parallel on the large-scale expression and purification of the pDS113 culture.
We also wanted to make sure that the problem does not lie in the cell lysis. Therefore we used a different lysis machine for each culture.
Transformation, expression and induction were carried out according to protocol. For the pDS113 culture the Maximator Cell homogenizer was used again to lyse the cells. The pDS115 culture was lysed with the Cell Lyser 1.
The different lysis method did not impact the outcome of the uninduced vs induced gels. We proceeded with the purification and did not encounter further indication that the protein expression level was as low as the initial gel indicated.
The unexpected gel pattern therefore remains unexplained, but did not result in any consequences for the success of the protein recovery.
The protein purification was carried out according to protocol, and the vectors provided by the Jinek Lab.
The purification process included a Ni-NTA pulldown, an overnight TEV cleavage and dialysis, an ion-exchange chromatography and finally a size-exclusion chromatography.
When running the SDS-PAGE gel after the size exclusion chromatography we observed a double band around 148 kDa. We were unable to separate these two protein populations due to their small differences in size. It seemed plausible that the TEV cleavage had been incomplete and the two bands corresponded to the cleaved and uncleaved protein.
Before the protein was used in further experiments a sample was sent to mass spectrometric analysis at the Functional Genomics Center Zurich to confirm our theory. They were able to confirm the identity of the protein as the cleaved LbCas12a and also noted the presence of a second, slightly bigger protein which might correspond to the uncleaved protein. Based on this we assumed that our purification had yielded a mixed population of TEV-cleaved and uncleaved LbCas12a protein.
We learned that the TEV-cleavage conducted during the overnight dialysis was insufficient to correctly cleave all of the product. To avoid mixed populations in further purifications we would need to separate the cleaved protein from the uncleaved.
To avoid the same issue as in the first purification, we applied what we learned and added an additional separation step to the purification. As the uncleaved protein contains a malE fragment, an MBP trap would be able to separate it from the cleaved protein.
We followed the same protocol for the purification, but added a MBP Trap in tandem to the size exclusion column.
Gels were carried out after Ni-NTA-pulldown, dialysis and size exclusion chromatography. Again a double band was observed in the SDS-PAGE gel after dialysis, but only a single band was visible in the final gel after the SEC and MBP trap chromatography.
Our conclusion that the double band represented a mixed population of cleaved and uncleaved LbCas12a protein has been further confirmed by these results. The MBP Trap resolved the problem by filtering for the cleaved protein. Our product now consists of pure cleaved LbCas12a and the protocol has been adapted for future use.
To verify the activity of our LbCas12a protein, we used a synthetic single-stranded DNA (ssDNA) reporter labeled with the fluorophore ROX at the 5’ end and the quencher BHQ2 at the 3’ end (ROX-N12-BHQ2). In its intact form, the quencher suppresses the ROX fluorescence. Upon cleavage of the reporter by the activated LbCas12a nuclease, the fluorophore is separated from the quencher, resulting in a measurable and visible fluorescence signal at around 602 nm. This allowed us to visually demonstrate the activity and specificity of our system.
The LbCas12a protein, the crRNA and the ssDNA reporter (ROX-N12-BHQ2) were used at a concentration of 100 µM. For a single experiment we used a 50 µL Cas12a mixture: 0.25 µL crRNA, 0.21 µL LbCas12a, 0.1 µL ssDNA reporter, 0.5 µL RNase inhibitor (40 U/ µL), 5 µL NEBuffer 2.1 (10X), 7.5 µL symmetrical RPA product of the HPV16 L1 gene, and 36.44 µL nuclease-free water.
To ensure proper mixing, we first combined all of the reagents except the LbCas12a protein and the crRNA in a single tube and vortexed the mixture. Afterwards, we added the Cas12a protein and crRNA and mixed gently by pipetting up and down. Then we incubated the tube. All in all we tested 3 different incubation temperatures (25°C, 30°C and 37°C) as well as different incubation times (25°C for 60 and 90 minutes, 30°C for 30 and 60 minutes and 37°C for 30 minutes).
During this experiment, we noticed that the concentration of the ssDNA reporter was initially too low to allow visual detection of a color change. The reaction mixture was transparent at the start, and no color shift could be observed by eye. However, under UV light, a faint red fluorescence confirmed that the Cas12a reaction had occurred. To test whether the low reporter concentration was the limiting factor, we added 9 µL of additional ROX-labeled reporter (100 µM) and 1 µL NEBuffer 2.1 to one of the previously incubated reactions.
After 30 minutes at 37°C, the mixture turned from the initial blue to a violet. This change was clearly visible without any UV light or other specialized equipment. Repeating the incubation for a further 30 minutes did not result in a stronger color, suggesting the reaction had reached saturation.
Still, the color intensity was not strong enough for robust visual detection. We concluded that increasing the reporter concentration further would improve visibility and consistency of the signal in future assays.
Legend:
Building on our first assay, where a visible color change was observed only at 37°C, we set out to test whether LbCas12a retains activity at other temperatures, including room temperature. For this we increased the concentration of the ssDNA reporter to improve signal intensity. By systematically varying incubation temperature and time, we aimed to identify the temperature range in which LbCas12a remains active.
For this assay, the LbCas12a protein, the crRNA and the ssDNA reporter were all used at a concentration of 100 µM. For each reaction, we used 50 µL of the following Cas12a mixture: 0.25 µL crRNA, 0.21 µL LbCas12a, 8 µL ssDNA reporter, 0.5 µL RNase inhibitor (40 U/ µL), 6 µL NEBuffer 2.1 (10X), 7.5 µL symmetrical RPA product of the HPV 16 L1 gene, and 27.54 µL nuclease-free water.
The LbCas12a protein and the crRNA were gently mixed in a separate tube by pipetting (vortexing was avoided to protect enzyme activity). The ssDNA reporter, the RNase inhibitor, the NEBuffer 2.1, the symmetrical RPA product and the nuclease-free water were combined and vortexed briefly. Finally, the two mixtures were put together and mixed gently before incubation.
The reactions were incubated at four different temperatures 37°C (positive control), 32°C, 25°C, and room temperature (~ 23 °C). Each condition was monitored by photographing the tubes at the start, and again after 30, 60, and 90 minutes of incubation.
The reaction tubes which were incubated at 37°C and 32°C showed a clear colour shift after 30 minutes of incubation. After another 30 minutes of incubation, the reaction tubes incubated at 25°C and at room temperatures also showed a change in colour. From this experiment we learned that LbCas12a functions most efficiently at elevated temperatures, producing strong and rapid signals at 37°C and 32°C. Nonetheless, the enzyme also retains activity at 25°C and even at room temperature, although it requires longer incubation times of at least one hour to achieve visible results. This finding suggests that the assay is feasible under ambient conditions, which is particularly valuable for our intended application. In the next cycle, we will focus on getting quantitative results for our dry lab with the help of a plate reader. The plan will also be to integrate a negative control, which was not done here.
| Temperature / Sample | 30 minutes of incubation | 60 minutes of incubation | 90 minutes of incubation |
|---|---|---|---|
| Room temperature (approx. 23 °C, Tube 5) | - | + | + |
| 25°C (Tube 4) | - | + | + |
| 32°C (Tube 3) | + | + | + |
| 37°C (positive control, Tube 2) | + | + | + |
Legend:
After verifying Cas12a activity in tube-based assays, the next step was to demonstrate whether target detection could also be achieved in a lateral flow format. We used the Milenia Biotec HybriDetect kit together with a FAM-/Biotin-labeled ssDNA reporter. To systematically assess sensitivity, we tested two concentrations of reporter (50 nM and 100 µM) in combination with two input amounts of symmetrical RPA product (1 ng and 0.1 pg, equivalent to a 10,000× dilution).
We prepared two separate mixes to protect Cas12a activity:
After vortexing tube A briefly and mixing tube B gently by pipetting, tube B was added to tube A to reach a total volume of 20 µL. The mixtures were incubated at 37 °C for 20 minutes. The full reaction volumes were then applied to the sample pads of lateral flow strips, which were placed upright into 100 µL of assay buffer at room temperature.
We prepared four different strips:
After 2-5 minutes, all strips displayed the control line to confirm validity. The test line was visible on each strip, demonstrating successful Cas12a-mediated target recognition and cleavage.
Legend:
This experiment confirmed that LbCas12a detection can be combined with lateral flow readout. According to literature [5], 50 nM reporter should be sufficient, and indeed this worked reliably with both high and diluted RPA inputs. However, even at the higher reporter concentration (100 µM), results were equally positive, suggesting robustness of the system. Importantly, clear signals developed within just 2 minutes across all tested setups. This shows that the Cas12a assay is not only functional but also rapid and compatible with a paper-based format, providing a strong proof of concept for its diagnostic potential.
Building on our experience from the LbCas12a assays, we proceeded to collect a range of data points to support the dry lab in testing their models. This experimental cycle consisted of five distinct experiments, all conducted simultaneously due to logistical constraints. Since the Pelkmans lab did not have access to a plate reader, we collaborated with the Plückthun lab to obtain quantitative readouts. The five experiments focused on measuring background fluorescence, kinetic parameters, the fluorescence signal threshold, a calibration factor, and enzyme denaturation characteristics.
We conducted five assays, each targeting a specific parameter of the LbCas12a system. All reactions used a 100 μM stock concentration of the ssDNA reporter (ROX-N12-BHQ2). The experiments were assembled as follows:
Experiment 1: Reporter Concentration Optimization
Two versions of the Cas12a reaction were prepared, differing in the volume of ssDNA reporter used: 8 μL (positive control) and 4 μL. All other components remained constant.
Experiment 2: Background Fluorescence
LbCas12a reactions were assembled without DNA template to measure background fluorescence levels. We performed five such reactions, to enhance the reliability of the results.
Experiment 3: Fluorescence Threshold Determination
RPA product was diluted to create a series of samples with the following concentrations: undiluted (1 ng/μL), 1:10, 1:100, 1:1’000, 1:10’000, and 1:100’000. These dilutions were then incorporated into standard Cas12a reaction mixtures.
Experiment 4: Free Fluorophore Calibration
Two Cas12a reactions were assembled to generate free fluorophores. One was treated with 2 μL of 0.5 M EDTA directly after the assembly. This was used as a negative control. Then both of them were incubated for 60 minutes at 37 °C. The EDTA was added in order to stop the enzyme activity. After the 60 minutes incubation time, we also added 2 μL of 0.5 M EDTA into the other previously untreated reaction. The reaction where we added the EDTA after the incubation was then used to create a dilution series for the calibration curve generation.
Experiment 5: Enzyme Denaturation
Aliquots of LbCas12a enzyme sufficient for four reactions were incubated at elevated temperatures (45 °C and 55 °C) for 10 minutes to induce partial denaturation. Control aliquots remained at standard temperature. All samples were then incorporated into LbCas12a assays.
Experiment 1:
Reactions with 4 μL and 8 μL of the ssDNA reporter were incubated at 37 °C. Fluorescence readings were taken at 5, 15, and 30 minutes to compare signal intensity and determine the impact of reporter concentration on the detection sensitivity.
Experiment 2:
The five no-DNA control reactions were incubated under standard conditions (37°C, 30 minutes), and fluorescence was measured to assess background signal levels.
Experiment 3:
All six dilutions of the RPA product were tested under standard reaction conditions (37 °C, fixed reaction volume). Fluorescence output was measured to determine the minimal DNA input needed to surpass background fluorescence, effectively establishing the system’s threshold.
Experiment 4:
The reaction treated with EDTA after the incubation (now containing free fluorophores) was serially diluted. The fluorescence was measured across the dilution series to generate a calibration curve, allowing conversion of raw fluorescence values into quantitative output. The other, undiluted reaction was measured as well, providing a reference point.
Experiment 5:
Denatured enzyme samples were used in reactions incubated at 37 °C for either 45 or 75 minutes. Control reactions with fresh enzymes followed the same protocol. Fluorescence measurements allowed comparison of enzyme activity post-heat treatment, providing insight into thermal stability and optimal handling conditions.
With these experiments, we were able to determine a range of parameters for our dry lab models.
Experiment 1:
In this experiment, our aim was to determine the kinetic parameters of the LbCas12a enzyme. However, due to a misunderstanding between the wet lab and dry lab teams, we only obtained data for two substrate concentrations, which is insufficient for a reliable parameter estimation. To overcome this limitation, we consulted a published study [4] that reports kinetic values for LbCas12a under various experimental conditions. From these, we selected the conditions most comparable to our own setup and used the corresponding parameters for our reaction model. Further details can be found on our Reaction Model page.
Experiment 2:
It was crucial for us to determine the background fluorescence in the absence of target DNA. This was the only way to accurately calculate the fluorescence generated by the reaction itself.
Experiment 3:
With the dilution experiment, we were able to determine the experimental threshold that must be exceeded to detect a signal. Based on this, our reaction model allowed us to establish the limit of detection (LOD), defined as the minimum number of DNA copies required to generate a detectable signal and thereby indicate the presence of an infection.
Experiment 4:
This experiment provided the necessary data to determine a linear calibration factor of 130.73 a.u./nM. Using this factor, we were able to calculate the concentration of cleaved reporter molecules from the measured fluorescence.
Experiment 5:
The Dry Lab has chosen a different method than initially intended to predict the test’s shelf life. Consequently, the results of this experiment were not used. For more details, see the Reaction Model page.