Design:

Our Design began with the decision to explore antimicrobial peptides (AMPs) for agricultural applications. However, we wanted to test a peptide that had not yet been applied against citrus pathogens. This led us to Professor Eduardo Festozo page from Universidade Estadual Paulista (UNESP), who has studied the peptide CTX for over ten years in his laboratory. This peptide was isolated from Hypsiboas albopunctatus, a small frog native to the Brazilian Cerrado. It consists of 21 amino acids and acts through the “barrel-stave” mechanism , inserting into microbial cytoplasmic membranes, forming pores that destabilize the membrane and ultimately lead to cell death. This property prevents the development of microbial resistance, highlighting CTX as a promising alternative to conventional antibiotics. Professor Festozo provided us with a 2 mg chemically synthesized sample of CTX, enabling us to begin our first tests.

We designed an assay to evaluate CTX's effectiveness against major post-harvest fungal pathogens— Penicillium digitatum and Geotrichum candidum. The protocol was based on a small-scale plate assay to quickly visualize antimicrobial activity. We selected concentrations— 10 µM, 25 µM, and 50 µM—to determine the minimum inhibitory concentration (MIC). This setup allowed rapid, low-volume screening while providing clear inhibition patterns.

Build:

To carry out the first tests with CTX, we collaborated with Professor Taicia, an expert in citrus pathogens. She granted us access to her laboratory facilities, where experiments were conducted under the supervision of her graduate students and specialists in plant pathology. This ensured biosafety compliance and increased the reliability of our assays. Fungal strains of Penicillium digitatum and Geotrichum candidum were prepared from her group's collection, providing well-characterized and reproducible models for post-harvest citrus diseases. With access to controlled growth conditions, sterile media preparation, and experienced technical guidance, we built a robust experimental framework that allowed us to test the peptide with confidence and generate reproducible results.

Test:

The assays were performed in 96-well plates (200 µL per well), where each well was inoculated with approximately 10⁵ spores of the target fungus. CTX solutions were applied at the concentrations defined during the design stage (10 µM, 25 µM, and 50 µM). Plates were incubated at 25 °C, without a photoperiod, for 48 hours. Fungal growth was monitored visually, and inhibition was assessed by comparing colony development across treatments. A commercial fungicide (Imazalil) was included as a positive control, while untreated wells served as negative controls. This setup allowed us to confirm CTX's effectiveness and to identify the concentration at which the strongest inhibition occurred.

Learn:

From these tests, we observed that CTX was effective against both pathogens. It showed complete inhibition of Penicillium digitatum at 25 µM and Geotrichum candidum at 50 µM after 72 hours. After 120 hours, 50 µM was sufficient to inhibit both fungi . These MIC results provide a solid foundation for future dosage modeling in field applications. Overall, our findings validate CTX as a promising antimicrobial candidate and guide our next step: testing CTX against Huanglongbing (HLB, "Greening"), the most devastating citrus disease worldwide.

Greening is currently the most devastating citrus disease worldwide. We recognized the importance of attempting to provide a solution to this challenge. However, working with the causal bacterium, Candidatus Liberibacter asiaticus, is extremely difficult since it is non-cultivable in laboratory conditions. Direct plant assays are also complex, time-consuming, and require large amounts of peptide. To overcome these obstacles, we partnered with the Centro de Citricultura de Cordeirópolis (IAC) to design an indirect but rapid test using infected citrus leaves.

The experimental design involved collecting leaves from infected trees and excising approximately 15 mg of tissue from the petiole tip for DNA extraction (baseline sample). The remaining leaf tissue was placed in a 50 mL Falcon tube containing 2 mL of treatment solution and incubated at room temperature for 7 days. After incubation, the same amount of tissue (15 mg) was excised again from the petiole, and DNA was extracted. Both DNA samples (before and after treatment) were sent to a specialized diagnostic clinic for qPCR quantification of Candidatus Liberibacter asiaticus, allowing us to compare bacterial loads and evaluate treatment effectiveness. The DBTL framework was fundamental for the development of this strategy, since we needed to run three internal DBTL cycles to fully verify the feasibility and robustness of the assay, as follows:

1st DBTL cycle: Would Liberibacter survive in excised leaf tissue for the duration of the experiment?

To address this question, we set up a pilot assay in which infected leaves were incubated for 7 days in Falcon tubes containing either water or culture medium. DNA was extracted before and after incubation and analyzed by qPCR. The results confirmed that the bacterium remained detectable after 7 days, with only a slight reduction in concentration. From this cycle, we learned that Liberibacter was able to persist in excised leaf tissue for the entire incubation period, providing the foundation to proceed to the next DBTL cycles.

2rd DBTL cycle – Was CTX effective against Candidatus Liberibacter (CLas)?

With the assay validated in the previous cycle, we proceeded to evaluate whether CTX could reduce the bacterial load in infected citrus leaves. For this purpose, excised leaves were incubated for 3 days in Falcon tubes containing CTX solutions at the concentration previously defined during the design stage (100 mM). As a positive control, Oxytetracycline at 100 µg/mL was used, since this antibiotic has been reported in the literature as effective in reducing bacterial load in Greening-infected plants. After incubation, DNA was extracted before and after the treatments. A conventional PCR was initially performed to confirm the presence of Candidatus Liberibacter asiaticus (CLas), and all samples tested positive. Bacterial quantification was carried out by qPCR; however, due to a delay in result delivery, it was not yet possible to determine whether there was a reduction in bacterial population based on Ct values.

Bacterial quantification was carried out by qPCR. This experiment aimed to directly evaluate the potential of CTX as an antimicrobial agent against the citrus greening pathogen under controlled conditions. For this purpose, eight samples were analyzed, separated according to treatments, and collected before and after the experimental period. The results indicated that there was no significant reduction in the bacterial population in the samples after treatment (table). However, this lack of reduction was also observed in the positive control, which should have shown a decrease, suggesting that the exposure time used in the experiment may have been insufficient. Therefore, we suggest that in future assays the treatment period be extended in order to more accurately assess the efficiency of the procedure. In addition to this modification, we also considered the application of CTX directly to induced roots of cuttings derived from Greening-infected plants, since the root system has greater efficiency in absorbing molecules than the petiole and roots obtained from cuttings would present a more homogeneous bacterial load. This approach has not yet been carried out due to the time required for root induction, but it represents a promising alternative to test the effectiveness of the peptide.

Our work aimed at producing CTX in microbial cell factories, but we understood the inherent challenge of expressing AMPs in microorganisms due to their potential toxicity. To develop a viable strategy, our team selected three initial hosts: Escherichia coli, Saccharomyces cerevisiae, and Aspergillus oryzae. Each offered distinct advantages and productive potential. E. coli could be engineered rapidly, providing a fast system for initial validations. S. cerevisiae represented a more robust platform, capable of growing to high cell densities in large-scale fermenters, similar to those already used for bioethanol production in well-established industrial processes. Finally, A. oryzae, a filamentous fungus, was chosen for its reputation as a powerful protein biofactory with a strong natural capacity for secretion. Because AMPs often harm the very hosts intended to produce them, it was critical to evaluate CTX's toxicity against these expression systems. This step was essential not only for anticipating production challenges but also for shaping the engineering strategy we developed to safely express CTX in microbial platforms.

Design:

We designed a growth inhibition assay to assess whether CTX had toxic effects on each host organism. The experimental design mirrored the logic used for pathogen testing, with different peptide concentrations. The goal was to determine the minimum inhibitory concentration (MIC) that impacted host growth and to compare susceptibility across species.

Build:

At our PI's laboratory, Prof. André Damasio (LEBIMO), we obtained the three expression hosts (E. coli, S. cerevisiae, and A. oryzae) as well as the equipment required for the experiments. The CTX peptide used in the assays was provided by Prof. Eduardo Festozo, as described above.

Test:

The assays were performed in 96-well plates (200 µL per well). Each well was inoculated with approximately 0.1 OD of E. coli or S. cerevisiae, and with 5 uL of 10⁶ spores of A. oryzae. CTX solutions were applied at the concentrations defined during the design stage. Plates were incubated under optimal conditions for each organism (E. coli at 37 °C and S. cerevisiae/A. oryzae at 30 °C). Microbial growth was monitored visually, and inhibition was assessed by comparing biomass development across treatments.

Learn:

The tests showed that CTX strongly inhibited the growth of E. coli and Aspergillus oryzae, Saccharomyces cerevisiae displayed higher tolerance at the tested concentrations, although its growth was still slightly affected. These results highlighted the need for a protective expression strategy, which guided us to design CTX as a fusion with highly expressed carrier proteins—ensuring safe production in the host, followed by controlled cleavage to release the active peptide.

After verifying the toxicity of the peptide against the selected hosts, our team developed a strategy to enable CTX production. Since its mechanism of action targets cellular membranes through peptide interactions in solution ^[1], we hypothesized that coupling CTX to a carrier protein could inactivate its antimicrobial activity and allow safe expression inside microorganisms. To test this hypothesis, we selected sfGFP as the carrier protein because it is highly soluble, easy to detect, and well characterized as a reporter. We decided to begin by expressing the sfGFP–CTX construct in E. coli and Aspergillus oryzae to validate this approach.

Design:

We designed a fusion construct to protect CTX during expression and enable clean post-processing:

We selected sfGFP BBa_I746916 as the carrier protein because it is highly soluble, easily detectable, and well characterized as a reporter. The K7 construct included a flexible 3×GS linker (BBa_J18921) to provide spatial separation from sfGFP; dual TEV protease sites (BBa_K20750037) flanking the CTX to enable precise release; and an 8×His tag (BBa_K4422007) to allow purification by Immobilized Metal Affinity Chromatography (IMAC). After purification, TEV cleavage yielded a small, tag-free CTX ready for downstream use. Considering CTX’s small size (~2.3 kDa), we planned its separation from the larger carrier fragments (~30 kDa) via ultrafiltration using a 10 kDa MWCO membrane, allowing CTX to pass through while retaining the carrier (size-based cleanup).

Build:

We decided to express sfGFP–CTX in two distinct hosts— Escherichia coli (bacterial system) and Aspergillus oryzae (filamentous fungus). These complementary expression platforms allowed us to compare production performance and assess whether the construct would inhibit host growth, informing both scalability and biosafety for downstream manufacturing.

For the E. coli system:

To construct the expression vector for CTX, we designed a fusion strategy with sfGFP, enabling easy detection of protein production during in vitro assays. The DNA fragments required for this construction were provided by FASTBio, including the HIS–TEV–AMP cassette, while the expression backbone pJL1–sfGFP was kindly supplied by BioLinker®. The pJL1–sfGFP plasmid provided by BioLinker was first solubilized following standard procedures. The plasmid pellet was resuspended in 50 µL of elution buffer after centrifugation (1 min at 13,000 rpm) and gently mixed. DNA concentration was measured by Nanodrop, yielding a concentration of 50 ng/µ and transformed it into E. coli DH5α by heat-shock. Transformation was carried out with 1 µL of solubilized DNA, followed by recovery in LB medium. Transformed cells were plated on LB agar supplemented with kanamycin. Two plating conditions were tested: 10 µL of recovered culture and the remaining volume spread across separate plates. Plates were incubated at 37 °C for 15 hours, resulting in colonies carrying the backbone plasmid.

Two colonies were selected from the plate inoculated with 10 µL of transformed E. coli. Each colony was inoculated into a 50 mL Falcon tube containing 3 mL of LB medium supplemented with 3 µL of kanamycin. Cultures were incubated at 37 °C and 180 rpm for 16 hours in a shaker. Plasmid DNA was then extracted and purified using the Monarch Miniprep Kit, following the manufacturer’s protocol. Quantification by Nanodrop confirmed high yields of plasmid DNA:

• pJL1–sfGFP C1: 221.06 ng/µL
• pJL1–sfGFP C2: 292.66 ng/µL

For the construction of pIGEM001 – sfGFP–CTX, the pJL1–sfGFP vector was divided into two fragments to facilitate amplification. In parallel, the cassette K7–CTX insert (149 bp), provided by FastBio©, was amplified from pUC K7CTX. All fragments were subsequently purified and assembled using the NEBuilder DNA Assembly Kit, following the manufacturer’s protocol.

Putative colonies were screened by colony PCR and the correctly assembled clones produced a 1,074 bp product. Two correct colonies C4 and C6 were miniprepped and validated by XbaI restriction digest, which generated the expected 2.7 kb fragment. These colonies were stored in the freezer - 80°C using glycerol 30%.

for the A. oryzae expression system:

We amplified the sfGFP–CTX fragment (~1000 bp) from our constructed plasmid (pIGEM001) by PCR using primers containing homology regions for insertion into pUC57-IS1 (~5000 bp). The pUC57-IS1 plasmid belongs to the DIVERSIFY system, which allows the integration of a gene of interest (GOI) into the IS1 region, thereby disrupting the uidA gene. We linearized the pUC57 plasmid using primers with homology to our K7.

• pUC57 empty: 13 ng/µL
• GFP–CTX insert: 72 ng/µL

These purified fragments were then used for the NEBuilder® DNA assembly reaction using 3:1 concentrations by Ligation Calculator.

The NEBuilder® assembly reaction was treated with 1 µL of DpnI at 37 °C for 1 hour to remove the template plasmids. Following digestion, the entire assembly mixture was transformed into chemically competent E. coli cells. Transformants were plated on LB agar supplemented with ampicillin and incubated at 37 °C overnight.

The transformation produced numerous colonies, including several satellite colonies. The 12 largest colonies were selected for screening. The Colony PCR (cPCR) screening was performed using the primers, which yield a 1200 bp product for positive clones (successful integration) and a 2200 bp product for negative clones (empty vector), using NEXT TUPÃ Polymerase.

From the colony PCR, candidates 8 and 12 appeared to contain the correct construct. Both were selected for Miniprep plasmid extraction and quantified by Nanodrop, yielding:

• C8: 50 ng/µL
• C12: 30 ng/µL

To perform a second round of confirmation, the plasmids were digested with BamHI restriction enzyme. This analysis was designed to verify the presence and integrity of the sfGFP–CTX construct.

~200 ng of each plasmid was digested with 1 uL BamHI FastDigest in the FDgreen buffer by 1 hour in 37°C.

Aspergillus oryzae Transformation with pIGEM002 (C12):

For the transformation of A. oryzae strain Ory7, we used the confirmed construct pIGEM002 (C12) obtained in the previous assembly. The transformation protocol used in this work was the same from the DIVERSIFY system.

Buffers:

• Aspergillus Transformation Buffer (ATB): 1.2 M sorbitol, 50 mM CaCl₂, 20 mM Tris, 0.6 M KCl
• Aspergillus Protoplastization Buffer (APB buffer): 1.1 M MgSO₄, 1 M Na₂HPO₄, 1 M NaH₂PO₄, pH 5.8
• PCT (PEG–Calcium–Tris): 50% (w/v) PEG 8000, 50 mM CaCl₂, 20 mM Tris, 0.6 M KCl.

The first transformation step is protoplasting. The spores were harvested directly from agar plates using 20 mL of MMG/YPD and inoculated into a 250 mL Erlenmeyer flask containing 100 mL of medium, incubated at 30 °C, 180 rpm for ~15 hours. After sufficient biomass was obtained, the culture was filtered through Miracloth, and the resulting mycelial “cake” was collected. The mycelial cake was washed with ~20 mL of APB buffer and transferred to a 50 mL Falcon tube with 20 mL of APB containing lysozyme (VINOTASTE) at 40 mg/mL (800 mg total). The mixture was incubated at 30 °C, 150 rpm for 2–3 hours. Protoplast formation was checked under the microscope after ~1 h 45 min, and again after 3 hours, confirming successful protoplasting.

After enzymatic digestion, the culture medium was filtered through Miracloth, and the flow-through was adjusted to a final volume of 40 mL with ATB buffer. To this suspension, 5 mL of diluted ½ ATB was slowly added, generating a two-phase system. The mixture was centrifuged at 3000 × g (acc 9, dec 4) for 12 minutes, resulting in two visible phases. The protoplasts were located in the upper phase, which was carefully collected (~5 mL) using a micropipette and transferred to a fresh 50 mL Falcon tube. The suspension was then brought up to 40 mL with ATB and centrifuged again at 3000 × g (acc 9, dec 9) for 12 minutes, producing a protoplast pellet. The supernatant was discarded, and the pellet was gently resuspended in 1 mL of ATB. Protoplast concentration was checked using a Neubauer chamber, and counts above 10⁶ protoplasts/mL were recommended for transformation. Aliquots of 100 µL were prepared and stored in 0.6 mL microtubes for immediate use.

The pIGEM002 plasmid was digested with SwaI at 25 °C for 1 hour to release the donor DNA fragment (~3.5 kbp) to facilitate homologous recombination in A. oryzae.

Transformation plates:

For the transformation, 150 µL of protoplast suspension (~1 × 10⁶ protoplasts/mL) was thawed directly from −80 °C storage. The transformation mix included 3–10 µg of total DNA (donor DNA and CRISPR guide plasmid), corresponding to 25–50% of the total reaction volume. To this suspension, 200 µL of cold PCT buffer (4 °C) was carefully added. Since protoplasts are highly sensitive, the buffer was added slowly along the tube wall without pipette mixing. The mixture was incubated for 10 minutes at room temperature. Next, 250 µL of ATB buffer was gently added using the same approach, avoiding direct mixing to preserve protoplast integrity. The transformation mixture was then spread onto a selective regeneration medium supplemented with 1.2 M sorbitol and incubated at 30 °C to allow colony development.

The X-Gluc plate with 6 candidates from transformation plates T1 and T2 and Ory7 Wild type in the center. (+) are protoplast cultivation without selection (with Uracil/Uridine) and (-) are protoplast with selection media (without Uracil/Uridine).

For the first screening step, spores from the three candidate (1, 2 and 3) transformants were inoculated into 10 mL of MMG medium (1% + 2% maltose) and incubated for 72 hours. This cultivation allowed us to obtain both secretome for protein analysis and biomass for DNA extraction. Approximately 10 µg of total protein from the culture supernatant was loaded onto a 12% SDS-PAGE gel to check for expression of the fusion protein sfGFP–CTX (expected size: 35 kDa).

The Biomass was harvested from static cultures after 48 hours in 10 mL dish plates. Around 100 mg of biomass was collected and used for DNA extraction, while the remaining culture medium was saved for SDS-PAGE protein screening. The biomass was ground into a fine powder (cotton stick with liquid nitrogen) and resuspended in 500 µL of DNA extraction buffer (0.5% SDS, 0.2 M Tris-HCl pH 8, 0.025 M EDTA pH 8, 0.25 M NaCl). After cooling on ice for 5 min, 100 µL of 8 M potassium acetate was added and mixed by inversion. Samples were centrifuged at 13,000 rpm for 15 min, and the supernatant was collected. This step was repeated, followed by DNA precipitation with 300 µL of 100% isopropanol and centrifugation (13,000 rpm, 15 min). The DNA pellet was washed with 1 mL of cold 70% ethanol, centrifuged, air-dried at 42 °C for 20 min, and resuspended in 50 µL of warm water. Finally, 2 µL of RNase (10 mg/mL) was added, and the samples were incubated at 65 °C for 30 min before storage at −20 °C.

Confirmation PCR was performed using extracted genomic DNA from the candidates as template, the expected fragment size of positives was 1276 bp. The plasmid pIGEM002 was included as a positive control template. All three candidates were confirmed by PCR.

Test:

E. coli cultivation, sfGFP expression and CTX isolation:

The pIGEM001–sfGFP–CTX expression vector was transformed into E. coli BL21. Pre-cultures (5 mL LB medium supplemented with 50 µg/mL kanamycin) were grown overnight at 37 °C with shaking (200 rpm) and used to inoculate 500 mL of LB medium containing 50 mM sodium phosphate buffer (pH 7.0) and kanamycin. Cultures were grown at 37 °C until reaching an OD₆₀₀ of ~0.6, at which point expression was induced with 0.5 mM IPTG. Cells were incubated at 30 °C for 16 h.

Cells were harvested by centrifugation (4,000 ×g, 20 min) and resuspended in lysis buffer (20 mM sodium phosphate buffer, pH 7.0, 100 mM NaCl, 5 mM imidazole). Lysozyme (1 mg/mL final) and DNase I (25 U) were added, and the suspension was gently agitated for 1 h. Cells were then disrupted by sonication on ice (15 min total, 10 s on/10 s off, 70% amplitude). Cell debris was removed by centrifugation (10,000 ×g, 30 min, 4 °C), and the clarified lysate was used for affinity purification.

Purification was performed by Immobilized Metal Affinity Chromatography (IMAC) using 1 mL of Talon cobalt resin. The resin was washed with 30 mL ultrapure water, equilibrated with 20 mL binding buffer (20 mM sodium phosphate, pH 7.0, 100 mM NaCl, 5 mM imidazole), and incubated with the lysate for 15 min. The unbound fraction (flowthrough) was collected, and the resin was washed with the binding buffer prior to elution with 20 mM sodium phosphate, pH 7.0, 100 mM NaCl, and 500 mM imidazole. Eluted fractions containing sfGFP–CTX were analyzed by SDS–PAGE.

For CTX release, elution fractions were buffer-exchanged into tris buffer using centrifugal concentrators. TEV protease donated by NextVitro was added at a 1:50 (w/w) enzyme-to-protein ratio, and the reaction was incubated at 30 °C for 16 h. The cleavage mixture was passed through a 10 kDa molecular weight cut-off (MWCO) centrifugal filter, allowing the ~2.3 kDa CTX peptide to pass through while retaining sfGFP and TEV protease. The filtrate containing CTX was quantified by UV absorbance at 280 nm (ε = 5,500 M⁻¹ cm⁻¹) and stored at −20 °C until further use.

Quantification of CTX after TEV cleavage. The positive control CTX-UNESP (1 mg/mL) was diluted 5×. Samples CTX ft1 and CTX ft2 correspond to 1 mL of flow-through obtained from a 10 kDa filter after TEV cleavage. The results confirmed the presence of CTX in the purified fractions. From 1 L of E. coli cultivation, we obtained approximately 20 mL of purified sfGFP–CTX after column chromatography. For cleavage, we applied 50 µL of TEV protease to 1 mL of purified sfGFP–CTX, which yielded ~0.1 mg of CTX peptide. Based on this ratio, we estimated a production of ~2 mg of purified CTX per liter of E. coli culture, requiring about 1 mL of TEV protease for the cleavage step.

SDS-PAGE analysis of samples related to peptide purification and separation from GFP. Lane 1: reference sample provided by UNESP. Lane 2: our peptide sample concentrated 10×. Lane 3: our peptide sample before concentration. Lane 4: flowthrough fraction of our non-concentrated sample treated with TEV protease. Lane 5: residual material retained in the concentrator after centrifugation. Our sample was treated with TEV protease to cleave and release the target peptide from GFP.

Aspergillus oryzae cultivation and sfGFP expression:

With integration confirmed, we next performed expression assays to evaluate whether sfGFP–CTX was stably expressed and secreted under our chosen conditions. For this, we cultivated the transformants in medium containing 1% glucose + 2% maltose. The rationale was that sfGFP-CTX was engineered with the spGLA signal peptide, which directs the fusion protein to the secretory pathway. Since spGLA is maltose-inducible, the presence of maltose in the medium should trigger export of sfGFP–CTX into the extracellular environment.

We applied 20 uL (about 10ug of protein) of secretome into SDS-PAGE gel. A faint band at approximately 35 kDa—the expected size of sfGFP–CTX—was observed in all candidate transformants but not in the wild-type A. ory7 control, indicating successful integration and expression of the construct.

The SDS-PAGE was performed using 20 µL (~10 µg of protein) of secretome in gel and revealed a band at ~35 kDa in all three candidates, consistent with the expected size of the sfGFP–CTX fusion protein. Taken together, these results suggest that expression of sfGFP–CTX successfully occurred in the transformants.

To verify the presence of sfGFP-CTX expression in our candidates, we measured the fluorescence of the secretome from A. oryzae strain ory7 and the transformants C1, C2, and C3. The samples were analyzed in a 200 uL microplate reader using excitation/emission settings of 470 ± 15 nm and 515 ± 20 nm, respectively. Fluorescence values were recorded as relative fluorescence units (RFU), which provide a direct comparison of GFP signal across the different secretomes.

These results confirm higher fluorescence levels in all transformants compared to the parental strain, which does not express GFP. Taken together with PCR confirmation of construct integration and SDS-PAGE analysis, these data validate the successful production of sfGFP-CTX in all candidate strains. Based on protein quantification, we estimated a secretion of ~500 mg/L of total protein, of which approximately 20% corresponded to sfGFP–CTX (~100 mg/L). Since CTX accounts for ~10% of the sfGFP–CTX mass, this corresponds to ~10 mg/L of CTX.

Learn:

We confirmed the production of sfGFP–CTX in two different hosts: E. coli and the filamentous fungus Aspergillus oryzae. This result demonstrated that the fusion strategy successfully inactivated the peptide’s toxicity, allowing normal host growth while enabling later recovery of CTX after cleavage. In E. coli, the fusion protein was successfully extracted and purified using column chromatography. After TEV protease cleavage, the estimated yield was ~2 mg of CTX per liter of culture. However, we were not able to clearly visualize the peptide on SDS-PAGE gels, most likely due to its low concentration and small molecular weight. This highlighted the need for more sensitive analytical techniques. In future cycles, HPLC will be applied to accurately detect and quantify CTX at such low concentrations, ensuring reliable characterization of the peptide.

In A. oryzae, we confirmed both the integration of the construct and the production of the sfGFP–CTX fusion protein by SDS-PAGE and fluorescence. Although we did not have sufficient time to purify CTX from the secretome, this would represent the next logical step of the project. Based on protein quantification, we estimated ~10 mg/L of CTX, which is 5x more than E. coli production. Remarkably, this concentration was achieved under static liquid cultivation, suggesting that yields could be significantly improved under agitated fermentation conditions.

These production data were also incorporated into our Entrepreneurship section, where we evaluated the economic feasibility of CTX production. By comparing yields from different processes, together with purification costs and scalability, we developed projections to assess whether CTX could be manufactured at a viable cost for large-scale application in Brazilian citriculture.

Yeasts such as Saccharomyces cerevisiae have long been used in biotechnology due to their safety and robustness in large-scale bioprocesses, such as ethanol production. They can grow to high cell densities using inexpensive substrates, making them an attractive platform for producing recombinant proteins and peptides. By leveraging yeast’s natural potential, we aimed to create a cost-effective and scalable strategy for CTX production. Although S. cerevisiae demonstrated greater resistance to CTX compared to the other hosts, we still applied a fusion-based strategy. This approach was essential because even partial growth inhibition can compromise yield and long-term stability in large fermentations. By coupling CTX to carrier proteins, we ensured both safe intracellular expression and the possibility of controlled release of the active peptide after production.

Design:

Taking advantage of the experience of our scientific team leader Bruno Batista, who participated in iGEM 2019 with Team SÃO-CARLOS:BRAZIL working on a project that used surface display to aggregate melanin on the yeast cell wall, we developed the idea of coupling CTX to the yeast surface and then releasing it efficiently. To explore the possibility of producing CTX in yeast, we designed a strategy that combines yeast surface display with a sequence-specific chemical cleavage system (SNAC-tag) to enable cost-effective production ^[3]. In this system, CTX is genetically fused to the yeast cell wall protein Aga1p, which is naturally anchored to the yeast surface via a GPI-anchor. Unlike the traditional Aga1–Aga2 display system, this design does not require fusion of the protein of interest to Aga2p, thereby avoiding dependence on disulfide bond formation between Aga1p and Aga2P ^[4]. To allow peptide release, we inserted a SNAC-tag between the linker–V5 epitope–Aga1p fusion and CTX. The SNAC-tag enables highly specific cleavage in the presence of Ni²⁺ ions, releasing CTX with a precise N-terminal sequence. This strategy eliminates the need for proteolytic enzymes such as TEV protease, which are often costly and introduce additional steps in downstream processing.

Design:

The plasmid pCY002_p426GPD-ScAGA1, kindly donated by Gustavo Seguchi, was digested with XhoI. Primers were designed to amplify the CTX sequence from pIGEM001, introducing XhoI sites at both the 5’ and 3’ ends; the 3’ end also included the sequence for the SNAC-tag. After obtaining and digesting the fragments, they were ligated using T4 ligase and transformed into E. coli. Colony PCR confirmed the correct construct, which we named pIGEM004 – CTX-SNAC-ScAGA1. Subsequently, a miniprep was performed to recover the plasmid, which was then used to transform S. cerevisiae (BY4272). Yeast candidates that grew on selective medium (lacking uracil and uridine) were sequenced to confirm successful transformation.

Test:

We cultivated yeast expressing Aga1p-CTX and the control strain expressing only Aga1p for 48 hours in YNB medium. From each culture, 10 mL of biomass (equivalent to OD600 ~2) was collected and resuspended in 10 mL of cleavage buffer (1:1 ratio with OD600) for 18 hours at 25 °C and 200 rpm. After incubation, the samples were centrifuged at 3,000 rpm, and the supernatant was collected. The samples were concentrated ~30x and CTX presence was then measured using a Nanodrop to assess peptide production in both CTX and control samples.

The average of the samples was calculated, and the value from the control was subtracted, as the control did not contain the peptide in the membrane. Since the Ni²⁺ treatment could not cleave anything, it was considered potential noise.

The CTX peptide was recovered from 1 mL of Saccharomyces cerevisiae culture at OD₆₀₀ = 2.0 following treatment with nickel cleavage buffer for 18 h at 25 °C. At this cell density, the biomass concentration is estimated at ~0.8 g/L, corresponding to 800 µg of dry biomass per milliliter of culture. From this sample, we obtained ~12 µg of CTX, which represents ~1.5% of the total treated biomass. The production estimation titer was 12 mg of CTX per liter of yeast culture.

Learn:

We learned that our fusion and cleavage strategy can successfully release CTX from yeast biomass, reaching measurable yields. Although the peptide represented ~1.5% of the treated biomass, about 12 mg/L, this result confirmed that yeast can act as a feasible production host. However, we also observed that the nickel buffer may interfere with absorbance readings in the Nanodrop, since some signal was detected even in the negative control without CTX expression. This indicates that our current quantification could be overestimated. To address this, in future cycles we plan to apply HPLC-based methods for accurate characterization and quantification of the produced peptide. Nevertheless, the approach showed strong potential: using yeast biomass as a production platform together with nickel-assisted cleavage could significantly reduce production costs, which is a key factor for scaling CTX manufacturing in an economically viable way.

Aspergillus oryzae is well known for its exceptional ability to secrete large amounts of proteins, particularly amylases, whose main role is the degradation of polymers such as starch and maltose ^[5]. Among the secreted proteins, amylases are consistently among the most abundant. The A. oryzae strain contains three copies of amylase genes (AmyA, AmyB, and AmyC) in its genome, which ensures their strong expression. Previous studies have shown that the heterologous expression of proteins in filamentous fungi is often very low, usually in the milligram range, whereas the expression of native homologous proteins such as amylases can reach gram levels ^[6]. To take advantage of this natural secretion capacity, our team designed a strategy to couple CTX to the three amylases of A. oryzae (AmyABC). By fusing CTX to these highly expressed proteins, we aimed to leverage the fungus’s powerful secretion system and thereby achieve high-yield production of the peptide.

Design:

The amylases are present in three copies in the genome of A. oryzae. These AmyABC genes are homologous, sharing the same promoter sequence, while their terminators differ: AmyB and AmyC share the same terminator, whereas AmyA has its own unique terminator. Our strategy was to fuse K7–CTX at the C-terminus of each amylase, inserting it immediately before the stop codon. This design would allow the fungus to continue producing its natural amylases, while simultaneously secreting CTX coupled to them. To achieve this, we designed a CRISPR/Cas9 editing strategy with two guide RNAs (gRNAs) to target all three amylase loci simultaneously. As donor DNA, we used synthetic fragments (obtained through IDT sponsorship) containing the fusion sequence, enabling homology-directed repair to insert the K7–CTX module as illustrated in the scheme below.

Build:

The gRNAs were designed using the EuPaGDT online tool. To increase CRISPR/Cas9 editing efficiency, we selected two gRNAs targeting the amylase locus: one located in the middle of the gene and the other positioned near the stop codon. This dual-target strategy was intended to maximize the likelihood of successful cleavage and insertion of the donor DNA. The gRNAs were cloned into the pFC9330 system used in DIVERSIFY by NEBuilder.

The donor DNA was received from IDT as a linear gene fragment. To facilitate its use, it was designed to be cloned into the multiple cloning site (MCS) of pRS426 upon arrival. The donor fragment contained 750 bp of upstream and downstream homology arms to enable efficient homologous recombination in A. oryzae. In addition, to prevent potential re-cleavage by one of the guide RNAs, a codon modification (W369F) was introduced into the sequence.

From each transformation, six transformants were isolated and screened by PCR to confirm K7-CTX insertion at the target locus. The positive candidates should have 530 pb, while the negative 370 pb. As we can see below, all candidates were negative, indicating the absence of cassette integration.

Test:

We performed spore PCR to confirm the integration of CTX into the three amylase genes (amyA, amyB, and amyC). However, all candidates tested negative, indicating that the insertion was not successful.

Learn:

Due to time limitations, we were not able to complete the full construction of the amyABC–CTX system during this cycle. Although no positive transformants were obtained in this first attempt, the strategy remains promising. We raised some hypotheses to explain the negative results, such as the possibility that the gRNAs were not efficiently cleaving the desired sites or that transformation efficiency needs to be optimized. Despite these challenges, our team plans to continue developing and refining this approach in future cycles. Coupling CTX to highly secreted amylases could provide a powerful strategy to increase peptide yields and enhance the overall feasibility of the production system and open the doors to solid fermentation production.

Every year, the Brazilian orange juice industry generates millions of tons of residues during juice extraction. These by-products are traditionally destined for a variety of secondary uses (LDC), but their potential remains largely underexplored. Our team saw in this challenge an opportunity: to turn citrus waste into a resource for sustainable peptide production. By harnessing the natural ability of Aspergillus oryzae to grow on complex substrates and secrete large amounts of amylase enzymes ^[7], we aimed to establish a low-cost and eco-friendly system for CTX production through our AmyABC–CTX coupling strategy. To achieve this, we explored the use of solid-state fermentation (SSF) with orange peel residues as the growth substrate.

Design:

We first set out to determine whether Aspergillus oryzae (Ory7) could grow on orange peel residues and secrete amylases under solid-state fermentation (SSF). Our hypothesis was that SSF on citrus waste would support robust fungal growth and enzyme secretion, providing a viable base for peptide production across our coupling strategy. Once this baseline was established, our second objective was to cultivate genetically engineered A. oryzae carrying the AmyABC–CTX fusion strategy, to assess whether CTX could be co-produced via high-expression amylase carriers under the same SSF conditions.

Build:

We searched for studies at UNICAMP that had used orange bagasse for enzyme production and found a doctoral thesis that employed juice industry residues as substrate ^[8]. From this, our Human Practices team contacted Gabriela Macedo, from the Department of Food Engineering, who kindly provided us with orange peel residues and valuable advice on how to set up the cultivation.

Test:

Orange peel residues donated by Professora Gabriela Macedo, were processed using a pulse blender and sieved through two meshes (1.68 mm – Tyler 10 and 0.35 mm – Tyler 42), resulting in an intermediate particle size. According to the work, an average size of 0.7 mm is commonly used, but this sieve was not available in the laboratory, so we combined the two mesh sizes as an alternative.

Each 125 mL Erlenmeyer flask was prepared with 30 mL of the ammonium sulfate (5 g/L) and 2.4 g of orange peel substrate (1.68–0.35 mm particles). Flasks were autoclaved at 121 °C for 20 minutes. Since supplementation with uracil/uridine was forgotten prior to sterilization, it was added afterwards. The cultures were inoculated with 1 mL containing 10⁶ Ory7 spores. After cultivation, 20 mM acetate buffer (pH 5) was added to the fermented substrate at a ratio of 10 mL per gram of substrate. The samples were shaken at 200 rpm for 1 hour, filtered through paper to remove biomass, and the flow-through was stored as the secretome.

Protein quantification of the secretome was carried out by the Lowry method. The day 0 control (no fungus added) showed a relatively high protein signal, which we attribute to compounds released from the substrate. However, as seen in later gels, these proteins were likely consumed by Ory7, since strong bands visible at day 0 were not present at days 3 and 5. To avoid overestimation the value from day 0 was removed from total protein quantification of days 3 and 5.

Each well was prepared with 20 µL of sample and 5 µL of loading buffer. Samples were heated at 95 °C for 5 min and then applied in gel. The electrophoresis was run at 110 V for 20 min followed by 130 V for 1 h, stained for 4 h with coomassie blue, and destained for 24 h with distilled water.

A visible band at ~50 kDa appeared in the day 3 and day 5 samples, consistent with the size of amylases. This result confirmed that A. oryzae was able to grow on orange peel biomass and secrete proteins, validating the feasibility of using citrus residues as substrate for protein production. Protein quantification of the secretome by Lowry revealed approximately 1100 mg/L of total protein. SDS-PAGE analysis suggested that about 60% of this protein fraction corresponded to amylases, equivalent to ~660 mg/L in the culture supernatant. Since CTX (3 kDa) was fused to the amylase backbone (50 kDa), we estimated that ~6% of the amylase mass could correspond to CTX, yielding ~40 mg/L of peptide. In the experiment, 2.4 g of orange peel residue was incubated in 30 mL of nitrogen solution plus 20 mL of protein wash buffer, resulting in a final concentration of 40 mg/L of CTX. This corresponds to a total of 2.0 mg of peptide, which scales to approximately 830 mg of CTX per kilogram of residue under solid-state fermentation.

Learn:

Through solid-state fermentation, we confirmed the growth of Aspergillus oryzae and estimated a yield of ~830 mg of CTX per kilogram of orange peel residue, using the amyABC-CTX strategy. Although we were not able to complete the construction of CTX fused to the amylases (Part 6), this strategy indicated the highest potential titer among all approaches tested as you can see in Entrepreneurship page. Importantly, it also leveraged a low-cost by-product of the orange juice industry, reinforcing our commitment to circular economy and sustainability. These findings highlight the strong potential of coupling CTX to highly secreted amylases as a scalable, cost-effective production strategy for future cycles, and guide our next steps in refining this approach.