Team:uOttawa/Human Practices
Human Practices
Table of contents:
Summary:
Being an independent team of undergraduate students, we quickly learned that inexperience in synthetic biology and financial constraints limit the success of young teams. Thus, our project endeavoured to tackle these two obstacles. Thanks to our Virtual Protocol Donation Box, we analyzed, tested, and optimized a plethora of protocols from other successful iGEM teams. We compiled an easy-to-read protocol handbook designed specifically to assist new learners of synthetic biology. The handbook is equipped with software models designed by our dry lab to assist with primer design and yeast cloning. We allied with the University of Ottawa’s DEGREE program and invited students from various educational backgrounds and levels to test our protocols. We analyzed their performance and their feedback to ameliorate our protocols. Based on their comments, we filmed succinct videos to accompany each procedure. Our protocols are specifically designed to train amateurs of synthetic biology using cost-effective techniques. Our collaborators, who worked with higher order eukaryotes, recommended that we make the competition accessible to young synthetic biologists who are interested in working with eukaryotes. Consequently, we developed a library of BioBrick plasmid backbones that are simultaneously compatible for cloning in both E. coli and S. cerevisiae using the basic techniques explained in our protocols. To further overcome the financial hurdles of the competition while keeping up-to-date with modern cloning techniques—and to address common documented issues with traditional cloning that were noted by our collaborators and during the making of our yeast-compatible plasmid backbone library—, we developed a DIY Gibson Assembly kit to minimize the cost of cloning and provided an easy-to-read manual to make the kit. We provide the necessary enzyme sequences, cost-free, to the Registry of Standard Biological Parts.
1. The Handbook of Tested and Optimized Synthetic Biology Protocols for Amateurs
a. Background: Science, Finances, Politics
Technical complications of synthetic biology, in addition to financial constraints, are especially problematic for young scientists. We can speak from experience: iGEM is the first opportunity for all our team members to work in synthetic biology, and the learning curve was very steep. Not only we started the competition with hardly any knowledge of restriction enzymes, we had no funds to finance the team. To make matters worse, a new political party took power in our province and insisted on making our science life extra-difficult by cutting-down public education funds, which ultimately prevented our access to funding also. Although our prospects seemed bleak, given our inexperience and financial standing, we had a vision: make synthetic biology accessible to amateurs through simple, tested, optimized, and cost-effective training protocols. Our idea expanded to more than just a protocol handbook as we progressed through the project—continue reading for more details.
b. Virtual Protocol Donation Box
Thus, began a period of intense protocol review. We collected a plethora of tested protocols from a variety of sources. The first source was iGEM’s website. These protocols inspired us because they provided a brief introduction to explain the purpose of each procedure. Upon testing the protocols, however, we noticed that the protocols consistently recommended using less DNA, which assumed that the experimenter had a high experimental success rate, which was not our case. For example, in iGEM’s DNA extraction from a kit plate protocol, it recommends that only 1 µL of immersed DNA be transformed into home-made chemically competent E. coli. We found that volume to be very little and we had very limited growth on our antibiotic selection plate. Instead, we modified that protocol to indicate that all immersed DNA be used for the transformation. This enhanced growth on antibiotic selection agar plates.
c. University of Ottawa DEGREE Program
As explained above, we developed a set of robust training protocols that could be easily understood and performed by undergraduate students with limited research experience.
The training protocol was designed to replicate the workflow of developing new BioBricks. It included procedures such as DNA restriction digestion, DNA ligation, E. coli transformation, E. coli inoculation, E. coli colony PCR, plasmid DNA miniprep, and gel electrophoresis.
As a part of our initiative to get more students interested in synthetic biology and to make synthetic biology more accessible, we collaborated with the University of Ottawa to host 4 students from the DEGREE (Discover and Engage Graduate Research Experience) program for three days. This program aimed to introduce undergraduate students to research performed at the University. These students, who had limited research experience, were provided the opportunity to perform our training protocols.
All students, with the exception of one student, had at least one red colony present on their plate. We inoculated 3 colonies (or the maximum number of red colonies) from each individual’s plate in liquid LB kanamycin. For the student who did not have any red colonies, she continued her experiments using a backup plate that was prepared when the protocol was being tested. Each individual also performed a colony PCR of RFP. The results of the colony PCR can be seen in the following gels:
The gels show a single band around the 1000 base pair mark, which represents RFP. There was only one student (student A) whose colony PCR did not work. Given that her positive control also did not work, the PCR failure can likely be attributed to improper technique. We also ran everyone’s initial digestion product on a gel to assess whether or not the DNA was completely digested:
The gel shows two bands at the correct length in every lane: the upper band represents PSB1K3, which is 2204 base pairs long. The lower band represents RFP, which is 1069 base pairs long. Two lanes show additional bands of different lengths, but the desired two bands are still present. These additional bands can be attributed to contamination. After the program, we sent each student a questionnaire to ask for feedback on our protocols.
d. Refined Protocol Handbook with Protocol Videos and Primer Designer
The feedback from the DEGREE visitors who tested our protocols was very helpful, and we used it to refine the protocols and make them clearer. For example, one student stated that although the protocols seemed somewhat abstract at first, they made sense once we put them to practice in the lab. Because someone with experience cannot always be present to provide assistance to students who are trying out the protocols, this gave us the idea to film video protocols to accompany our written protocols. We also noted that students struggled with the concept of primer synthesis. Therefore, Miléna Sokolowski, the dry lab member, designed a tool to assist with the making of the PCR primers. Please visit her software/dry lab page to learn more.
The refined Handbook of Tested and Optimized Synthetic Biology Protocols for Amateurscan be found here.
The series of video protocols can be found on our YouTube channel.
You can also download our protocol videos directly from iGEM's server:
-Pipetting Protocol
-Restriction Digestion Protocol
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2. Rapid, Flexible, and Affordable Yeast Genome Engineering with BioBrick Standardization
a. Background: Facilitating iGEMer’s Access to S. cerevisiae Cloning
While rummaging through iGEM’s websites in search for protocols, we noticed that the BioBrick backbones were limited to work with E. coli. There were very few resources for amateur synthetic biologists interested in working with higher-order eukaryotes. This could be due to a multitude of reasons, namely that cloning in eukaryotes is technically more difficult, time-consuming, and costly. Because ourHandbook of Tested and Optimized Synthetic Biology Protocols for Amateursalso encompassed transformations in Saccharomyces cerevisiae, we believed that we should also work on making the competition more accessible to synthetic biologists interested in working with Saccharomyces cerevisiae. We felt doubly encouraged by Team UC Davis, who were also tackling the issue of the absence of eukaryotic representation in iGEM, save in mammalian cell lines. In brief, we develop a library of BioBrick plasmid backbones that are compatible for cloning in both E. coli and S. cerevisiae using the basic techniques explained in our protocols. The plasmid backbones adhere to the RFC 10 Standard and Type IIS Assembly and allow for the systematic and efficient cloning of a desired gene within target yeast chromosomal loci, Ade2, His3, Ade4, and Gal4 loci, via homologous recombination and are equipped with KanMX, NatMX, Ura3, and His3 yeast-selectable markers as well as RFP to enable colorimetric selection in E. coli. This library reduces the number of steps, and the amount of troubleshooting, preceding a yeast transformation. Setti Belhouari spearheaded this project by designing and constructing the RFC Standard 10 backbones, based on pSB1K3, that are compatible with homologous recombination in S. cerevisiae as part of her honours thesis project in the fourth year of her undergraduate studies. Setti Belhouari and Taylor Lanosky also expanded the plasmids by designing them such that they are also compatible with Type IIS Assembly. Taylor Lanosky constructed the remainder of the Type IIS Assembly yeast-compatible plasmid backbones.
b. RFC 10 Standard Yeast-Compatible Plasmid Backbones
In addition to complying with the BioBrickTM standard specifications, which enables the usage of the platform in the classic BioBrick Standard Assembly process, our goal was to update and design BioBrick plasmid backbones to enhance site-directed genomic modification via homologous recombination in S. cerevisiae, the most efficient method of genome modification in this organism. To do so, BioBrick backbone, pSB1K3, needed to include 1) upstream and downstream sequences that are homologous to various yeast loci allowing for homologous recombination, 2) an auxotrophic or an antibiotic selection marker that allows the identification of the yeast clones possessing the desired genome modification, and 3) a multiple cloning site that allowed the insertion of a desired coding sequence (Figure 13).
In order to comply with the BioBrick Standard, we chose to improve an existent BioBrick backbone. pSB1K3. Although the ultimate goal of the plasmid is to facilitate cloning in Saccharomyces cerevisiae, initial updating of the plasmid in E. coli facilitates manipulation and cloning of the plasmid as well as the cryopreservation of the E. coli strains and their submission to the Registry of Standard Biological Parts to allow other research groups to benefit from our work. Furthermore, any subsequent modifications which we perform on the backbone do not introduce any illegal restriction sites that are found in the ‘prefix’ (EcoRI, XbaI, and NotI) and the ‘suffix’ (SpeI, NotI, and PstI).
Because the process of homologous recombination in yeast results in the replacement of a pre-existing gene with a heterologous DNA insert, we chose to design our plasmids such that they each contain the upstream and downstream regions homologous to either the His3, Ade2, Ade4, and Gal4 loci, because they are non-essential genes whose absence does not result in cell death when the host is supplanted with the appropriate media. Furthermore, Ade2 and Ade4 loci are common targets for direct chromosomal modification because they provide colorimetric selection. The disruption of the Ade2 locus results in the formation of red colonies due to the accumulation of the red pigment P-ribosyl aminoimidazole, which facilitates screening of the desired clones in S. cerevisiae. Subsequent targeting of the Ade4 locus, on the other hand, results in the elimination of the red pigment, which again helps identify the desired new clone which are now white colonies.
As indicated in Figure 15, we carefully selected the upstream and downstream homology sequences to consist each of 100 bp and a high GC content, to optimize yeast homologous recombination at the target locus, while avoiding the inclusion of restriction digestion sites that are found in the multiple cloning site. More importantly, the upstream homology sequences consist of 100 consecutive base pairs of the loci’s respective promoters that come after the upstream transcription activating sequences, such as TATA boxes. This ensures that the upstream activating sequence in the endogenous target locus is undisrupted after homologous recombination, allowing for the formation of the transcription complex necessary for the transcription of the desired insert. The downstream homologous region, on the other hand, contains 100 bp directly following the stop codon of the endogenous yeast locus, which consists primarily of the transcription termination sequence. This insulates the desired DNA insert from any downstream coding sequences found in the chromosome in which it is inserted after yeast transformation.
Global overview of the construction of the plasmid library:
To complete Step 1 (Figure 17), Setti PCR amplified the homology sequences to include two additional restriction sites: an MfeI and an NsiI cut site at the 5’ and the 3’ ends, respectively. Then, she digested the amplicon with MfeI and NsiI. On the other hand, she digested a pSB1K3 plasmid containing RFP between the prefix and the suffix using EcoRI and PstI. Because the MfeI and the NsiI protruding ends are complementary to EcoRI and PstI protruding ends, respectively, the upstream and downstream homologies can be ligated to pSB1K3 using T4 ligase, without reforming the restriction sites. This resulted in the elimination of the RFP insert and the growth of white colonies of LB kanamycin. She screened 2, 3, 3, and 4 white colonies for the His3, Ade2, Gal4, and Ade4 upstream and downstream homologies in pSB1K3, respectively, and found that all colonies tested positive for the desired homology sequences (Figure 18)
To achieve Step 2, which consisted of adding a yeast-selectable marker between the prefix and suffix, a colony PCR reaction, Setti amplified a yeast kanamycin (KanMX) selection maker from a pre-existent yeast strain expressing the marker. This PCR reaction allowed her to flank the KanMX selection marker with an MfeI restriction site on the 5’ end, as well as the prefix and the suffix on the 3’ end (Figure 19). She digested the amplicon with MfeI and PstI, whereas she digested each of the four homology-containing plasmids (Ade2, His3, Ade4, and Gal4) with EcoRI and PstI. The complementary annealing of the MfeI protruding end and the EcoRI protruding end, as well as those of PstI, results in the ligation of the selection marker between the upstream homology and the prefix of the homology-containing plasmids. The ligation restores the prefix and the suffix and does not produce a restriction-site where the MfeI cut-site anneals with the EcoRI cut-site. She transformed the ligate in E. coli and grew it on LB kanamycin. (E. coli expresses resistance to kanamycin due KanR in the pSB1K3 backbone, not due to the KanMX insert between the upstream homology and the prefix. KanMX confers resistance to G418 or Geneticin® in S. cerevisiae.)
Anticipating a similar high efficiency ligation as the one during which which she ligated the homology sequences to pSB1K3, Setti screened 3 colonies in each search for Ade2::KanMX, His3:KanMX, Ade4::KanMX, and Gal4::KanMX. (In this notation, the value appearing before colon represents that target locus homology flanking the value after the colon. For example, Ade2::KanMX refers to the plasmid, in Figure 10, containing the Ade2 target locus homology sequences flanking the KanMX selection marker and the multiple cloning site.) Because none of the 12 colonies screened contained the desired KanMX marker, she performed a 5’ dephosphorylation of the digested homology-containing plasmids to prevent the plasmids from re-circularizing before they anneal to the digested KanMX amplicon. Only 1 out of the 45 screened colonies for the Ade2-homology-containing plasmid across 3 replicates contained the KanMX marker (Figure 20). Thirty-six colonies were screened for His3::KanMX, 27 for Ade4::KanMX, and 27 for Gal4::KanMX, but with no success.
A primary suspect for the decrease in ligation efficiency was ethanol contamination. Because the restriction endonucleases used, MfeI-HF and PstI-HF, are not heat-labile, digested DNA was column purified to eliminate the restriction enzymes prior to ligation. As indicated in Figure 21, following the column purification of the digested homology sequence and pSB1K3 (prior to the ligation that yielded 100% efficiency), the average 260 nm to 280 nm absorbance ratio was 1.88 ± 0.10, indicating pure DNA. However, following the column purification of the digested KanMX selection marker and the homology-containing pSB1K3, the average absorbance ratio increased to 2.63 ± 0.62, indicating contamination. Thus, BioBasic’s column purification protocol was optimized to include a 2-hour evaporation of ethanol prior to the elution of the digested DNA. Consequently, the absorbance ratio decreased to an average of 1.86 ± 0.09, indicating pure DNA.
Nevertheless, evaporation of the ethanol did not significantly increase the yield of ligation: out of the 28 screened colonies across three replicates, only one presented the Ura3 auxotrophic selection marker in the Ade4-homology containing backbone Figure 22, despite using 10 times more Ura3 to Ade4-homology-containing backbone. (During the ligation of the KanMX to the Ade2-homology-containing backbone, a molar ratio of 4 to 1 was used.)
To allow for direct comparison and to determine the effect of ethanol on the ligation of the selection markers to the homology-containing backbones, three ligation experiments were performed in parallel. In the first experiment, the KanMX selection marker and the Ade2-homology-containing backbone were directly ligated following digestion without column purification. In the second experiment, the selection marker and the backbone were digested, column purified with a 2-hour ethanol evaporation, and then ligated. The third experiment was identical to the second experiment, except that the eluted digested DNA was further vaporized overnight at 37°C and then re-suspended in distilled, de-ionized water prior to ligation. The first experiment yielded significantly more growth than the second and the third experiment (Figure 23). Five colonies were screened per plate for a total of 15 colonies screened per experiment. Plasmids were extracted via miniprep extraction, digested with PmeI, and then gel electrophoresed. The only colony which contained the successful ligation of the KanMX selection marker to the Ade2-homology-containig pSB1K3 was the result of the first experiment (Figure 24). Nevertheless, the result was not significant enough to demonstrate that the elimination of ethanol contamination enhanced ligation.
Having eliminated the problem of ethanol contamination (by optimizing BioBasic’s column purification protocol), the ligation of the selection marker to the homology-containing backbone was almost identical to the ligation of the homology sequence to the pSB1K3 backbone. Nevertheless, the overall average efficiency of the ligation dropped from 100% to a negligible value (0.72 ± 2.74%) from the former to the latter ligation, respectively. One difference remained, however: During the former ligation, the homology sequences were replacing RFP. Following the transformation, only white colonies were screened in a mixture of white and red colonies. However, during the subsequent ligation of the selection markers to the homology-containing backbones, colorimetric selection was not possible (compare Figure 17 and Figure 19). To determine if the absence of RFP resulted in the increase in selection of false positives, RFP was introduced between the prefix and the suffix of all homology-containing plasmids (Ade2, His3, Ade4, and Gal4). Both plasmids and RFP were digested with EcoRI and SpeI prior to ligation. Interestingly, because both EcoRI and SpeI are heat-labile, column purification was not necessary, and ligation directly followed digestion, resulting in an average ligation efficiency of 51.1 ± 9.8%. This demonstrates that the elimination of a column purification step — given that the restriction endonucleases could still be heat-inactivated prior to ligation — significantly increases ligation efficiency (second column to the right column in Figure 25).
RFP colorimetric selection, however, did not reduce the prevalence of false-positives (Figure 26). In a procedure similar to that illustrated in Figure 19, Ade2::RFP plasmid was digested with EcoRI and PstI, while the KanMX amplicon was digested with MfeI and PstI. The digested DNA was column purified, then ligated and transformed. A total of twelve white colonies were screened across three plates with their plasmids extracted and digested with PmeI to identify the KanMX insert. None proved positive.
In an effort to compare the purported traditional cloning approach to modern cloning (Figure 19), the KanMX selection marker was ligated to the Ade2-homology-containing backbone through Gibson Assembly®. Gibson Assembly’s recommended protocol was followed66: Both the backbone and the selection marker were PCR amplified with 20 bp homologous ends. Both amplicons were column purified and then incubated in Gibson Assembly Master Mix at 50°C for 15 minutes. When the backbone was PCR amplified from the Ade2::RFP plasmid, none of the twelve screened colonies across three plates incorporated the desired Ade2::KanMX in pSB1K3. On the other hand, when the backbone was PCR amplified from the RFP-less Ade2-homology-containing plasmid, only one colony, out of twelve across three plates, incorporated the Ade2::KanMX in pSB1K3 (Figure 25). Thus, again, the use of RFP colorimetric selection did not reduce selection of false-positives. Gibson Assembly did not offer a significant advantage to traditional cloning, but it reduced the number of colonies screened (12 versus 45).
Because Gibson Assembly® required less screening for the identification of positive clones, this cloning method was used for the creation of the His3::NatMX and the Gal4::His3 pSB1K3 plasmids, following New England BioLab’s recommended procedure, as described above. Out of 10 colonies screened, across three plates, 2 contained the desired His3::NatMX plasmid in pSB1K3, for an average ligation efficiency of 16.67 ± 14.43 %. Similarly, out of the 12 colonies screened, across three plates, only 1 contained the desired Gal4::His3 pSB1K3 plasmid (Figure 26). Because we found Gibson Assembly to be quite costly for our modest team, we also endeavoured to make our very own DIY Gibson Assembly kit, spearheaded by George Liu (see below).
Emily Lam courageously tried to find an alternative to column purification to see if she can improve the yield of our ligations. Such an alternative would greatly facilitate the expansion of the plasmid library. After some research, she found MunI, which is an isoschizomer of MfeI supplied by ThermoFisher. She tried assembling the plasmid using the same process as explained above in which MunI replaced MfeI, but this did not increase efficiency (Figure 27).
Emily also tried inserting RFP in between the BioBrick prefix and suffix to add colorimetric selection. After ligating KanMX, there were many white colonies. However, none of them contained KanMX after digesting their plasmid miniprep with pmeI, resulting in a gel similar to that of Figure 27.
At this point, we began to believe that the poor efficiency may be a result of a problem with the protocol. Thus, Emily repeated the protocol using RFP instead of KanMX. RFP was incorporated into the homology-containing backbones and transformed onto 3 plates. Approximately 10% of the colonies were red, which indicated that our approach had an approximate efficiency of 10%. Because our ligation efficiency without using isoschizomers was far higher (up to 65%), this suggests that there is something inherently disadvantageous about the annealing of two cohesive ends resulting from the isoschizomer digest.
Emily also tried adding the yeast-selectable marker between the upstream homology and the prefix using Type IIS assembly. We created primers for KanMX and RFP containing BsaI recognition sites, which would cut in a manner that created EcoRI and SpeI-compatible sticky ends. Approximately 97% of the colonies on the RFP plate were red, and they were confirmed after digesting the minipreps with PmeI (Figure 28 A). On the other hand, 10 colonies from the KanMX plate were tested, and none of them contained the KanMX segment (Figure 28B). Thus, although we succeeded in creating our yeast-homology platform, further troubleshooting of the traditional endonuclease approach failed to increase efficiency.
We contented ourselves with the library that we had built thus far and decided to expand the plasmid backbones that we made to make the compatible with Type IIS Assembly. Both Setti Belhouari and Taylor Lanosky designed the plasmid backbones. Taylor Lanosky constructed the remaining Type IIS plasmid backbones discussed in the section below.
c. RFC 10 Compatible Standard and Type IIS Yeast-Compatible Plasmid Backbones
Firstly, we looked at the iGEM website requirements for RFC 1000 (Type IIS compatibility) and realized that BsaI sites were necessary to incorporate in both our inserts and backbones. The easiest way to do this was to use PCR amplification. Although each plasmid is unique, the same general steps were followed during the creation of each one (see the Type IIS protocol for a more detailed explanation):
1. Isolate the insert from a backbone which is different from the backbone that will be in the plasmid we are trying to create (ie. RFP from RFP pSB1C3 if creating an RFP pSB1K00 plasmid)
2. Isolate the backbone from an insert which is different from the insert that will be in the plasmid we are trying to create (ie. pSB1K3 from CEN9 pSB1K3 if create an RFP pSB1K00 plasmid)
3. Create primers that will allow for the addition of BsaI sites within the promoter prefix and terminator suffix.
4. PCR insert and backbone
5. Ligate insert and backbone together
Note that we wanted to make sure that we took RFP from a backbone other then pSB1K3 and removed pSB1K3 from an insert other than RFP to ensure that if we had red colonies following the transformation, we knew that it was a product that we made rather than one that was pre-existing.
During the creation of these primers, we came across some challenges. Firstly, we knew that we wanted our plasmids to remain RFC 10 compatible so that anybody working with them could use either Type IIs assembly or standard assembly. The easiest way to ensure this happened was to put the BsaI sites outside of the RFC 10 prefix and suffix. Next, we wanted to be certain that the Type IIS (BsaI) cut sites remained present after ligation so that if we wanted to use them for basic Type IIS assembly or even more advanced Type IIS assembly (ie loop assembly) in the future, we could. Below are images of our primer sets (Figures 29 and 30): you will notice that the BsaI cut sites are inverted such that they remain part of the backbone even after digestion. Also, they are comprised of “junk” DNA (4-5 base pairs downstream of the recognition sequence) to allow for annealing with RFP.
After Taylor PCR’d both the insert and backbone with their respective sets of updated primers, we gel’d to verify that our products were the correct length. We knew that pSB1K00 (pSB1K3-Type IIS compatible) should have been 2238 base pairs and that RFP-Type IIS compatible should have been 1103 base pairs. Below are images of gel of the verified PCR products (Figure 31). Note that we use Thermofisher’s 1 kb Plus DNA ladder.
Once the PCR product was verified, a PCR purification was done and the concentration and purity was recorded using Thermofisher’s nanodrop machine. Taylor then digested both the insert and backbone with BsaI, heat shocked the digest, ligated the digested insert and backbone together, and transformed the entire reaction mixture. Note that the detailed protocol can be found under the protocol Type IIS Compatibility-For Basic Parts. A verification of 13 colonies was done to ensure that we had created the desired plasmid. Of the colonies screened, 100% were verified. Note that the transformation itself was 97% efficient. Images of the gel verification can be found below (Figure 32).
After verifying that our new protocol is efficient, we decided to use it to create a library of plasmids which users would be able to utilize to synthesize samples in multiple assembly formats. We wanted to build plasmids that were very useful and versatile, so we chose to work with our already-existing plasmids which contain yeast homologies. If we could develop these, then we would have a platform that was both RFC 10 and RFC 1000 compatible, and due to the fact that they have yeast homologies, compatible with both Saccharomyces cerevisiae and Escherichia coli as well.
We had the following 8 plasmids with yeast homologies; Ade2 Homologies pSB1K3, Ade2::KanMX-pSB1K3, Ade4::Ura3-pSB1K3, Gal4::His3-pSB1K3, Ade4 Homologies pSB1K3, Gal4 Homologies pSB1K3, His3 Homologies pSB1K3, His3::NatMX-pSB1K3. We designed a unique set of primers for each one to add the promoter prefix and the terminator suffix (as we previously did for pSB1K3 above). Note that besides what we have added with primers (consisting of junk and the terminator suffix/promoter prefix) the rest of the plasmid was already made. Below are images of the primers we have constructed and the resulting plasmid with that the primers would be adding included. Note that “EXSP” refers to the prefix and suffix adjacent to one another.
Once the primers in, Taylor followed the same process of PCR’ing and gelling to verify that we had the correct lengths.
Note that from this time on we will refer to pSB1K00 and not pSB1K3 due to the fact that they are now Type IIS compatible.
Taylor chose to use our yeast homology linearized plasmids (now Type IIS compatible) as the “backbone”, and the Type IIS compatible RFP we originally created as the insert. This choice was beneficial because it saved us time from having to make another insert Type IIS compatible and we could use the colorimetric selection it would provide to our benefit. We then followed the same protocol as we did to make RFP pSB1K00 a Type IIS compatible plasmid for each of the 7 new backbones. There was plenty of growth from each transformation with a majority of red colonies. Due to the high success rate of our first attempt at creating Type IIS compatibility, we decided to screen only 1 red colony from each transformation. Plasmids were extracted via miniprep and then digested with BsaI. The lower band near 1000 bp corresponds to RFP, whereas the higher band in the non ladder column represents the yeast-compatible plasmid backbone as represented in Figures 41 and 42.
In summary, our team has created an efficient protocol with the purpose of building Type IIS compatible basic parts and used this protocol to construct a library of Type IIS and RFC 10 compatible plasmids that permitted cloning in both E. coli and S. cerevisiae. Please see below the list of plasmids/plasmid backbones made to date.
1. Ade2::EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271000
2. Ade2::EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271008
3. Ade2::KanMX-EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271004
4. Ade2::KanMX-EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271012
5. His3::EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271002
6. His3::EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271010
7. His3::NatMX-EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271007
8. His3::NatMX-EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271015
9. Gal4::EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271003
10. Gal4::EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271011
11. Gal4::His3-EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271006
12. Gal4::His3-EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271016
13. Ade4::EXSP pSB1K3(RFC10 backbone only) – BBa_K3271001
14. Ade4::EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271009
15. Ade4::Ura3-EXSP pSB1K3 (RFC10 backbone only) – BBa_K3271005
16. Ade4::Ura3-EX-RFP-SP pSB1K3 (RFC10 backbone only) – BBa_K3271014
17. Ade2::EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271017
18. Ade2::KanMX-EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271012
19. His3::EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271019
20. His3::NatMX-EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271023
21. Gal4::EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271020
22. Gal4::His3-EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271024
23. Ade4::EX-RFP-SP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271018
24. Ade4::Ura3-EXSP pSB1K00 (RFC10 and Type IIS backbone) – BBa_K3271022
25. RFP pSB1K00 (RFC10 and Type IIS backbone) --BBa_K3271031
The final step in the creation of the plasmid library consisted of testing and validating that the plasmids can indeed perform homologous recombination in S. cerevisiae.
d. Testing and validating the plasmid library.
Because the plasmids in the library were made by cloning in E. coli, we had already validated that they are compatible with cloning in E. coli. Next, they had to be validated for cloning in S. cerevisiae. We chose the 4 above underlined plasmids. By confirming that they perform homologous recombination in yeast, we would have also validated all the plasmids in the library because we would have proved that the homology sequences facilitate the process of homologous recombination, whereas the selection markers are functional. Both Setti Belhouari and Victoria Feng validated the plasmid library. Setti Belhouari, who had previous experience cloning in S. cerevisiae, demonstrated that the plasmids, do indeed permit homologous recombination in yeast. Victoria Feng, who had no previous synthetic biology experience, demonstrated that the plasmids indeed facilitate cloning in S. cerevisiae.
Setti PCR amplified a yeGFP cassette (pGPD-yeGFP-tCYC1, BBa_K3271029) flanked by a prefix and a suffix was digested with NotI and directly ligated within the prefix and the suffix of each of the NotI digested plasmids (following the heat-inactivation of NotI endonuclease). The average ligation efficiency of yeGFP into all four plasmids was 36.54 ± 17.06% (see centre-most column in Figure 25). Figure 43 shows the gel electrophoresis of the PmeI digested plasmids from 13 screened colonies following E. coli transformation of aspired ligated Ade4::Ura3-yeGFP in pSB1K3. Presence of yeGFP within the target-loci homologies of the plasmids was further confirmed by PCR using primers complementary to the upstream and downstream homologies as well as primers complementary to the promoter and terminator of yeGFP.
The yeGFP cassette in the Ade2, Ade4, and Gal4-targeting plasmids (Ade2::KanMX-yeGFP, Ade4::Ura3-yeGFP, and Gal4::His3-yeGFP) was PCR amplified, then column purified, and then transformed in S. cerevisiae in duplicate, resulting in an average of 18.5 ± 2.1, 122.7 ± 39.5, and 0.7 ± 0.6 colonies per plate, respectively. In a preliminary assay, patches of the Ade2::KanMX-yeGFP transformation were subjected to Alexa 488 protocol, causing an average of 15.0 ± 7.1 % to fluoresce (Figure 44A). A colony PCR was conducted on a seemingly fluorescing colony to ensure the insertion of the yeGFP cassette in the Ade2 locus (Figure 44B).
Victoria repeated the above process independently. She replaced RFP with the yeGFP cassette in the underlined plasmids from the above section. She digested the plasmids using PmeI and gel electrophoresed them, resulting in Figure 45. She then transformed the DNA using the Handbook of Tested and Optimized Synthetic Biology Protocols for Amateurs into S. cerevisiae, and obtained the abundant growth depicted in Figure 46. Therefore, our plasmid backbone library indeed facilitates cloning in S. cerevisiae while remaining compatible with cloning in E. coli via RFC 10 and Type IIS Assembly.
3. Frugally Modernizing Synthetic Biology Techniques
a. Background: When Traditional Techniques Fail
As explained in section 2b, the use of traditional endonuclease approach for the building of the plasmid library was error-prone for nor foreseeable cause. The use of Gibson Assembly salvaged the process. However, Gibson Assembly is more costly than traditional endonucleases. Therefore, rather than purchasing it, we decided to make our own homemade DIY Gibson Assembly Kit and provide a manual for the making of the kit. We assumed that other teams would be facing similar challenges, so we endeavoured to share all elements of our kit with other iGEM teams through the Registry of Standard Biological Parts. George Liu spearheaded the making of the kit.
b. DIY Gibson Assembly Kit
To promote the sustainability and longevity of iGEM uOttawa, we set out to create cheaper, alternative reagents for future and ongoing projects. Our main target this summer was to isolate and purify a set of “homemade” enzymes: Phusion polymerase (PHU), Taq ligase (TAQ), and T5 exonuclease (T5X).
By consulting the New England Biolabs and Thermo Fisher Scientific websites, George identified the source organisms of each commercial enzyme. He was able to locate and identify the DNA sequence for PHU from its US patent, but the other enzymes proved to be more difficult. After cross-referencing several academic papers and journals, George located the amino acid sequences for TAQ and T5X on UniProt. Using IDT software, we converted the amino acids into DNA sequences optimized for expression in E. coli. In silico, we added NcoI and SalI restriction sites to each coding sequence. Finally, we designed primers and ordered our sequences from IDT.
Upon receiving the synthesized DNA, George performed PCR to amplify the DNA samples. The results are shown below.
Using restriction digestion (NcoI + SalI in NEBuffer 3.1) and T4 DNA ligase, George inserted each coding sequence into a pHis-Parallel 2 expression vector. He transformed the resultant plasmids into DH5α E. coli, plated on LB-Amp, and grew them overnight. He then randomly selected colonies for miniprep, digested the purified DNA (NcoI + SalI in NEBuffer 3.1), then used gel electrophoresis to screen for successful inserts. (Figures 49 and 50).
The initial screening showed very faint bands for T5X (likely due to the expression and toxicity of the enzyme), so we performed PCR again on the ligated plasmids using primers for the T5X insert. The results are shown below:
From the successful colonies, George transformed the verified plasmids into a Rosetta E. coli strain, which he plated on LB with ampicillin (Amp) and chloramphenicol (Cm). He amassed 10 colonies and transferred them to a liquid growth medium with 100mL of liquid LB (Amp+Cm), letting it grow on a shaker at 37°C until the OD reached between 0.6-0.8. George transferred a 10mL aliquot to each of 3 larger flasks containing 1L of 2xTY liquid media. He grew the cultures on a shaker at 37°C until the OD600 reached 0.1-0.3, then shifted the cultures to the 25°C shaker. He let the cultures grow until the OD600 reached 0.6-0.8.
George made a sample pellet from 3mL of the uninduced sample (freezing immediately in liquid nitrogen), then added IPTG (at 0.25mM) to each flask to induce expression. He returned the flasks to the 25°C shaker and let the proteins express for 3 hours. Afterwards, George made another pellet from 1.5mL of the induced sample.
The pellet for T5X compared to Ptp1, another protein Dr. Rudner was studying, is shown below.
To test for effective induction, George lysed the samples by bead beating them in 250 µL of 1X Sample Buffer (50mM Tris 8.0, 300mM NaCl, 10mM imidazole, 10% glycerol) + 1mM PMSF. George ran 15µl of each sample on a 15% polyacrylamide gel and used Coomassie staining to assess induction. The results are seen below:
Dr. Adam Rudner took charge of expressing and purifying HF Phusion Polymerase. We tested the functionality of the Phusion Polymerase throughout the entire project. The polymerase can amplify DNA of different lengths and at different elongation temperatures, as seen in the gel electrophoresis figures above.
Taq ligase (BBa_K3271026), T5 exonuclease (BBa_K3271025), and HF Phusion Polymerase (BBa_K3271027) have been submitted on the Registry of Standard Biological Parts.
The next step in the preparation of the kit would be to purify TAQ and T5X, then independently test each enzyme’s catalytic function. The manual for the preparation of the DIY Gibson Assembly Kit has been provided to assist other iGEMers in the making of the kit. Note: this manual still includes the purification steps because they have been successfully implemented by Dr. Rudner in the making of the homemade polymerase. The manual also includes a list of reagents needed to assemble the final Gibson Assembly Master Mix buffer.
The success of the homemade Phusion has already been extraordinarily effective in cost reduction. Our lab has high hopes that the other two enzymes work just as well, and we hope to test our DIY Gibson Assembly kit by year’s end.
If that works out, it would be a huge step towards making synthetic biology accessible for everyone! And who knows what’s next? Restriction enzymes? Transcription factors? When you can do it yourself, the only limit is the extent of your own imagination!