Engineering

Project Design:
Explore how PlasMission progressed through three major design plans and innovative ways to overcome technical barriers, initiate validation processes, and reinforce hypotheses. Witness how we internalize conjugation technologies into stopping beta-lactamase bacteria and shape the ultimate plan for conjugation and reliable validation of both successful conjugation and efficacy of BLIP.

Wet Lab:
Find out how PlasMission navigates through a total of four transformation steps in constructing a complete genetic circuit, each carefully designed to assemble functional modules with precision and compatibility. See how we employ the RP4 conjugation mechanism to facilitate horizontal gene transfer, enabling the donor bacteria carrying the BLIP gene to successfully deliver the target gene into recipient bacteria harboring ampicillin resistance. This process not only demonstrates controlled genetic exchange but also validates our conjugation system as a reliable platform for plasmid mobilization.

Dry Lab:
Simulate models of bacterial conjugation and the interaction between BLIP and β-lactamase. Our team built two computational models to understand how our BLIP systems fight antibiotic resistance. The first model focused on how plasmids spread between bacteria through conjugation, helping us estimate how efficiently the BLIP genes could transfer. The second model, created in COPASI, explored how BLIP interacts wit β-lactamase to block its activity and protect antibiotics. This allows us to compare the efficiency BLIP-I and BLIP-II degrading BLA. Together, these models helped us predict system performance even when lab experiments couldn’t be completed on time, giving us a clearer picture of how our experiment will and should work.

Human Practices:
Take a look at how PlasMission integrates synthetic biology, antibiotic resistance, and other applications into education and outreaches. A total of 6 interviews, 13 physical outreaches, and 3 education activities. In between each we make mistakes, reflect, and rebuild each interview and outreach methods.

Design
Explore how For BLIP expression and validation purposes, our insertion gene sequence is composed of RP4 oriT promoter, two RBS, BLIP-I/BLIP-II gene expression, fluorescent protein, and a terminator. We used the screen sequence function provided by Integrated DNA Technology (IDT) PlasMission progressed through three major design plans and innovative ways to overcome technical barriers, initiate validation processes, and reinforce hypotheses. Witness how we internalize conjugation technologies into stopping beta-lactamase bacteria and shape the ultimate plan for conjugation and reliable validation of both successful conjugation and efficacy of BLIP.

Build
Once everything was put together, the first version of codons had a complexity score of up to 78.1 for BLIP-I, and 87.22 for BLIP-II. We adjusted codons according to IDT’s optimizing guidelines. The first version of optimization was done with codon optimization tools provided by IDT, Benchling, and Snapgene.

Test
After optimizing all the lines that either had repeated sequences that were too long, or a too high number of GC content with codon optimizing tools listed above, the amino acids appeared to be different from the initial protein sequence. This process was repeated several times, all resulting in different amino acid sequences, different from our targeted sequence.

Learn
In this process we learned and adapted from only using codon optimization tools, to adjusting some parts of the gene sequence by manually. Although this process was more complicated and had a relatively higher error rate, after doing the same process of optimization for three times, we managed to get a result that was the same and authenticated.

Design
To validate the successful conjugation and transfer of the BLIP gene, we designed three complementary approaches. First, we used fluorescent markers, embedding green and red fluorescent proteins in the recipient and donor plasmids, respectively—successful conjugation would produce transconjugants expressing both colors. Second, we applied an antibiotic resistance assay, using selective plates to distinguish donors, recipients, and transconjugants based on their survival patterns. Third, we performed PCR amplification targeting BLIP-I and BLIP-II sequences to confirm their presence in transconjugant colonies. After further research, we refined our validation strategy by introducing optimized plasmid designs to enhance efficiency and reliability.

Build
Through this planning process, we identified potential challenges and limitations in our validation setup during later times when we received our distribution kit. There were no available pSB1K3 in the distribution kit, making it difficult for us to carry out the plan.

Test
Additionally, although the current validation method provides us with roughly estimated efficacy of BLIP, there is no direct proof of BLIP actually working against beta-lactam resistant bacteria.
Furthermore, the first validation method required a confocal for both colors to show at the same time, but we do not have access to it.

It was also predictable that the survival rate of the transconjugant bacteria to survive dual antibiotic plate might be too low for effective validation.

Learn
To conclude, this first validation would not be an ideal way to confirm the efficacy of BLIP and successful conjugation. The survival of the transconjugant on Kanamycin and Chloramphenicol plate is too unstable to provide reliable results and is unable to prove the functionality of BLIP.

Design
After the first cycle, we refined our system to directly demonstrate the efficacy of BLIP against ampicillin-resistant bacteria. Initially, we believed that working with ampicillin-resistant strains might not be permitted under the iGEM High School Safety White List, making validation challenging. However, after researching iGEM safety regulations and consulting biosafety experts, we confirmed that ampicillin resistance falls under β-lactamase resistance (Ambler Class A) — a category that is safe for BSL-1 high school teams.
This confirmation allowed us to proceed with a scientifically valid and safety-compliant design that effectively showcases BLIP's inhibitory potential.

Build
We established a new validation workflow to assess conjugation and inhibition more effectively:
- Continued using PCR and gel electrophoresis for molecular confirmation.
- Implemented cross-plating of donor and recipient cells on three antibiotic conditions: Chloramphenicol (Ch), Ampicillin (Amp), and Dual Ch + Amp. These conditions allowed us to compare survival patterns and evaluate BLIP’s inhibitory function.
- Introduced color-based differentiation for better visualization: Red colonies (mCherry) = donor strain, blue colonies (lacZ/X-gal reaction) = recipient strain, purple/mixed colonies = transconjugants carrying both markers. This combination of antibiotic selection and phenotypic color screening enabled precise differentiation among donor, recipient, and transconjugant populations.

Test
The cross-plating validation method successfully distinguished all cell types under selective conditions. Although IPTG did not arrive in time, we proceeded with blue-white screening using X-gal alone. Because the recipient plasmid (BBa_J435320) carries a leaky lac operon, some basal expression of lacZ occurs even without induction. As a result, the recipient and transconjugant colonies displayed a weak blue coloration, confirming that the color-based differentiation still functioned as expected.

For BLIP-II, no colonies appeared on the Amp plates, demonstrating complete inhibition of β-lactamase activity.
For BLIP-I, a few transconjugant colonies grew on Amp plates, indicating that inhibition was partially effective under the tested conditions.

Learn
Through this validation cycle, we learned the importance of experimental adaptability and quantitative refinement. Even without IPTG, the leaky expression of lacZ in BBa_J435320 was sufficient to visualize conjugation events, confirming that our screening system was robust and reliable.

The partial inhibition observed in BLIP-I transconjugants suggests that BLIP activity may vary depending on expression levels or antibiotic concentration. If additional time were available, we would perform dose–response assays using graded ampicillin concentrations to better characterize the efficacy range of BLIP-I. These insights will inform the next design cycle and help optimize BLIP constructs for more consistent β-lactamase inhibition in the future.

Design
We aimed to amplify the BLIP-I and BLIP-II gene sequences using PCR to generate sufficient DNA for cloning and downstream analysis. The goal was to obtain visible, high-quality DNA bands with sufficient concentration for ligation and digestion, while minimizing non-specific amplifications.

Build
To achieve this, we planned a 10 µL PCR reaction containing:
- 1 µL of BLIP template DNA (same for BLIP-I and BLIP-II)
- 0.5 µL each of VR and VF2 primers
- 3 µL of ddH₂O
- 5 µL of 2x Taq Master Mix

We referenced the experimental handbook provided by our collaborating team, NYCU Formosa, and online resources to design our PCR program:
- Initial denaturation: 95 °C for 5 min
- 25 cycles of:
→ Denaturation: 95 °C for 30 seconds
→ Annealing: 55 °C for 30 seconds
→ Extension: 72 °C (1 min per kb + 30 seconds)
→ Final extension: 72 °C for 10 min
→ Hold: 4 °C
Based on sequence lengths including mCherry, we set the expected extension time to 1 minute 30 seconds for BLIP-I (1,609 bp) and 2 min 30 s for BLIP-II (1,982 bp).

Test
In the first trial, we observed strong and bright DNA bands at the expected size but also faint trailing smears behind the main band.
This suggested potential primer dimerization or underextension, likely caused by an insufficient extension time. According to the procedure, the extension step for BLIP-I should be 2 minutes; however, we failed to account for the additional length of the mCherry sequence in our construct, leading to a miscalculation of the required extension duration.

Learn
We concluded that short extension time contributed to the background products. We modified the extension to 2:00 for BLIP-I and 2:30 for BLIP-II, which yielded cleaner bands suitable for downstream cloning.

Design
Use PCR to amplify BLIP-I and BLIP-II at sufficient yield and purity for downstream restriction digestion and cloning.

Build
Building on the previous cycle, where the PCR conditions were optimized, we aimed to further improve the yield and clarity of the BLIP-II amplification. After observing dim bands and low concentrations in several attempts, we modified the procedure while keeping the optimized temperature profile from DBTL cycle 1.

We maintained an initial denaturation at 95 °C for 5 minutes, followed by 35 total cycles (instead of 25) to increase DNA yield. Each cycle included denaturation at 95 °C for 30 seconds, annealing at 55 °C for 30 seconds, and extension at 72 °C for 2 minutes (BLIP-I) or 2 minutes 30 seconds (BLIP-II), ending with a final extension at 72 °C for 10 minutes and hold at 4 °C. The total reaction volume was increased to 20 µL to compensate for some DNA loss during the gel extraction.

Test
Restriction digests on post-cleanup PCR products were more reliable.
Correcting the extension time removed most primer-dimers/smearing (trailing bands).
Bands were brighter than earlier attempts, though still not as intense as the first two runs.

Learn
The earlier smearing/dimerization in cycle 1 was driven by extension time mis-setting. With the corrected time setting, the specificity has greatly improved. The persistently dim bands/low concentration despite many scaling up and improvements suggest DNA template degradation due to repeated freeze-thaw.

Lesson learned: We should always aliquot templates to avoid repeated freeze-thaw process and subsequent template degradation.

Overall, the revised program produced clean, digest-ready amplicons, enabling downstream cloning with higher consistency.

Design
To isolate pure fragments of BLIP-I, BLIP-II, and pSB1C3 using agarose gel electrophoresis while preventing too much DNA loss.

Build
We built the gel extraction protocol with the reference to the protocol from the NYCU lab manual. To purify DNA from the agarose gel, the desired bands are excised and transferred to labeled tubes. Add Buffer B after cutting the desired band on the gel, and incubate the mixture at 65 °C until the gel dissolves completely. The solution is transferred to a spin column, washed sequentially with Buffer W1 and Buffer W2, and centrifuged to remove impurities. The DNA is then eluted twice with 20 µL of ddH₂O, and the purified DNA concentration is measured before storage at -20 °C.

Test
After gel extraction, we can only evaluate whether we got our desired bands and if they have high enough concentration after restriction digestion and running gel electrophoresis.

09/23 Gel electrophoresis of BLIP-I, BLIP-II, and pSB1C3 restriction digestion

After several rounds of gel extraction and restriction digestion, the gels consistently showed faint or even non-visible DNA bands. We made multiple adjustments to reagent volumes and conditions, but none resulted in stronger bands. Eventually, after going over resources and the troubleshooting guide provided by GeneDireX, we discovered that Buffer W2 from the gel extraction kit had not been properly diluted with ethanol. This oversight likely caused inefficient DNA precipitation and binding to the silica column during the wash step, and therefore led to the low DNA recovery, as suggested by the weak or undetectable bands observed in our earlier attempts.

Learn
We invested a great deal of time and effort trying to identify the cause of our dim bands and low DNA concentrations. Overlooking the need to dilute Buffer W2 with ethanol significantly impacted our experimental progress, leaving us with limited time for subsequent adjustments and trials. This experience taught us an important lesson, always double-check the instructions for kits and reagents before starting any experiment to avoid losing valuable time over an easily preventable mistake.

Design
Aims to isolate pure fragments of BLIP-I, BLIP-II, and pSB1C3 using agarose gel electrophoresis while preventing too much DNA loss.

Build
After figuring out we should dilute our Buffer W2 with ethanol beforehand, we used the same protocol from the NYCU experiment notebook.

Test
To our surprise, the DNA concentration remained low even after properly diluting Buffer W2 with ethanol. While this adjustment slightly improved the results, it became clear that another underlying factor was still causing the dim or non-visible bands observed after gel extraction. After researching and brainstorming over potential reasons, we suspected that pH level of ddH₂O may be ideal, as the GeneDireX manual explicitly stated the ideal pH range to be 7.0-8.5. After measuring the pH of our ddH₂O, we found that the pH was around 5-6, which could be the major reason for low elution strength. To ensure consistency, we modified the procedure by replacing ddH₂O with Tris Buffer, as suggested by the manual.

Learn
Through this process, we learned that even small details in reagent choice can significantly affect DNA recovery efficiency. Replacing ddH₂O with Tris buffer greatly improved the elution, as Tris helps maintain a stable pH that prevents DNA from adhering too tightly to the column membrane and protects it from acid-induced degradation. This experience emphasized the importance of understanding the chemical mechanisms behind each step rather than following protocols mechanically, allowing us to troubleshoot more effectively and improve the reliability of our gel extraction results.

Design
Aims to isolate pure fragments of BLIP-I, BLIP-II, and pSB1C3 using agarose gel electrophoresis while preventing too much DNA loss.

Build
Following the protocol provided in the Gene DireX Inc. gel extraction kit, we purify DNA from the gel. 500 µL of Buffer B is added to the gel slice and incubated at 65 °C until fully dissolved. The solution is transferred to a spin column, washed with Buffer W1 and Buffer W2, and centrifuged to remove impurities. To make the DNA more concentrated, the DNA is eluted with 25 µL instead of 50µL of Tris Buffer (instead of ddH20 due to acidity), and stored at -20 °C.

Test

Restriction digestion results using the DNA samples purified by the modified protocol. The bands are visible and clean.

The purification went well and we purified our desired bands. After restriction digestion, we see bands indicating successful gel extraction. Proving that the acidic ddH2O is the reason for past failed attempts, and replacing it with Tris buffer works more effectively, we achieved clearer digested bands and substantially higher DNA yield.

Learn
We learned that the acidic ddH₂O used during gel purification was the primary cause of low elution efficiency and reduced DNA concentration. By switching to Tris buffer for elution, we significantly improved DNA recovery and stability, ensuring sufficient yield for downstream reactions.

Using the purified PCR products from this optimized workflow, we proceeded with restriction digestion, ligation, and transformation, all of which were successful. This marked a major engineering milestone in our cloning workflow, demonstrating that our systematic optimization effectively resolved previous bottlenecks and enabled reliable plasmid construction.

Design
After ligation, our recombinant plasmid is ready to be introduced into the bacteria via transformation. To make our donor and recipient ready to take in the recombinant plasmid, we made competent cells for S17-1 and DH5α.

Build
Taking the protocol from the RBC HIT Competent Cell Protocol Book. To prepare competent E. coli cells, 0.6 mL of overnight culture was added to 12 mL of LB broth and incubated at 37 °C with shaking for about 1 hour until the OD₆₀₀ reached 0.2-0.3. The culture was chilled on ice, centrifuged, and washed twice with ice-cold 10% CaCl₂ to make the cells competent. Resuspend the final pellet in a small volume of 10% CaCl₂ for immediate heat-shock transformation for the most effective approach, or in 15% glycerol + CaCl₂ solution for long-term storage at –80 °C.

Test
After incubating the overnight culture, added to 12 mL of LB Broth, and incubate in 37 °C with shaking for 1 hour, we got OD₆₀₀ = 0.275, which is within the desirable range of 0.2~0.3 OD. However, only a very tiny and almost non-visible pellet is observed after the first spin down. We further centrifuge it a few more times in order to get bigger pellets, but the pellets were still extremely small. As a result, the procedure was halted and we had to redo a new batch.

Learn
From the tiny pellets observed in our first competent cell preparation attempt, we realized the failure was likely due to low cell/DNA concentration. In addition, repeatedly centrifuging the culture may have further reduced cell potency. This experience taught us that both incubation timing and protocol accuracy are critical. To improve, we planned to incubate the overnight culture longer to achieve a higher OD and greater DNA concentration, and we also modified the procedure accordingly.

Design
Aimed to prepare E. coli S17-1 with higher OD values to ensure sufficient cell density and improve competency. By optimizing the preparation conditions, we intended to maximize transformation efficiency and establish both strains for use in later conjugation experiments.

Build
We took the protocol from the RBC HIT Competent Cell Protocol Book. To prepare competent E. coli cells, 0.6 mL of overnight culture was added to 12 mL of LB broth and incubated at 37 °C with shaking for about 3-4 hours until the OD₆₀₀ reached 0.4-0.6. The culture was chilled on ice, centrifuged, and washed twice with ice-cold 10% CaCl₂ to make the cells competent. Resuspend the final pellet in a small volume of 10% CaCl₂ for immediate heat-shock transformation for the most effective approach, or in 15% glycerol + CaCl₂ solution for long-term storage at -80 °C.

Test
After incubating the overnight culture for more than 4 hours, we measured an OD₆₀₀ of 0.8, indicating high cell density. Unlike our first attempt, a visible pellet was obtained after the initial spin down, showing clear improvement. However, we could not proceed directly to heat-shock transformation since no plasmid was yet fully prepared for transformation. As a result, we stored 10 tubes as glycerol stocks at –80 °C. Unfortunately, after being stored for several weeks, the S17-1 glycerol stocks lost competency, and subsequent transformation attempts were unsuccessful.

Learn
From the loss of competency in our glycerol stock, we realized that S17-1 cells are highly sensitive to long-term storage and quickly lose their ability to take up DNA. This taught us the importance of using freshly prepared competent cells rather than relying on frozen stocks for transformations. We also learned that timing the preparation of competent cells to align with plasmid availability is critical to avoid unnecessary storage and reduced efficiency.

Design
For our third cycle, we aimed to prepare E. coli S17-1 cells with a high OD value and ensure they were freshly made chemically competent. Unlike our previous attempts, where glycerol stocks lost competency or the pellets were too small, this time we planned to use the cells immediately after preparation. This approach was designed to maximize the likelihood that the donor strain would successfully take up the plasmid.

Build
We followed the same protocol as in our second cycle, but with one key change: instead of making glycerol stock of the competent cells, we used them immediately after preparation for transformation.

Test
After incubating the overnight culture for 4 hours, we measured an OD₆₀₀ of 0.392 (~0.4), and a clear, visible pellet was obtained after the initial spin down. We then directly performed heat-shock transformation using the plasmid we had miniprepped earlier. Following the transformation, clear colonies were observed, confirming that our approach with freshly made competent cells was successful.

Learn
We learned that the freshness of competent cells is crucial for transformation success. Using freshly prepared S17-1 cells significantly improved our results, highlighting how subtle differences in preparation can directly determine experimental outcomes.

Design
The initial design of our conjugation model was to input quantitative data from wet lab experiments directly into our mathematical equations framework in order to produce realistic simulations and visualizations of bacterial conjugation.

Build
To model the dynamics of donors, recipients, and transconjugants, we applied mathematical formulas that describe recipient and transconjugants in an inversely proportional manner. These formulas capture the essential behavior of conjugation: as transconjugants increase, recipients decrease correspondingly, while donors provide a stable source of plasmids.

Test
During finalization, we found that assuming donors remain constant did not fully capture real-life dynamics, as donors actually grow over time. Additionally, due to temporal constraints in the experimental flow, the wet lab was unable to provide the necessary data within our modeling timeline.

Learn
Although real-life experimental data from the wet lab were unavailable within the project timeline, our dry lab team took the initiative to research existing literature and derive our own parameter values. This approach allowed us to refine our conjugation model despite the lack of direct measurements.
Moreover, computational modeling enabled us to simulate scenarios that the wet lab could not feasibly achieve, providing a broader platform for exploring plasmid transfer dynamics.

Design
We developed a conjugation model that simulates both donor growth and no-donor-growth scenarios. The no-donor-growth version provides a clear view of the inverse relationship between transconjugants and recipients, isolating the direct effect of conjugation without the added complexity of donor proliferation. Because we operate as a dry lab, we are able to construct and analyze this simplified model—something that would be difficult or impossible to replicate in the wet lab—thereby offering unique insights into plasmid transfer dynamics under controlled conditions.

Build
We proceeded to construct the model using parameter values derived from accepted experimental ranges: donor and recipient populations of 1.0×10⁶ and 1.0×10⁷ cells/mL, respectively, reflecting the common 1:10 ratio used in conjugation experiments. This enables us to maintain progress despite the absence of primary data.

Test
The values were implemented into our equations to model how the donor, recipient, and transconjugant populations change over time. This helped us ensure that our model behaves in a relatively realistic way and makes biological sense under the assumptions we set. After inputting these values, we generated graphs to provide clearer visualization of the population dynamics and make the results more intuitive to interpret.

Learn
Simulation results show that in the no-growth model, donors remain constant, serving as a stable plasmid source and making the inverse relationship between recipients and transconjugants clearly visible. When donor and recipient growth are included, cell numbers increase exponentially, amplifying plasmid transfer due to the expanding donor population. By comparing these two scenarios, our dry lab simulation achieved something the wet lab cannot easily replicate—modeling and visualizing plasmid transfer under controlled, hypothetical conditions—highlighting both the direct impact of conjugation and the amplification effect of cell growth on resistance gene spread.

Design
We simulate a model in COPASI to explore the interaction between BLA and its inhibitors-BLIP-I and BLIP-II. This design step defines the key parameters and reaction networks needed to compare how each BLIP variant affects Beta-Lactamase (BLA) expression and activity.

Build
We build the model by implementing the defined reactions and expression profiles of BLA, Ampicillin (Amp), BLIP-I, and BLIP-II into COPASI. This includes specifying rate laws, initial concentrations, and any relevant interactions to ensure that the model represents the biological system.

Test
We test the model by running multiple simulations to evaluate BLA dynamics under the influence of each BLIP variant. These simulations generate time-course data, allowing us to compare the inhibitory efficiency of BLIP-I and BLIP-II in modulating BLA activity.

Learn
We learn from the analysis of the simulation outputs and graphs, which show that BLIP-II achieves more effective inhibition of BLA than BLIP-I. These insights inform future model refinements and experimental strategies for validating BLIP-mediated control of BLA.

Design
At the beginning of our Human Practices work, our goal was to gather expert insights on antibiotic resistance, gene transfer safety, and the ethical implications of conjugation-based systems. We designed interview questions intended to guide our project direction and validate our ideas. However, our early drafts were too broad or unrelated, making it difficult for interviewees to provide specific or actionable feedback.

Build
We conducted several initial interviews with professors and field experts, using the original question sets. During these sessions, we realized that some questions lacked focus or were too general to yield meaningful responses. To improve, we began collaborating internally across subteams, especially between the Wet Lab, Dry Lab, and HP members. This was to ensure that future interview questions were directly connected to our project's experimental design and goals.

We also established a post-interview reflection process, where team members would discuss what worked well and what could be improved for future stakeholder engagements. In addition to that, we wanted to see our impact in our outreach community, thus we held interviews with in-school students and the head principal.

Test
In subsequent interviews, the revised question sets produced more insightful and relevant feedback. Interviewees were able to engage in deeper discussion about our project's biosafety considerations, educational value, and real-world applications. These focused interviews provided information that directly informed our project implementation and wiki documentation.

The addition of post-meeting team discussions helped us identify key takeaways more clearly and brainstorm how each piece of feedback could be integrated into our design and outreach strategy.

Learn
We learned that Human Practices requires the same iterative refinement as wet-lab experiments — careful planning, testing, and reflection lead to better outcomes.
Through this cycle, we discovered that:
- Collaborative question design with relevant subteams produces more meaningful and technically grounded discussions.
- Focused, specific questions yield deeper expert insights compared to general or abstract prompts.
- Conducting post-interview debriefs ensures that valuable feedback is not only documented but also actively integrated into project development.
This iteration marked a significant improvement in how we approach stakeholder engagement, transforming our interviews from simple Q&A sessions into productive dialogues that shaped our project's ethical and scientific direction.

Design
In the initial phase, our objective was to investigate the relationship between early childhood creativity and their understanding of antimicrobial resistance (AMR). We hypothesized that introducing AMR through interactive storytelling and games would allow young children to better grasp hygiene practices and the dangers of antibiotic misuse. This was grounded in the idea that instilling basic awareness at an early age could foster responsible habits and future scientific curiosity.

Build
In order to construct this outreach initiative, we designed a series of hands-on learning stations tailored to children of ages 4~5. This included reading a hand-illustrated storybook titled, “Lil Joey and the Jungle’s Sick Story” which introduced the concept of AMR and how we can prevent them. Other activities that promoted young creativity in the field of synthetic biology include decorative games like “Build-A-Bug-Biolab” that allowed kids to showcase their interests in editing their own bacterias to make changes to the fields of biology. In addition to that, our team prepared materials that can be further used in the future for children of all ages, this includes our coloring book, educational videos, and more. This program was crafted to appeal to children’s senses of imagination while simplifying scientific principles for their age level.

Test
The two-day program was implemented at Star Academy in Hsinchu, successfully engaging over 130 kindergarten students across multiple rotating activity stations. The children demonstrated active participation during storytelling sessions by frequently raising their hands to ask questions and volunteering their ideas on preventing the spread of germs and infections. They also made insightful comments about medication misuse when prompted during the story of “Lil Joey and the Jungle’s Sick Story”. In the Build-A-Bug Biolab station, students creatively designed “problem-solving bacteria,” reflecting their ability to grasp abstract biological concepts through play. Many children drew bacteria with “superpowers” like fighting sickness, eating bad germs, or healing wounds, early evidence of their developing understanding of microbial functions. Teachers and staff noted the high level of excitement and retention, and several expressed interest in incorporating similar modules into their own teaching. This strong engagement validated our approach and confirmed that early childhood education can lay foundational knowledge for topics like AMR and responsible health behavior.

Learn
After our outreach at Star Academy, we realized that for younger audiences, interactive movement-based learning and simplified analogies significantly improved engagement and understanding. While our original plan already included visuals and stories, seeing how quickly students responded to physical games like the “No Germs Alive” activity helped us shape our future outreach planning by encouraging us to incorporate more interactive elements and clear debrief moments, even for older age groups, to ensure the understanding of AMR concepts in both the medical and environmental field through multi-sensory learning.

Design
In this cycle, our objective was to explore the relationship between community awareness and attitudes toward antibiotic use in vulnerable populations. We hypothesized that elderly patients, with proper guidance, would adopt more responsible behaviors to reduce AMR risks. This was grounded in the belief that increasing public literacy could reduce misconceptions and improve health outcomes.

Build
Firstly, we identified the primary aim of our outreach and constructed our plan around it. We carefully designed brochures that clearly explained key elements: the beginning of our journey and our goals, the purpose of our outreach, the impact and harm of the issue we are addressing, and our previous collaboration with Ton Yen Hospital. Once the brochures were completed, our Human Practices team carefully planned the outreach schedule and determined how best to approach the audience. To ensure the effectiveness of the outreach, we collaborated with Ton Yen Hospital once again and developed detailed interactive questions about the dangers of AMR. In addition, we prepared small rewards, provided through our collaboration with the Hsinchu Health Bureau, for participants who answered the questions and demonstrated their support.

Test
To assess the effectiveness of our outreach strategy at Ton Yen Hospital, we directly engaged with the local elderly population. During our information booth sessions, we distributed brochures that contained information about the project and constructed interactive Q&A sessions. The questions, which were based on common misconceptions about antibiotic use allowed the elderly community to actively participate and interact with the team. We asked a total of 5 questions where people voluntarily raised their hands to answer questions.

Learn
Through this activity, we learned that antimicrobial resistance still remains unfamiliar to older age groups, with many people having misconceptions regarding prescription and taking antibiotics. We also acquired the information that iGEM isn’t as well known for Taiwanese citizens, and the synthetic biology field still remains more foreign.

Design
In this cycle, our objective was to examine the relationship between student career aspirations and their engagement with AMR education. We hypothesized that linking AMR prevention to students’ future goals in medicine or biology would increase both interest and responsibility toward scientific issues. This was based on the notion that contextualizing synthetic biology within real career pathways enhances relevance and motivation.

Build
We initiated the process by designing a lesson plan to map out the slideshow we'd use during the showcase. The presentation included an opening question, relations to the SDGs, brief overview of the issue of antimicrobial resistance, our team, interactive questions, and an exit ticket. Our questions were mainly based on the feedback we’ve acquired from previous interviews and past teams’ works. Not only do we intend to observe the relationship between student career aspirations with AMR education, it is also meant to align with the Sustainable Development Goals (SDGs) to show how our project shows mindful connection.

Test
To assess the impact and actual implementation of our experiment, we consulted other advisors beforehand about whether they think the lesson will be comprehensible for students. We also asked if there could be connections to broader and more complex topics, as well as the implementation of our event into the curriculum.

Learn
They advised us to include more visuals such as videos to draw people's interest, enhance comprehension, and keep them attentive, which we then used to amend our slides accordingly. From this event, we learned that for our outreach purposes many students got inspired by our events and have an interest in pursuing a career in similar fields.