Week 1

Digestion and Isolation of pSB1C3 Backbone.

The purpose of digesting the pSB1C3/PArsRGFP construct was to separate the backbone from the former GFP insert. Tacoma RAINmakers sought to isolate the pSB1C3 backbone employed in their 2017 construct, as the GFP reporter complex was no longer desired. Apparent disadvantages of GFP indication in biosensors are ultraviolet readings. The RAINmakers prefered a chromoprotein that produces color in the visible spectrum. Enzymes XbaI and SpeI were used to cleave the terminator sites of the vector, freeing the pSB1C3 backbone. NEB resources confirmed that the Cutsmart Buffer 2.1 is the most compatible with these particular enzymes, providing an optimized environment for digestion. Combining the reagents listed in Table 1.0, the reaction was set at 37ºC in a water bath for 1 hour and 45 minutes. This reaction was completed in duplicate to increase statistical probability of desired backbone DNA.

Following completion of pSB1C3 digestion, Tacoma RAINmakers employed a standard procedure listed in the protocol page as “Agarose Gel Electrophoresis.” The purpose of this gel was to confirm if the backbone DNA had been successfully isolated from the undesired GFP insert. As cited in Figure 1, the expected pSB1C3 bands were located at 2000bp. A gel extraction process, also outlined in the protocol section, was completed in order to remove and contain the digested pSB1C3 DNA.

Fig. 1. DNA gel of digested parts.

Week 2

Digestion and Ligation of spisPink, amilCP, and PcArsR Inserts

Using IDT stock solutions of chromoprotein and arsenic regulatory DNA, Tacoma RAINmakers completed a standard restriction digest (see protocol page for further information). The notable difference between insert and backbone digestion are the enzymes. Each insert included a SalI enzyme site, which is not compatible with the BioBrick suffix/prefix of the vector. Instead, the SalI site is used to ligate the chromoprotein to PcArsR, rendering the ends of this complete insert as compatible for sticky-end ligation to pSB1C3. After combining reagents listed in Table 2.0, reaction was held in a water bath at 37°C overnight.

Once the SalI site in spisPink, amilCP, and PcArsR was successfully sticky-ended, Tacoma RAINmakers were prepared for ligation of each chromoprotein to the arsenic regulator. 60ng of each part were used in order to ensure that there would enough DNA material for the ligation. Having combined the substances from Table 2.1, the reaction was set to ligate overnight at 16°C. Afterwards, SalI was heat inactivated at 80°C, ensuring a complete denature, since the enzyme was no longer required.

Week 3

Digestion and Ligation of Insert (amilCP/spisPink + PcArsR) to pSB1C3 Backbone

Following the ligation of the chromoprotein and PcArsR, the complete insert was digested with enzymes compatible with our pSB1C3 backbone. This process allows sticky-ended ligation in the next step, which increases the chance of a proper insert-backbone ligation. 84 ng of insert DNA was pipetted into the reaction alongside the other reagents mentioned in Table 3.0. The digestion was set at 37°C in a water bath for 1 hour and 30 minutes.

Tacoma RAINmakers combined the reagents listed in Table 3.1 to ligate the completed insert to the vector. The reaction included a negative control that contained only vector DNA. A notable process involved in ligation reactions is calculating DNA volumes. Typically, a 1:3 ratio of vector to insert ensures that there is a balance of both parts. The RAINmakers employed the NEBioCalculator to determine how many moles were in 1ng of vector (2070bp) and 1ng of insert (1488bp). This calculation translated to 0.8µL of vector and 20µL of insert. The ligation occurred at 16ºC overnight and was heat inactivated at 80ºC for 20 minutes the following morning.

Initiation of Individual Chromoprotein and Regulator Plasmid Design

A standard procedure in the Tacoma RAINmakers project is PCR amplification. This process is listed under the protocol page as “Insert PCR Amplification.” In preparation for ligation of inserts (amilCP, spisPink, and PcArsR) into the vector, all insert DNA must be amplified from its original limited stock. Once the PCR reaction has exited the thermocycler, gel electrophoresis must be employed to assess the efficacy of the amplification. As pictured in Figure 3, both the spisPink and amilCP bands successfully appeared at about 1000bp, and the PcArsR expressed at about 550bp. Unfortunately, the PcArsR negative control produced DNA bands, which suggested contamination during the PCR amplification process.

Following a successful PCR amplification confirmed by the gel, Tacoma RAINmakers performed a standard gel extraction. With much more insert DNA, RAINmakers were prepared to design three additional plasmids containing single inserts for isolated testing.

Fig. 2. DNA gel of PCR products.

Week 4

Transformation of Arsenic Construct and PCR Screening

TThe positive control (pSB1C3 + Insert) and negative control (pSB1C3 only) were transformed using DH5 Alpha Competent E. coli cells. Although the iGEM protocol for transformation states that 1µL of DNA is sufficient, 2µL of DNA were used for both the positive and negative control plates to ensure enough DNA existed for multiple colonies to grow. Transformation reactions were incubated overnight at 37ºC within a 14-18 hour time period. An extended description of the standard transformation process is listed under the protocol page.
The RAINmakers ran a PCR screening for both the positive (vector and insert) and negative (vector only) controls. This PCR allows us to amplify our cloned DNA, which is necessary to do because it will help us determine whether or not the ligation of the vector and insert was successful. The product after our PCR will be used to run a gel, which will help us see which colonies have the vector and insert and which only have the vector. After determining the volumes of the reagents, we decided to add an extra 20% uncertainty to each reagent in the master mix since small volumes of liquid can get stuck in the pipette. We did all these calculations based on the fact that we planned on doing 8 PCR reactions (8 PCR tubes). Before beginning the process of adding all the reagents to our PCR tubes, we needed to determine how long the extension part of PCR should be based on the base pair count of our product. Running a PCR simulation of Snapgene allowed us to find the exact base pair count of our positive control, which is 1597 base pairs. From this, we found that the extension period should be 1 minute and 36 seconds.
After adding all of our reagents together into 8 PCR tubes, we needed to select single colonies from transformation plates from 6/19/18. We determined that 6 of our PCR tubes would have the positive control, 1 would be the negative control and the last tube would have no DNA. Once we selected a single colony and put the DNA into a PCR tube, we streaked it onto a new plate (cut up in 8 different sections for each 8 different colonies) and labeled them 1, 2, 3, 4, 5, 6 (positive control), 7 (nothing) and 8 (negative control). We then incubated this new plate so that more colonies would grow. Finally, we put our PCR tubes in a thermocycler and ran our PCR.

Blunt-End Digestion and Ligation of pSB1C3 and Inserts

Designing plasmids with individual inserts becomes slightly complicated, as each insert contains a SalI site that is not compatible with the BioBrick prefix/suffix of the backbone. Hence, Tacoma RAINmakers preferred blunt-end ligation, rendering the incompatible ends irrelevant.
Additionally, each insert was digested with its respective enzyme and filled in with T4 polymerase in preparation for blunt-end ligation.

Week 5 - Week 7

We re-started the cloning process with fresh PCR products and more vector. We re-ligated our pSB1C3 backbone with our PcArsR+SpisPink/PcArsR+amilCP insert several times with only negative results. We decided to re-ligate PcArsR + chromoprotein (spisPink or amilCP), as there were speculations that something went wrong in the initial ligation of the two. This ligation contained 3 uL of PcArsR, 3 uL of chromoprotein (spisPink or amilCP), 1x T4 DNA Ligase Buffer, 1.5U T4 DNA Ligase, and molecular water to a final volume of 10 uL. We ligated at 16ºC overnight, then heat killed at 80ºC for 20 minutes.

After this ligation was complete, we digested the newly ligated insert with XbaI and SpeI to get it ready for ligation with the pSB1C3 backbone. This digestion contained 1 uL of Buffer 2.1, 1 uL of XbaI, 1 uL of SpeI, and the newly ligated insert DNA. We did an overnight transformation and then ran PCR on the colonies. To see the results we ran it on a 1% agarose gel and saw that the insert did not ligate to the vector. This sequence of events repeated itself for many weeks, to great frustration.

Week 8

Restart cloning process

After failed attempts to clone the circuit, we decided to start again from the beginning. In the digestion of the backbone, we followed the iGEM protocol, but modified it by doing an overnight digestion at 16ºC instead of a 30 minute digestion. The same digestion was performed on PcArsR, spisPink, and AmilCP. We heat killed the ligase at 80ºC for 20 minutes the next day, and digested the new insert with XbaI and SpeI. After digestion, we did an overnight ligation of the vector and the insert using the slightly modified iGEM protocol with an overnight incubation.

Testing ligation success before transformation

The team spent a lot of time transforming without successful insertion of the circuit, and needed a better way to determine the quality of ligation product that was being put into the cells. We reasoned that since PCR can amplify the tiniest amounts of DNA, we may be able to use this to screen the ligation products.
To screen the ligation, we ran end point PCR on the newly ligated parts using our standard PCR protocol. Surprisingly, the PCR confirmed that the vector and insert successfully ligated together! We then transformed our full circuit using the transformation protocol on the iGEM website, and mini-prepped using the Promega Wizard Plus SV Kit.

Fig. 3. DNA gel of ligation showing successful ligation.

Week 9

Bioengineering Summer Camp

The team took a week off from lab work to run a summer bioengineering camp for 9th and 10th grade students. We had a lot of fun teaching the students and used many tools gained from iGEM, like letting students streak out bacteria containing RFP (positive transformation control) and GFP (interlab positive control).

Week 10

Circuit testing

We started the process to test the complete arsenic circuit. In vivo testing of full circuits with reporters amilCP and spisPink was performed with 1uM/2uM/10uM concentrations of arsenate/arsenite. RAIN's 2017 iGEM team had ran an experiment testing 150uM arsenic which killed the cells during the experiment. A second experiment was conducted where 25uM arsenic was used. This did not kill the cells and GFP was produced. This is the maximum amount of arsenic we want to test.

The limits of detection for our circuit to be viable and to compete with current testing equipment is below 100ppb. At 50ppb soil and water is considered unsafe for consumption or play. At 100ppb the soil or water is deemed worthy for excavation due to its potential health risks. Therefore, we will test 1uM arsenic (equivalent to 75ppb), 2uM arsenic (150ppb) and 10uM arsenic (750ppb).

10mL of liquid culture was created for both spisPink and amilCP the cultures will be set up as follows:

The cultures were observed at 1 hour, 3 hours, 24 hours and 48 hours for visual solution color change. During the entirety of the experiment, no color change or production occurred.

Repeat circuit testing with higher concentration of arsenite/arsenate

Testing of the amilCP and spisPink in vivo was unsuccessful at 1uM/2uM/10uM. There was also an attempt to grow CFU plates, however, these plates were left in the incubator for two days and overgrew. RAIN's 2017 iGEM team was able to see production of GFP at 25uM concentration of arsenic so this experiment will lie closer to these values.

Three concentrations or arsenic will be tested this time: 17.65uM (equivalent to 1324ppb), 21.42uM (1606ppb) and 25uM (1875ppb).
4mL of liquid culture was created for both spisPink and amilCP circuits.

After growing overnight, there were no color changes in the cultures.

Week 11

More cloning

Since we had a lot of trouble seeing any reporter expression, the team wanted to clone the individual parts of just the regulator and just the reporters. This will allow us to see if the reporter will turn on when its not being repressed with ArsR. Unfortunately, we ran into the same problems as before. By this, we mean that the ligation reactions were not demonstrating successful insertion of our DNA into the vector.

Week 12-13

Different members of the team kept trying to clone the individual parts. The individual parts were amplified using our standard PCR protocol. Following PCR amplification, the reactions were run at 120V on a 1% Agarose gel. After seeing in the light box that the PCR was successful, the PCR products were extracted using a standard protocol and stored at -20°C.

Fig. 4. DNA gel of PCR products.

The purified parts and pSB1C3 were digested overnight with XbaI and SpeI. The ligation was performed the next day using reactions contained 4.1μL of digested insert, 1.35μL digested linear pSB1C3, 1u T4 DNA ligase, and 1x T4 ligase buffer, and Molecular Water to a final volume of 10μL. It was incubated overnight at 16C. Then, the ligation was tested with PCR primers amplifying the region between the XbaI and SpeI sites on pSB1C3. The PCR reactions contained 1x GoTaq Flexi Green Buffer, 0.15mM DNTP, 1.5mM MgCl2, 0.63u GoTaq Polymerase, 0.5μL of Ligation Product, 0.38μM FWD primer, 0.38μM REV primer, and molecular water to a final volume of 10μL. We ran it on a 1% agarose gel and saw no bands of the desired size.

In order to test the amilCP reporter, we created a circuit with a high expression T7 promoter, amilCP, and the same double terminator used in our other circuits. We began cloning by digesting IDT stock solutions of T7/amilCP with EcoRI-HF and PstI, following iGEM standard protocols. We then ligated T7/amilCP with pSB1C3, using the iGEM ligation protocol. To screen the ligation we used our PCR protocol. After running PCR products on an agarose gel, we found that the ligation failed.

Week 14

We repeated the ligation at 4°C overnight. The reactions contained 5μL digested insert, 5μL digested linear pSB1C3, 1U T4 DNA Ligase, 1x T4 Ligase Buffer, and molecular water to a final volume of 15μL. The ligation was tested in the same way, and once again produced no bands.

The third attempt at ligation contained 10μL digested insert, 2.5μL of a plasmid with pSB1C3 that we digested using XbaI and SpeI, 1u T4 DNA Ligase, 1x T4 Ligase Buffer, and molecular water to a final volume of 20μL. It was left at room temperature for 1 hour, then moved to incubate at 16°C overnight. The ligation was tested in the same way as the previous 2 attempts. The gel had strong bands at 75bp, the size of the empty vector, and faint bands in the correct location for successful ligations.

Fig. 5. PCR of successful ligation. Extremely faint bands are visible in the 500-1000bp range of Samples 2, 3, and 4!

Transformations for the PArsR-spisPink construct were performed following the iGEM transformation protocol. The transformations were screened with colony PCR reactions containing 1x GoTaq Flexi Green Buffer, 0.15mM DNTP, 1.5mM MgCl2, 0.63u GoTaq Polymerase, 0.38μM FWD primer, 0.38μM REV primer, and molecular water to a final volume of 10μL.

Fig. 6. Colony PCR screening. Many colonies were screened to get just 2 positive colonies for the spisPink construct.

In continuation of the amilCP test circuit, we went back to the stocks and digested T7/amilCP with EcoRI-HF and PstI, again following iGEM protocols. After digesting, we ligated the product, this time with a ratio of backbone to insert at 5:1, in an effort to make the reaction more efficient. We screened with our standard PCR protocol and found that the ligation was a success (fig. 7.).

Fig. 7. T7/amilCP ligation screening. Sample 1 shows a band just under 1000bp, the correct size for ligated product.

Week 15

Now that one colony has been confirmed to contain the spisPink construct, we needed to transform the remaining ligations. Transformations followed the usual protocol, and colony screening by PCR.

With the ligation of the amilCP test circuit was complete, we proceeded to transform it according to iGEM protocols. Visual examination of the plates after incubation showed no signs of bacterial color change. We screened the colonies with our PCR protocol and found that the full circuit was present (fig. 8.). This likely means that there was a flaw in circuit design. Members of our team have speculated that this was due to the lack of spaces in the sequence around the promoters and ribosome binding sites.

Fig. 8. T7/amilCP transformation screening. Bands are visible just under 1000bp for Samples 2, 3, and 4. This corresponds to the size of T7/amilCP.

Week 16

After successfully transforming all our individual parts, we finally were getting ready to do minipreps for them. We used the Promega Wizard Plus SV Miniprep Kit and used the protocol provided by Promega to do our minipreps. The whole process did not take too long and soon after, we now had minipreps for pSB1C3/PcArsR, pSB1C3/PArsR-spisPink, and pSB1C3/PArsR-amilCP and stored them in the -20ºC freezer until it was time to send them to iGEM.