Goal:

Design NOS construction and express Nitric Oxide Synthase (NOS) gene.

Achievements:

1. Construction of NOS

2. Run SDS-PAGE to confirm expression in E. coli

This year, our team aims to reduce intraocular pressure (IOP) through the contact lens with engineered E. coli that can produce Nitric Oxide (NO). We aimed to make our plasmid express NOS and support our bacteria to bind to the contact lenses. Since E. coli doesn't produce nitric oxide, we ordered a sequence containing a lacO-T7 promoter, B0034 RBS, and bsNOS. We ligate the sequence by inserting it into the pSB1C3 plasmid and combining it with another sequence (pLac, RBS, csgA, RBS, csgD, LacI) from IDT, then transformed into E. coli DH5-Alpha. By doing so, NOS can convert L-arginine into nitric oxide, then release it into the eyes^[1].

To ensure the plasmid contains those two sequences, we conducted PCR with their primer separately. Fig. 1 shows that the plasmid contains both of the desired sequences.

In order to test the function of the T7 promoter, we transformed the plasmid into BL21(DE3). Next, to observe the effects of various IPTG concentrations on NOS expression, we performed SDS-PAGE with different IPTG concentrations. The bacteria is cultured for two hours and induced with IPTG for 12 hours. In Fig. 2 we can observe that the first to the third lane express a similar thin band. Meanwhile, we can see a thicker band in the fourth lane, which means it has a higher protein expression when the IPTG concentration is 1 mM.

Goal:

Control NOS production by IPTG inducible systems to produce more nitric oxide.

Achievements:

1. Measure the NOS activity in different IPTG concentration at different time

2. NOS kinetic test

As we decided to use an IPTG inducible system, we conducted an experiment to determine whether IPTG concentration and induction time can control the production of NO. Therefore, we test the IPTG system by using different concentrations and induced at different times. Here, we are using E. coli BL21(DE3) strain. As seen in Fig. 3, the nitric oxide production is both time dependent and IPTG dependent. Thereby, we can observe whether the IPTG inducible system can effectively produce nitric oxide in a certain induction time.

After confirming the production of NO can be induced by IPTG, we test the kinetic of NOS. With Fig. 4, the bacteria are cultured for 12 hours and induced with 0.1 mM IPTG for 2 hours. We then applied the homogenized bacteria and substrate to the cornea of the porcine eye to observe whether the concentration of nitric oxide will increase in the eye or not. The substrate runs out within 20 min and produces approximately 1.6 nmol nitric oxide. Through the graph below, we can observe that when there is 1.6 nmol nitric oxide in the porcine eye, it will diffuse into the cornea and the concentration of nitric oxide will increase six times higher when it reaches aqueous humour.

We are taking advantage of the IPTG inducible system to control NOS expression, thus producing nitric oxide. From all the experimental results above, we have proved that our system is functioning well. Starting from NOS construction, confirm NOS expression, until NOS functional test, we have provided convincing results to support our ideas.

Goal:

Enhance biofilm production and improve the binding affinity of our engineered bacteria.

Achievements:

1. Construction of csgD and csgA

2. Run SDS-PAGE to confirm expression

3. Functional test to measure the amount of biofilm production and binding affinity

We are very grateful that one of our PIs, Professor Masayuki Hashimoto, from the Institute of Molecular Medicine, with generosity, was providing us a BW25113 wild type strain, BW25113 △csgD, and PCA24N plasmid to supported our csgD and csgA experiment.

As a first step, we construct plasmid that only contains pLac-csgD on BW25113 and an improved version that contains NOS, csgD and csgA using gBlock provided by IDT. After that, we transformed into E. coli BL21(DE3) strain. Then, we use PCR to confirm that our bacteria carried csgD and csgA. The expected results are shown in Fig. 6.

Furthermore, we ran SDS-PAGE to check the protein expression of CsgD and CsgA. We cultured the bacteria for 2 hours, and added IPTG to induce for 12 hours long, then adjusted the OD₆₀₀ to three. We expected to see CsgD protein size around 25 kDa^[2] and CsgA around 15 kDa^[3]. The results below have shown the expected size we want for CsgD and CsgA protein expression.

Next, for further confirmation, we use a congo red dye to see the curli expression. If there is a biofilm, the absorbance value at 500 nm will be higher^[4]. We use BW25113 as control, BW25113 with csgD, and NOS-csgA-csgD on BL21(DE3). First, we cultured for 2 hours and added IPTG to induce. After overnight culture, we add congo red to 1 ml of bacteria and leave it for 2 hours. Next, we centrifuge to separate the supernatant and pellet. By using a microplate reader, we can measure the absorbance value at 500 nm and normalized by 600 nm wavelength. Through Fig. 7 below, we can see that the amount of biofilm is increased after adding csgA.

After conducting several research, we found a common way for biofilm staining by using 96-well plate assay. We soon discovered that most of the experimental error came from the staining procedure and the steps of removing supernatant which decrease the accuracy. We try to remove the supernatant by hitting the plate to the table. As shown in Fig. 8, the data standard deviation is high.

However, the standard protocols to test biofilm formation cannot directly support our claims to increasing binding affinity. Therefore, we come up with a simple and quantifiable measurement protocol for evaluating biofilm binding ability. To observe the adherence of bacteria to the contact lens surface, we culture the bacteria for 2 hours then induced with IPTG. Then, we proceed into a 96-well plate and leave for 12 hours of incubation. Afterward, we used external forces to test the bacteria binding affinity by free falling an object - a book, from different heights and measuring the OD₆₀₀ value to determine its concentration. As the concentration increases, the bacteria will have a greater binding affinity.

The results show the bacteria overexpressing CsgA-CsgD can stand greater force. Thereby, we can prove that overexpressing CsgA-CsgD not only enhances biofilm production, but our bacteria can also bind properly to the contact lens.

Goal:

To prevent our engineered bacteria surviving outside the contact lens.

Achievements:

1. Tc and heat inactivated cTc characterize

Our first kill switch design used TetR fused with VP16, tet-off system^[5] to control the Colicin E7(BBa_K117000) as our biosafety measure. We will put tetracycline analog into the contact lens, this will prevent the VP16 to activate the lysis gene. If the bacteria leave the contact lens, where the environment has low concentration of tetracycline analog, the lysis gene will be activated. (Fig. 1)

There are several options when it comes to tetracycline analog that can act on the tetR protein, changing its DNA binding affinity, however, according to Biomol GmbH-Life Science Shop, 50 mg of Anhydrotetracycline costs 54 US dollars^[6]. In search of a cheaper alternative, we turned to tetracycline(TC)^[7] and Chlortetracycline(CTC)^[8], which has been reported to bind to tetR protein and affect its affinity to tetO.

After many attempts constructing this plasmid resulted in failure, we turned to gene knockout as our biosafety masure. However, we can still characterize the effect of TC and CTC on TetR using iGEM BioBrick BBa-K611059. (Fig. 2)

BBa-K611059 has two open reading frames, a pTet promoter controlled GFP, and pBAD promoter controlled TetR. As we use arabinose to produce TetR to repress pTet, this allowed us to try different tetracycline analogs and observe GFP signal recovery to assess their potency. Since most tetracycline analogs have antimicrobial activity, we test two different ways to compensate for this effect, heat inactivation of tetracycline analogs and co-transform tetracycline resistance plasmid.

We first transformed the plasmid carrying BioBrick BBa-K611059 into BW25113, which cannot metabolize arabinose, and added different concentrations of heat inactivated CTC^[9] to see if the GFP signal is restored.

After 12hr of culture with different concentrations of heat-inactivated CTC and 0.0001% of arabinose, we can see as arabinose was added into the culture, the normalized GFP signal slightly decreased, suggesting BBa-K611059 functions correctly. As 20µg/ml heat-inactivated CTC can rescue the GFP signal, and 30µg/ml will result in higher GFP signal. Also, through the OD600 results we can also conclude the heat-inactivated CTC does not affect E. coli growth.

Despite showing that heat-inactivated CTC can activate pTet, we were not satisfied with the dynamic range, so we decided to challenge using TC itself, since this operon originated in response to antibiotic resistance. In concerns of TC will be deadly to E. coli itself, also there has not been reported heat-inactivation TC can activate pTet, we co-transformed a TC resistance plasmid, pACYC184, into BW25113 to conduct further experiment.

We first tried two different arabinose concentrations, 0.002% and 0.0002%, both concentrations show repress GFP signal. However, bacteria experienced growth retardation in 0.002% of arabinose, we speculate that the high arabinose concentration induces osmotic stress to the bacteria. Also we can see as TC concentration increases, although the GFP signal remains the same, the OD600 decreases.This indicates that TC can induce pTet to produce GFP but suppress bacteria growth. We speculate that pACYC184 can not withstand such high concentration of TC.

From these results, since both TC and heat-inactivated CTC can activate pTet, but TC seems to affect cell growth at higher concentrations although we supplied the bacteria with resistance plasmid, we can conclude that heat-inactivated CTC is a better inducer compared to TC.

Goal:

To prevent our engineered bacteria from surviving outside the contact lens.

Achievements:

1. Transform plasmid with T7 polymerase gene into WM3064

2. Successfully prove our bacteria will not survive without diaminopimelate (DAP)

We use the knockout strain WM3064 which deleted the gene dapA encoding for 4-hydroxy-tetrahydrodipicolinate synthase that is essential for cell wall synthesis^[10] and growth. For the growth, the strain WM3064 must rely on the supplement of diaminopimelate (DAP) in medium; in other words, the bacteria can’t survive without DAP. With this concept, we designed our kill switch for safety.

We transformed a plasmid carrying T7 RNA polymerase into WM3064 to control the expression of NOS by the T7-lacI promoter. After that, we ran SDS-PAGE to ensure the protein expression of NOS under the T7-lacI promoter. The expected protein size of NOS is 40 kDa. As seen in Fig. 14, our WM3064 with T7 RNA polymerase can successfully produce NOS.

In order to prove that E. coli WM3064 can’t survive without DAP, we spread our E. coli WM3064 with T7 RNA polymerase to different plates. As seen in this result (Fig. 15), the bacteria can survive on the plate with exogenous DAP. When there is no more DAP supplied, the bacteria cannot survive.