Linear Array Epitope

The results of template-repeated PCR (TR-PCR), shown in Figure 1, indicate that the length of the tandem-repeated sequence (TRS) varies with the concentration of the primer. We found the typical trend that the lower the concentration of primer, the higher the number of repeats of the TR-PCR products. However, the product concentration decreases when the primer concentration increases. In our project, the optimal primer concentrations are 0.2 μM for TRS-151*7 and 0.08 μM for TRS-110*3.

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Figure 1. The TR-PCR results of (a) TRS-151*7 and (b) TRS-110*3. The values listed on each represent the concentration of primer (in μM). The yellow arrow shows the typical ladder pattern of TR-PCR. M = DNA marker.

TRS-110*3 and TRS-151*7 obtained from TR-PCR were transferred to adaptor PCR (AD-PCR). The ladders shown in Figure 2 are the products obtained from AD-PCR. The sequences ranging in size from 200-400 bp and 50-200 bp, identified as TA-151 and TA-110, respectively, were extracted and ligated to TA vectors. Mutiple bands found in the results of AD-PCR due to the fact that there were several sites available for ligation of the adapter. Therefore, various lengths of sequences were generated during AD-PCR. These ligated sequences were then successfully transformed into DH5α competent cells. The colonies shown in Figure 3 suggest the bacteria survives due to the antimicrobial peptides (AMP) in the TA vector.

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Figure 2. The results of AD-PCR of (a) TRS-151*7 and (b) TRS-110*3. The value listed on each lane represents the concentration of primer (in μM). M = DNA marker.

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Figure 3. The culture plates suggest that TA-151 (a) and and TA-110 (b) were successfully transformed into DH5α competent cells.

The results of colony PCR show that the sizes of TRS-151*7 (Figure 4(a)) and TRS-110*3 (Figure 4(b)) are estimated to be 420 bp and 300 bp, respectively.

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Figure 4. Colony PCR of (a) TRS151*7 and (b) TRS110*3. The sizes of TRS-151*7 and TRS-110*3 are estimated to be 420 bp and 300 bp (including the M13 primers and some sequences on the TA-vector), respectively. The numbers above the lanes indicate colony number. M = DNA marker.

Plasmids from these colonies were extracted, and TRS were obtained using the restriction sites of Ncol and Hindlll on the TA vector. TRS-151*7 and TRS-110*3 have sizes of 200-300 bp (Figure 5(a)) and 90-100 bp (Figure 5(b)), respectively, which fits our expectations.

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Figure 5. The result of digestion of (a) TA-151 and (b) TA-110, which have sizes of 200-300 bp and 90-100 bp, respectively. M = DNA marker.

Finally, these TRSs were extracted and ligated to pET-29b(+). However, we ran into a problem during transformation of pET-29b(+) into DH5α. Currently, we are still seeking to optimize the transformation conditions.

Envelope Protein (E protein)

A plasmid containing the whole structural protein of the dengue virus was obtained from National Health Research Institute. The Envelope protein (E protein) with two HA tags from the plasmid were amplified using PCR. Figure 6 shows that the sequence of the E protein with the HA tags were amplified, matching the expected size of 1,551 bp.

Figure 6. E protein with two HA tags, matching the expected size of 1,551 bp. M = DNA marker.

We prepared pET-29a(+) as the vector for expressing E protein and confirmed in Figure 7 that plasmid extraction shows the size of pET-29a(+) is about 5,000 bp, which is close to the theoretical size, 5,371 bp. Also, we transformed the pET-29a(+)_E protein into DH5α and check the transformation using colony PCR. Figure 8 shows the size of E protein with two HA tags, and T7 promoter and terminator is about 2,000 bp, close to the theoretical size of 1,903 bp.

Figure 7. Plasmid extraction of pET-29a(+), matching the expected size of about 5,000 bp. M = DNA marker.

Figure 8. Colony PCR of pET-29a(+)_E protein, which has a size of about 2,000 bp, containing E protein with two HA tags, and T7 promoter and terminator. M = DNA marker.

pET-29a(+)_E protein was then transformed to BL21(DE3), cultured with 3 ml medium at 37 °C, and induced with 1.8 ml bacteria and 1 mM IPTG for 2 hours. Figure 9 shows the results from four different colonies with and without induction. The weak bands from 63 to 48 kDa indicate the E protein expression with the HA tag, which has a size of 61.2 kDa. However, it is too hard to see, and we are trying to improve the expression. Also, the bacterial culture was lysed with sonication, then separated with a high-speed centrifuge. It indicates that E protein found in the supernatant in Figure 10. The results were also confirmed using Western blot based on the HA tag, which is part of pET-29a(+). Figure 11 shows that the anti-His tag binds to the His tag, suggesting that the protein expression was successful.

Figure 9. SDS-PAGE of E protein from a small-scale culture. After induction with IPTG, the strong bands at about 61.270 kDa indicate the expressing of E protein. M = protein marker. NI = Non-Induction. I = Induction.

Figure 10. SDS-PAGE of E protein from a small-scale expression. The bacterial solution was lysed with sonication and separated with a high-speed centrifuge. The results indicate E protein is expressed in the supernatant. M = protein marker. Sup. = supernatant.

Figure 11. Western blot of E protein. An anti-His tag was used to bind to the His tag on the E protein. The stronger band on the both non-induction and induction lanes suggests the experiments were successful. NI = Non-Induction. I = Induction.

CLEC5A

A plasmid containing the CLEC5A extracellular domain with a Myc tag was obtained from OriGene Technologies. The CLEC5A extracellular domain and Myc tag were amplified using PCR. Figure 12 shows that the sequence of the CLEC5A extracellular domain with a Myc tag were amplified successfully, with a size of 564 bp. pET-29b(+) was prepared successfully as a vector, as shown in Figure 13, with a theoretical size of 5,370 bp. pET-29b(+)_CLEC5A was obtained by inserting the CLEC5A into pET-29b(+) and then transforming it into DH5α which was confirmed using colony PCR.

Figure 12. CLEC5A with a Myc tag, matching the expected size of about 600 bp. M = DNA marker.

Figure 13. Plasmid extraction of pET-29b(+), matching the expected size of about 5,000 bp. M = DNA marker.

Figure 14 shows the size of CLEC5A extracellular domain with Myc tag, and T7 promoter and terminator is about 900 bp, close to the theoretical size of 904 bp.

Figure 14. Colony PCR of pET-29b(+)_CLEC5A, which has an expected size of about 900 bp including the CLEC5A extracellular domain and Myc tag and T7 promoter and terminator. M = DNA marker.

pET-29b(+)_CLEC5A was then transformed to BL21(DE3) and induced to express protein with 1.8 ml bacteria and 1 mM IPTG for 2 hours. Figure 15 shows the results from four different colonies with and without induction. The strong bands near 28 kDa indicate expression of the CLEC5A extracellular domain with the Myc tag, which has a size of 26.6 kDa. The results were also confirmed using Western blot based on the HA tag, which is part of pET-29b(+). Figure 16 shows that the anti-His tag binds to the His tag, suggesting that the protein expression was successful.

Figure 15. SDS-PAGE of CLEC5A from a small-scale culture. After induction with IPTG, the strong bands at about 26.6 kDa indicate the expressing of CLEC5A. M = protein marker. NI = Non-Induction. I = Induction.

Figure 16. Western blot of CLEC5A. An anti-His tag was used to bind to the His tag on the CLEC5A protein. The stronger band on the induction lanes suggests the experiments were successful. NI = Non-Induction. I = Induction.

Finally, we expressed CLEC5A on a large scale. The bacterial culture was lysed using a French press, then separated with a high-speed centrifuge. Figure 17 indicates CLEC5A is always found in the pellets no matter how long they were induced.

Figure 17. SDS-PAGE of CLEC5A large-scale expression. The protein was induced for 30 minutes, 1 hour, 2 hours, 3 hours, and 4 hours, separately. The bacterial solution was lysed with a French press and separated with a high-speed centrifuge. The results indicate CLEC5A is always expressed in the pellet under these conditions. M = protein marker. Sup. = supernatant.

Conjugations of Primary Amines to the Glass Fiber Membranes

We expressed green fluorescent protein (GFP) as a mock E protein to show that that after modification, the glass fiber membrane can bind to the primary amines (side-chain amines) from a peptide or protein. We appreciate Mingdao iGEM 2020, who generously provided us with a plasmid containing the green fluorescent protein gene (GFPmut1, BBa_K1159311). The modified glass fiber membranes were immersed in GFP supernatant from the culture medium for 30 mins to form amide bonds. To verify that the experiments work, we measured the intensity of the fluorescence from GFP using a Synergy H1 Hybrid Multi-Mode Microplate Reader (also provided by Mingdao iGEM 2020). The intensity of membrane reaction with 0.5x and 1x GFP supernatant are shown in Figure 18. We carefully washed the membranes with double-distilled water after the reaction. The intensity was about twice as strong in the 1x supernatant as the 0.5x one, suggesting the conjugation experiments were successful. We also performed the same experiment with non-modified glass fiber membranes as a control, and the intensity of fluorescence is within the range of measurement error.

Figure 18. Fluorescence from modified glass fiber membranes reacting with 1x GFP stock, modified glass fiber membranes reacting with 0.5x GFP stock, and non-modified glass fiber membranes (control).

We also confirmed the reactivity of glass fiber membranes to amines using DNA primers (TRS-110R*7, BBa_K3648007) by measuring the fluorescence spectra with a plate reader (provided by Mingdao iGEM 2020). We used the same coupling conditions for GFP. Figure 19 shows that with the modification, the emission has a 0.5-fold enhancement in DNA conjugation.

Figure 19. The fluorescence spectra of DNA conjugated glass fiber membrane (blue), the control experiment (red), in which DNAs react with non-modified membranes, and the empty well (yellow). The spectra were obtained at an excitation of 320 nm.

Gold Nanoparticles (AuNPs)

Avoiding the aggregation of AuNPs

To conjugate the peptides to the AuNPs, pre-treatment of the AuNPs is necessary. However, during this process, aggregation of AuNPs could happen, which would result in failure of the experiment. We measured the sizes of AuNPs using dynamic light scattering shown in Figure 20.

First, we used MUA/MCH to modify 13-nm AuNPs by adding carboxylic groups to the surface. However, the AuNPs aggregated to a size of about 1,800 nm after we applied the EDC/NHS treatment. We found that if we replaced EDC with DCC, the size of AuNPs aggregation was reduced to about 350 nm. But 350 nm is not small enough, so we further tried replacing MUA/MCH with MHA /SB thiol, which was then treated with EDC/NHS. We obtained a significantly smaller size of 90 nm. We used these particles for the following experiments.

Figure 20. The size distributions of AuNPs modified using MUA/MCH + EDC/NHS (blue), modified using MUA/MCH + DCC/NHS (red), and modified using MHA/SB thiol + EDC/NHS (yellow)

Abbreviations: MUA: 11-mercaptoundecanoic acid; MCH: 6-hydroxy-1-hexanethiol; EDC: 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide; NHS: N-hydroxysuccinimide; DCC: N,N’-dicyclohexylcarbodiimide; MHA: 16-mercaptohexadecanoic acid; SB thiol: 1-(2-sulfosulfanylethylamino)tetradecane

Conjugations of primary amines to AuNPs

As a proof of concept that we are able to form the covalent bonds between the primary amines (from the PTRS) and AuNPs, we tried to conjugate the DNA primers (TRS-110R*7, BBa_K3648007) with the modified AuNPs.

We used Raman spectrometer verify if the experiments worked. We found that with the DNA conjugation, the Raman signals have a significant decrease (Figure 21). Although we have no model to explain this effect, we believe it resulted from interactions with the DNA, suggesting DNA can bind to AuNPs.

Figure 21. The Raman spectra of DCC/NHS modified AuNPs (yellow), and DCC/NHS modified AuNPs conjugated with 1 μM (blue) and 0.1 μM (red) DNA primers.

Abbreviations: DCC: N,N’-dicyclohexylcarbodiimide; NHS: N-hydroxysuccinimide