Wet Lab
To create the MicroMurals project, wet lab’s goals have been split into 3 parts: recreate chromoprotein expression from Team Uppsala’s 2012 and 2013 teams, enhance color intensity, and create bio-ink. Part 1 will be completed when we see color and use spectroscopy to confirm. Part 2 will have worked when transformed colonies with the auxochrome treatment are visibly seen to be more vibrant than without the auxochrome treatment. Part 3 consists of having these chromoproteins fused to csg (curli nanofiber) monomers and chemically crosslinked into a hydrogel to create the bio-ink needed for the bioprinters that our product development subteam created.Part 1: Recreate Chromoprotein Expression
Cycle 1:
Design: We will use chromoprotein parts created by Team Upsalla to express chromoproteins in E. Coli. In this first cycle, we attempted to transform parts 7J (BBa_K1033922, meffRed), 9L (BBa_K1033930, amilCP blue/purple), and 9N (BBa_K1033931, amilGFP yellow) from plate 6 of the 2021 DNA distribution kit into HMS174 (DE3) chemically competent cells. We also tested cells that were transformed in the previous cycle to see if they were still viable. Since chromoproteins are visible to the naked eye, the transformation will have worked when we see color qualitatively.
Build: As Team Cornell was sent the plasmids for the chromoproteins via the distribution kit, the first build process is simply to successfully transform the plasmid into E. Coli and view color expression. If testing goes correctly, the part 2 build will be planned to create the plasmids needed to enhance color.
Test: To test the chromoproteins, the parts that were detailed above from the iGEM 2021 DNA distribution kit were used within a transformation on plates with chloramphenicol antibiotic. After 48 hours, colonies grew but no color could be seen. Furthermore, the colonies grew abnormally with highly concentrated sections of cells and contamination.
Learn: After viewing the transformation, Team Cornell reviewed what happened during the transformation and came to conclusions that lead to a new design.
- Glass Beads used for spread plates were not sterile leading to contamination
- Petri dishes were also not sterile and could cause contamination
- Chloramphenicol antibiotic was made using water which later was learned to be insoluble allowing for non-transformed colonies to grow
Cycle 2: Stopping Contamination Issues
Redesigning Cycle 1 Design: After discovering more issues within part 1 of the design, Team Cornell implemented the following changes to design part 1:
- Created separate chloramphenicol solution with 100% dimethyl sulfoxide (dmso) and 95% ethanol in a 25mg/mL ratio, both to be tested to see which antibiotic solution works best.
- 100% ethanol was sterile filtered to ensure zero contamination
- DMSO was not sterile filtered as it was an unopened container
- New DH5 alpha E. Coli cells were ordered to perform the transformation due to viability concerns
- New chromoproteins were selected: Parts 7N (BBa_K1033925, spisPink) and 7P (BBa_K1033927, asPink)
- Control plates were added with no antibiotic or plasmid within the E. Coli, it will now be a standard addition
- Sterility considerations:
- The glass beads were placed in ethanol for 30 minutes and autoclaved immediately after.
- Petri dish packages are now opened inside a biosafety cabinet (BSC) to ensure no contaminants come into contact with plates.
- Any process done outside of a BSC will be performed near an alcohol lamp
The New Build: The new experiment would be to identify which solution of chloramphenicol, dmso or ethanol, would lead to the best transformation results. Five 100 mm plates were made: two containing chloramphenicol dissolved in sterile-filtered 100% ethanol, two containing chloramphenicol dissolved in 95% dmso, and one with no antibiotic. Standard transformation protocol was followed.
Testing the new build: Parts 7N (BBa_K1033925, spisPink) and 7P (BBa_K1033927, asPink) from plate 6 of the iGEM 2021 DNA distribution kit were used in this round of chemical transformation experiments. The plates were prepared as described in the previous section. After 18 hours, team members observed that one plate with chloramphenicol dissolved in dmso seemed to show better growth, while the other dmso plate had relatively little growth. A similar result was observed for the plates with ethanol, while the plate with no-antibiotic had many colonies growing.
Learn *continued*: After this round of transformation, conferring with subteam and team leads, and additional literature review, team members reviewed the results and came to these conclusions/changes about this experiment:
- Sterilizing the glass beads and petri dishes before use seemed to provide good results
- Chloramphenicol dissolved in both sterile-filtered 95% ethanol and 100% dmso provided better results than water-dissolved antibiotic. From this result, we will no longer use water-dissolved chloramphenicol.
- New information learned:
- The container of ethanol used was not actual ethanol, but instead a mislabeled waste beaker. We identified pure 95% ethanol that was for general lab use and used it to prepare a new stock chloramphenicol solution for the next cycle. A new 25 gram bottle of chloramphenicol was ordered to ensure sterility and viability would not be a concern.
Cycle 3:
Redesign, again: After research and discussion with team leads, we decided to only dissolve chloramphenicol in ethanol instead of DMSO. This development was informed by the realization that the ethanol we have been using for the past transformations was contaminated (explained in cycle 3). Any perceived E. Coli growth from the previous transformation was therefore not considered.
A concern was raised among team members that there may have been too much DNA in our resuspensions. We found the resuspended double stranded DNA concentrations via a nanodrop spectrophotometer and decided to perform this transformation with lower concentrations of resuspensions in the hope that this would result in a greater transformation efficiency [1].
| 2021 Plate 4 Well | DNA Concentration (ug/ML) |
| K9 | 113 |
| K11 | 94 |
| I15 | 32 |
| G17 | 37 |
| C21 | 34 |
| 2022 Plate 4 Well | DNA Concentration (ug/ML) |
| B2 | 5 |
| J6 | 11 |
| P8 | 12 |
| D11 | 11 |
| F12 | 13 |
Lastly, a new shipment of chloramphenicol and true 95% anhydrous ethanol allowed us to have a proper stock solution of antibiotic when making plates.
Build: Following these preparation steps, a transformation was performed to test the new antibiotic stock, as well as to see how plasmid concentrations impacted transformation ability. Both control plates were plated with untransformed cells (blanks). One control plate contained chloramphenicol antibiotic to test if the antibiotic is effective while the other contained no antibiotic to test if the cells were viable.
A proper test: Parts K9 (BBa_K1033919, gfasPurple), K11 (BBa_K1033932, spisPink) , and C21 (BBa_K1033933, asPink) from plate 4 of the 2021 DNA distribution kit were selected for this transformation due to these resuspensions having a high concentration; they were 113 μg/mL, 94 μg/mL, and 34 μg/mL, respectively.
Results:
Learn:
- Further research indicated that the optimal concentration of DNA for transformations is 10-20 μg/mL. A new standard step of our transformation protocol, now, is to check the concentration of our DNA via nano-drop to ensure we only use parts with this concentration.
- Since sterility issues were fully addressed, our current transformation protocol may require reconsideration.
Cycle 4:
Procedural Shift: Under the impression that our protocol was faulty, we decided to try a different procedure. Courtesy of DeLisa Lab at Cornell University, we tried their protocol which includes a second ice bath following the initial heat shock. Sterility is no longer a major concern as we have made every last effort to ensure all processes regarding transformations are done near an alcohol lamp or in a biosafety cabinet.
Another concern regarding the state of the plasmid samples from the 2021 iGem distribution kit was brought to light. Given their age and our lack of information concerning its previous handling, the DNA could be damaged. Therefore, we decided to try to take our samples from the 2022 distribution kits instead. Interestingly, when rehydrating these parts, there was noticeably less color in the solution. This is most likely because of the smaller concentration of DNA as tested with the nano-drop.
Comparing Methods: To ensure experimental control, we performed two transformations – one using the new, DeLisa Lab protocol (Experiment 1) and one using the old protocol (Experiment 2). Experiment 1 bacteria were transformed with DNA from well I15 (BBa_K592012, eforRed) in plate 4 of the 2021 DNA distribution kit. Experiment 2 bacteria were transformed with DNA from well F12 (BBF10K_003463, T4T5_AmilCP) in plate 2 of the 2022 DNA distribution kit. The resuspension of these parts had concentrations of 32 ug/mL and 13 ug/mL, respectively. We also included a chloramphenicol plate with untransformed cells as a control.
Testing: The original and new protocols can be found on our Experiments page. After 12 hours, the Experiment 2 plate showed no bacteria growth (bottom right):
Experiment 1 showed plenty of growth – which was clear after 12 hours – as well as clear red chromoprotein expression – clear after 24 hours.
To check if each experimental output really did express the plasmid, we created liquid cultures of both Experiment 1 and 2 bacteria to see if we could identify a color change in the LB. This protocol was recommended by iGEM Team Uppsala, the original creators of the chromoproteins we are using. After 36 hours, the result appears to be that the bacteria were not transformed as the LB did not change color:
Takeaways and Progress:
- Sterility is likely no longer an issue, since the cultures now clearly contain E. Coli colonies and not contamination.
- The transformation procedure that we are using required modification. The DeLisa lab protocol successfully did so.
- There could be an issue with the plasmid we are using.
Cycle 5:
Procedural Shift: Given that the DH5α EColi strain was the bacteria being transformed, we also decided to try a protocol specific to this strain. Our bacteria were sourced from New England Biolabs (NEB) and they provide a separate, albeit very familiar, transformation protocol for this strain. The NEB protocol included a longer ice bath after the original heat shock, as well as incubating the transformed bacteria at 37C with 250 rpm agitation. However, we edited the original protocol in order to streamline the process while also ensuring that the protocol would still work properly. Furthermore, instead of using LB after the second ice bath, the NEB protocol used SOC outgrowth. SOC has a higher concentration of nutrients compared to LB which makes it a better option for incubation.
Build: This experiment was done solely with the NEB protocol. We did not use our original protocol or the DeLisa Lab protocol. Parts from plate 3 of the 2021 DNA distribution kit were used for this experiment. Wells 2J (BBa_K592011, cjBlue) and 2L (BBa_K592011, cjBlue) were rehydrated with concentrations of 11 ug/mL and 84 ug/mL, respectively. The concentration of well 2J was within the optimal concentration range of 10-20 ug/mL, so only well 2J was used for this experiment.
Test Complications: During the transformation, the bacteria were left in the incubator for about an extra 10 minutes. Moreover, during the plating process, we realized that each eppendorf tube required two plates:1 with 20 uL of bacteria and 1 with 200 uL of bacteria. Due to complications with the autoclave, we only had two plates available. Therefore, we decided to plate 20 uL first. After new LB agar was autoclaved, we plated 200 uL of bacteria onto new plates.
The plates containing 20 uL of bacteria had little to no growth. Considering the low volume used, this is not an unexpected result.
Takeaways:
- Sterile technique was not properly followed during this transformation
- Troubleshooting: Petri dish package may have been left open, LB agar was not fully autoclaved
- The NEB protocol seems to provide good bacterial growth. A repeat of this experiment but with a control (transformation performed with the DeLisa Lab protocol in comparison with the NEB protocol) could provide more insight
Cycle 6: The Color at the End of the Tunnel
Procedural Shift: After the multiple cycles of transformations, we decided to make more procedural changes through the next transformation. Instead of using a well with previous resuspended DNA, we opened a new well, 9F, and started from scratch. The well 9F contained the pink chromoprotein BBa_K1033926 . Furthermore, instead of leaving the NEB 5-alpha Competent E. coli cells on ice to thaw until we were ready to transfer, we decided to only thaw the cells for exactly 10 mins before starting the NEB transformation protocol.
Build: After changing the thaw time in our protocol, we performed the NEB protocol and plated 2 plates with 20uL and 200 uL of the cells with the chromoprotein inside of it. After approximately 20 hours, E. Coli growth with pink color began to show.
- Thawing for too long may have caused the optical density of competent cells to go above the necessary 0.5-0.7 absorbance (at 600 nm). By shortening the thawing time to 10 mins, it is more likely the optical density stayed within that 0.5-0.7 range.
- With color successfully seen, the next step was to perform a lysis of the cells to extract color.
Part 2: Extracting Color
Cycle 1:
Design and Build: To start the lysis of the pink chromoprotein colonies, we used the NEB lysis protocol with an arbitrary amount of 200 uL of lysis buffer to break open 1 mL of liquid cultures that were made after color was expressed. This ended up being more buffer than the protocol recommended in order to provide more supernatant for chromoproteins to be inside in case the cells absorbed all the buffer and created too many bubbles.
Test: By doing the lysis protocol, this was to test the pink expression seen in the liquid cell cultures were chromoproteins and not pink contamination. If the supernatant after the protocol was pink, it would be known that the pink seen was chromoprotein expression. After the protocol was completed, the supernatant was successfully the same pink shade as the original cell colonies meaning the lysed proteins were the desired chromoproteins.
Learn: Through the lysis protocol, we were able to confirm that the chromoprotein BBa_K1033926 was successfully transformed into DH5-alpha cells and their color could be successfully extracted from those cells. Now that we succeeded with color, the next step was to perform a successful Gibson Assembly.
Part 3:
Cycle 1: Starting out Gibson Assembly
Design and Build: For our Gibson Assembly, we planned on creating a PCR colony with our forward and reverse primers and gene fragments of csgA alpha, gamma, csgC, E, and G. CsgA alpha were also linked with the three chromoproteins of focus (asPink, aeBlue, and amajLime). These biobricks were then used within the NEB Gibson Assembly protocol to combine the backbone pSB1C3 and the gene fragments to create a full plasmid. These plasmids were later transformed into E. Coli cells and placed within liquid cultures with IPTG solution in order to induce color.
Test: At a molar concentration of 0.4 pmol of DNA per part, we ran 4 consecutive Gibson assembly reactions. They were csgA-alpha + asPink, csgA-alpha + amajLime, csgA-alpha + aeBlue, and csgA-gamma. Transformation occurred 24 hours after Gibson Assembly and DNA purification. 24 hours after transformation, the following images were taken:
Learn: After the experiment, we discovered possible errors in our protocol such as too high concentration of the plasmid being transformed and not purifying the DNA before beginning the transformation.
Cycle 2:
Procedural Shift: After the first Gibson Assembly attempt where no color was seen in the liquid cultures, we decided to edit the procedure by adding a DNA purification step before transforming the plasmid and lowering the concentration of the plasmid in order to attempt to get better results. Furthermore, unpurified PCR products have many leftover enzymes, nucleotides, etc, so DNA purification would ensure that only the DNA would be present during Gibson assembly.
Design & Build: We used 0.4 pmol of each DNA part, using NEBio’s DNA calculator to calculate the proper volume. Overall, 14 PCR reactions were performed to ensure the 4 necessary Gibson reactions could be performed. Once the Gibson reactions were performed, colony PCRs were performed to ensure the success of the Gibson Assembly.
Test: An example of a PCR reaction used in preparation for Gibson assembly is shown below:
| Tube | Primers | DNA | Length of DNA part |
| 1 | CP_Green_csgC-F (35) + csgE-csgC_R(28) | rbs_csgC | 421 |
| 2 | CP_Pink-CsgC_F(33) + CsgE-CsgC_R(28) | rbs_csgC | 421 |
| 3 | CP_Blue-CsgC_F(31) + CsgE-CsgC_R(28) | rbs_csgC | 419 |
| 4 | BCKBN-PRMTR_F(25) + CsgC-CsgA_gamma_R(38) | promoter_rbs_csgA_gamma (39) | 1162 |
| 5 | CsgC-CsgE_F(27) + CsgG-CsgE_R(30) | rbs_csgE (20) | 465 |
| 6 | BCKBN-TRMNTR_R(24) + CsgE-CsgG_F(29) | rbs_csgG_term (22) | 1098 |
| 7 | CsgC-CP_Green_R(36) + Linker-amajLimeF(75) | amajLime(43) | 749 |
| 8 | CsgC-CP_Blue_R(32) + Linker-ae_blueF(73) | ae_blue(42) | 753 |
| 9 | CsgC-CP_Pink_R(34) + Linker-as_pinkF(71) | as_pink(41) | 756 |
| 10 | BCKBN-PRMTR_F(25) + amajLime-LinkerR(76) | prmtr_rbs_csgA_alpha_linker (40) | 845 |
| 11 | BCKBN-PRMTR_F(25) + ae_blue-LinkerR(74) | prmtr_rbs_csgA_alpha_linker (40) | 843 |
| 12 | BCKBN-PRMTR_F(25) + as_pink-LinkerR(72) | prmtr_rbs_csgA_alpha_linker (40) | 843 |
| 13 | TRMNTR-BCKBN_F(23) + PRMTR-BCKBN_R(26) | pSB1C3 | 2125 |
| 14 | CsgA_gamma-CsgC_F(37) + CsgE-CsgC_R(28) | rbs_csgC(18) | 415 |
Concentrations for the PCR reaction were in the range of 150 - 300 ng/uL. 4 Gibson reactions were run afterwards: csgA-gamma, csgA-alpha-linked-pink, csgA-alpha-linked-lime, and csgA-alpha-linked-blue. 0.2 pmol concentrations of each relevant DNA part were used. An example of a Gibson Assembly reaction can be found below, at 0.4 pmol.
| csgAα + aeBlue | csgAα + asPink | csgAα + amajLime | ||||
| Gibsone Assembly Component | Tube # | Volume (uL) | Tube # | Volume (uL) | Tube # | Volume (uL) |
| csgC | 3 | 0.57 | 2 | 0.50 | 1 | 0.70 |
| csgE | 5 | 0.56 | 5 | 0.56 | 5 | 0.56 |
| csgG | 6 | 0.97 | 6 | 0.97 | 6 | 0.97 |
| csgAα + Linker | 11 | 0.92 | 12 | 0.96 | 10 | 1.13 |
| pSB1C3 | 13 | 3.48 | 13 | 3.48 | 13 | 3.48 |
| Chromoprotein | 8 | 0.93 | 9 | 1.01 | 7 | 0.90 |
| dH2O | 2.57 | 2.52 | 2.26 | |||
| G.A. Master Mix | 10.0 | 10.0 | 10.0 | |||
Learn: The concentrations of these Gibson reactions were better after using the PCR-cleaned DNA parts. This was incorporated into our future protocol designs for our Gibson Assembly procedure. The colony PCR was performed, but results showed nothing on the electrophoresis gels. Amplifying the colony PCR using chromoprotein primers or csgA-gamma primers was a good decision as they would help troubleshoot issues of color production or lack thereof However, a better decision would be to use primers that amplify the csgA and chromoprotein, or a combination of more fragments. This way, we would have a more confident confirmation that the Gibson Assembly worked. On a gel, the band would be at around 750 bp (for chromoprotein only) but we were unable to see any band. Even better, we could miniprep liquid cultures of our colonies and send the resulting plasmids for sequencing to have further confirmation. This will hopefully be incorporated if time and funds allow.
Further, in our troubleshooting process, we learned from the NEB website that the strain we were using, DH5ɑ E. coli, was not as compatible with protein expression as other strains.
Cycle 3: Procedural Shift
Procedural Shift: After literature review of why our colony PCR might have failed, we ended up learning that BL21(DE3) E. coli would be a better candidate for protein expression rather than DH5ɑ E. coli. This is due to proteases produced by the latter strain that may interfere with proper protein expression. DH5ɑ E. coli is typically utilized for plasmid preparation, while BL21(DE3) is better for protein expression [2].
Design & Build: We utilized the workflow from the 2019 Manchester iGEM team [3] in which they used DH5ɑ to prepare their plasmids, mini prepped liquid cultures of these cultures, then transformed BL21(DE3) with the miniprep product to result in successful protein production.
Test: A miniprep was performed on 4 mL of liquid cultures made with pink CP expressing colonies from Figure 12. The NEB Monarch Plasmid Miniprep Kit was used. Though the kit suggested using cultures that had been incubating for 12-16 hours, we used a culture that was incubated for ~24 hours as it takes our transformed colonies with the pink CP longer to grow and express the pink color. After miniprep was completed, a transformation of BL21(DE3) cells was performed using the NEB protocol. 200 uL was plated onto a LB+Chlor plate and incubated for ~18 hours. Figure 16 shows a dense lawn of pink colonies, indicating successful plasmid transfer!
Learn: From this cycle we learned that plasmid transfer between DH5ɑ and BL21(DE3) is the proper method to use for protein expression. We also learned that a lower volume of the transformation recovery culture should be plated, since we obtained such a dense lawn during this trial. The success of this experiment suggests that we should be using this process for the expression of our fusion proteins. This method will be implemented next in order to fix the issue of non-expression in our transformed cells that should be expressing our final constructs.
Product Development
Bioreactor
Bioreactor Manual.Overview: As our primary product is a bacterially produced bioink with expressed chromoproteins, it is essential that we can provide a safe and efficient place for our bacteria to grow. This bacterial home must maintain a precise balance of several variables and react to the changing environment of the bacteria in real-time. In order to produce a large amount of bacteria without needing a giant growth chamber, we have chosen to create a continuous flow bioreactor. In essence, we supply nutrients and dilute basic solutions to meet the needs of the growing E. coli, which produce the csgA subunits necessary for the assembly of the bioink. When existing E. coli have expressed the desired csgA protein, they are pumped out of the growth tank to be replaced by new generations of bacteria to continue the process. This whole process is monitored by a fleet of sensors, which connect to multiple support systems in order to optimize our protein production.
Cycle 1
Design Process: The nature of our project requires a functioning bioreactor, which we were able to implement by building upon the bioreactor implemented last year in Cornell’s 2021 iGEM project, Collatrix [1]. Despite appearing functional, this setup had setbacks that prevented it from performing at its desired capabilities. Thus, the goal of our engineering cycle was to improve and optimize this former setup into a system that suits our new requirements.
Our primary concern was the scale of our project compared to that of last year. With the need for a larger volume of effluent in order to supply the 3D printer with a proper amount of printing material, we considered increasing from 2 jars to 3 jars. This would also provide 3 compartments for us to be able to grow 3 different types of modified E. coli that expressed unique chromoproteins for a larger range of color when 3D printing. Before finalizing this larger system, we decided to first optimize the existing smaller bioreactor.
As we brainstormed further ways to optimize, we made various sketches of what we wanted the updated version to look like.
Build Process: Once we began building the updated version of the bioreactor, we realized that a nozzle shown in Figure 2, while in theory is great for sampling, would cause issues. These issues being that it could lead to potential contamination of the cells within the bioreactor, and how to ensure sterility on the outside nozzle. Additionally, once we put the sensors onto the sides of the bioreactor, we noted that three motor fans would be too many and they would bump into the sensors. Thus, we opted for one motor fan for our next iteration.
Cycle 2
Our primary area of optimization was the mixer blades. From last year, the mixer blade was harnessed to a threaded rod and connected to the motor via a shaft coupler. Initial considerations wondered if the use of 2-3 total mixer blades per threaded rod would be effective in providing uniform mixing of bacteria. This was considered as we progressed through our CAD design process. Ultimately, this multi-stack mixer blade was not used because of the presence of the sensors in the motor as well as the uncertainty of actual improvement. Due to the sensors, there was a physical limit to how far up the motor blade could be positioned. Additionally, it was skeptical if having more motor blades would actually improve efficiency when the additional blades would not be positioned in the center of the solution, but instead near the top or bottom. Thus, we opted to remain with only one mixer blade per bioreactor.
The mixer blades themselves were redesigned to have a total of 6 wings as well as having a larger radius. This was our solution to having more uniform mixing by increasing the surface area of the blades, there would be more volume that was constantly in motion to create even mixing. We maximized the length of the mixer blades to provide clearance for the sensors necessary for the maintenance of the bacteria by gorilla taping the sensors to the walls of the container.
In addition to the mixer blades, we made improvements to the electrical circuit of the bioreactor. In Cornell Collatrix 2021, the final system was not tested due to the fact that one of the motors was reversed and inadvertently overflowed the base additive over the electrical wiring and boards. Our solution to this problem was in two steps: firstly, building a modular electrical box to house all our motor driver boards, arduinos, wires, and breadboard, and secondly, improving the standardization of wiring and code such that a miscommunication or misalignment would not result in harming our project.
The electrical box was made with recycled plexiglass from the construction of the bioreactor housing. The corners were joined and sealed with waterproof caulk and a thin foam sheet was fitted to the bottom of the casing to allow the parts to stay more static. A physical improvement to the circuit was the addition of a working LCD display that will be used to periodically confirm that the system is functioning properly. Furthermore, we were able to consolidate the circuit down from four motor drivers to three.
The standardization of wiring and code was done in part with the help of multi-colored wires. All sets of wires from motor drivers were consistent colors as seen in figure 8 and when applicable, wires connecting ground and power were black and red respectively. Additionally, all wires running from motor drivers to motors were color coded in pairs to make pin input into the code more systematic along with the addition of marking tape on the wires. The marking tape allowed us to identify which wire of the colored pair would map to the positive or negative terminal of the motor (we selected a marked wire as the positive end), such that if a wire were to come loose or needed adjustment it would be simple to reattach and maintain. These new implementations to our electrical system were in attempts to optimize and improve on what was learned from Cornell Collatrix. A comprehensive guide to our electrical system assembly can be found in Contribution bioreactor section.
Future plans for the electrical system involve the transfer of the breadboard wiring onto a perforated board. This soldered connection is more secure and reliable as compared to our existing breadboard and would ensure that our system is solid and could be transportable. Ideally, this would evolve into a custom PCB system that would truly solidify our wiring.
Overall, the structure of the bioreactor is similar to that of Cornell Collatrix. Notable changes include the addition of an output in the lid of the bioreactor that allows easy access for pipette to harvest grown cells. When not in use, this output is closed with a plug as seen on the right green lid in figure 9. Additionally, taking note of our previous bioreactor, the base housing (pink lid) can be seen positioned farthest away from the electrical box. An improvement to this base system would be the dual motor system installed on the pink lid. By consolidating two base containers to just one eliminates added risk as well as optimizes the number of motor drivers necessary for the electrical system.
Software: Similarly to the hardware, we inherited the software for the bioreactor from Cornell’s 2021 iGEM project, Collatrix. The code controls the major components for bacterial growth, such as pH, O2 concentration, temperature, and feed. With the aid of an Arduino, the native packages from the sensors that we utilized were installed and used to monitor these levels. La Chatelier’s principle will be applied in the Arduino such that when either pH, O2 concentration, or temperature are not at the optimal level, this would trigger a motor to input base, O2 bubbles, or adjust the sous vide machine to counteract the change to reestablish equilibrium.
| Conditions | pH | O2 Concentration | Temperature | |||
| Change | drop | rise | drop | rise | drop | rise |
| Effect | Input base | N/A | O2 bubbles | N/A | Sous vide machine | Sous vide machine |
The feed of the system is monitored differently and has to be modeled as the feed has to be supplied accordingly to the bacteria growth rate. It is necessary that the feed for the bacteria also increases exponentially to maintain optimum equilibrium between the feed and the bacteria. With a stepwise feed cycle, we are able to mimic an exponential growth curve by inputting increased amounts of feed at a set time interval, always maintaining more feed than the bacteria need. This implementation allows us to always have excess feed, minimizing the possibility of feed being a limiting factor, therefore resulting in better growth. An excess supply of feed also optimizes process time by minimizing the amount of calculation necessary for the Arduino to perform in the loop.
Testing
To test the bioreactor, it was set up in our lab space and tested in comparison to a shaker flask culture. All parts were sterilized according to our bioreactor manual. This test was meant to compare growth in both environments. Since our reactor was optimized for pH, oxygen, and feed to provide a better growth environment, we expected to see greater growth in the reactor versus the shaker flask.
During set up, one of the mason jars for the reactor cracked. Thus, we had to continue the testing with only one jar.
The reactor was set up with 1000 mL of LB broth, 1000 uL chloramphenicol solution, and inoculated to a starting OD of 0.04 with E. coli DH5ɑ that was transformed with the asPink chromoprotein. Though this was not our fusion protein, we wanted to observe growth that best reflected the transformed cells we would be using for the final product, so we decided the transformed pink colonies would be a better model to test with than un-transformed cells. Sodium hydroxide at pH 11 was used as a base to control for pH. The reactor was temperature controlled in a water bath at 37°C.
The shaker flask was set up with 60 mL of LB broth, 60 uL chloramphenicol solution, and inoculated to a starting OD of 0.04 using the same cells as the reactor. The flask was temperature controlled in a shaking incubator, shaking at 150 rpm. Due to the size of our incubator, we could not go higher than 150 rpm or the flask would tip and spill.
Both the reactor and flask were left to run for 40 hours. Samples were taken at 8 hour intervals for both. At first we began with single aliquots, but starting from hour 24 we took three aliquots so we could obtain average OD values in case of data outliers. Between hours 8 and 16 of the reactor, the tape we used to fasten some tubing had fallen off and we did not have extra to use to refasten them. This was interfering with the impeller as the tubing and wires were becoming tangled without being fastened by tape. Thus, the reactor was left for another 8 hours without stirring or oxygen control to avoid overflowing, but was maintained at 37°C. To maintain consistency, the shaker flask was also removed from shaking and kept at 37°C.
Figure 14 below shows the average OD600 readings for the 40 hour period, with error bars depicting the standard deviation. Both setups showed relative stagnation until between 32 to 40 hours, upon which cell density increased greatly. Based on this data, the flask exhibited a greater rate of growth at later times compared to the bioreactor, which was not what we expected.
Our team came up with several reasons for this observed pattern:
- Growth rate lowered between 8 and 16 hours due to lack of rotation/spinning
- Our transformed cells have typically taken longer than usual (i.e. compared to wild type E. coli DH5ɑ) to grow and exhibit any color, which suggests why the cell density did not increase until later times
- Feed was only periodically pumped into the reactor, and might not have been done so often enough to refresh the cultures
Another run of this experiment will be conducted to gain more data, especially since the standard deviations at later times are so large. Some fixes will also need to be made to the reactor, such as finding a more secure way to fasten tubing, editing the time between pumping feed, and ensuring pH adjustment is operating as intended. With these adjustments, we still hypothesize that the bioreactor will exhibit better growth than the flask.
3D Printer
Overview: This year our project is focused on the development of environmentally friendly bioart using engineered bacteria encoded to create a variety of colorful hydrogels that we can arrange in a design, and additionally supplementing this art by modifying the material to uptake carbon dioxide and (volatile organic compounds) VOCs to help the environment, essentially creating a living material that can perform all these functions. The primary method of delivering this hydrogel into our final art product is through a modified 3D bioprinter.
Our project this year will be focused largely on developing 3D bioprinting capabilities through the modification and upgrades to a traditional 3D printer. We will be using the base of a Makerbot Replicator 2 and adapting the printhead and print code to remove the plastic heating elements and become compatible with hydrogel extrusion. Inspired by the 3D printing systems of Dr. Joshi at Northeastern University and with feedback from Dr. Meyer at the University of Rochester, and previous work done by the TU Delft 2015 iGEM project, our team will be developing a new method of bioprinting using shear-thinning hydrogels printed without the use of specialized substrates, meaning printing can be accomplished on a wide range of surfaces to enable the creation of unique art [2, 4].
Hardware:
Cycle 1
The major hardware component of this project is the development of a biocompatible hydrogel 3D bioprinter. We will be accomplishing this through the modification of a 3D bioprinter to become compatible with hydrogel extrusion. Our base printer is a Makerbot Replicator 2, which will be altered in 2 primary ways: the adjustment of the print head, and the creation of a hydrogel reservoir to replace the traditional plastic extrusion roll.
Most household 3D printers print plastic through the use of a heating element designed to melt the extruded material, and quickly cool and resolidify in layers. Heating elements can heat the plastic to over 200 degrees Celsius. Temperatures this extreme would cause damage to our hydrogel and kill live bacteria in the hydrogel (thus eliminating any CO2 uptake capabilities of our art) [1, 3]. In order to avoid this, we will be removing the heating elements from the printing head of the Replicator. The removal of the heating element, which also serves as the extrusion nozzle, will then be replaced with a custom-designed hydrogel extrusion head. The extrusion head will feature a motor connected to a syringe needle through a series of gears, which will serve as the new extrusion head for our 3D printer. The exact diameter and extrusion rate will be determined through testing of model hydrogels [see extrusion testing for more information]. In order to house the needle and syringe, we will design a new 3D printed part to replace the carriage of the Replicator. Since the specialized part will be 3D printed, it is designed based on the existing carriage assembly, and will thus be easily interchangeable with the existing carriage, requiring fewer hardware and software changes to make the printer compatible.
An initial 3D printed design was created, however it was found to not fit using the originally determined CAD dimensions (specifically the base dimensions were off by 1 mm, and the syringe holder slot was not properly fitted to allow the syringe to slide in), and as such a second version was printed using the modified dimensions. The modified extrusion head was mated with the syringe, and then installed onto the print head (in place of the heated element) to be used for testing and further printing.
In order to feed the hydrogel to our extrusion head, we will be creating a new reservoir for our hydrogels, which will help serve as the “ink cartridge” for our printer. Initially, there are 2 proposed designs for the reservoir. The first would be to use a syringe preloaded with hydrogels, which is attached directly to a linear stepper motor that drives the extrusion of the hydrogels [1]. This design would utilize a similar hydrogel extrusion model to the Collatrix hydrogel extrusion form last year, with adjustments made to accommodate for the extrusion through tubing into the printer extrusion head. This design is limited by the size of the syringe, where each ink cartridge can only hold as much as the syringe volume, and would then need to be replaced. The second design would use a small tank, containing a larger volume of hydrogel, with tubing fed through a peristaltic pump into the printer tubing. In this design, the extrusion rate is directly proportional to the pump rate, and the use of a larger reservoir means that fewer ink refills are necessary, however, the use of a peristaltic pump for hydrogel extrusion may lead to complications with 3D printing. After testing and feedback from interviews our team decided to move forward with the syringe model as 20mL syringes hold enough ink for several development scale prints, and can be quickly swapped out to test different inks without needing to clear transport tubes as required with a pump based design.
Cycle 2
In order to further incorporate the new printing head with the old printer, a geared syringe holding block was built that could integrate with the previous motor used for plastic extrusion. This allows the former wiring and code used to run the printer to be seamlessly transferred to the syringe based system, removing the need for a new software package to be designed and installed. The initial printed model for this head was off in several dimensions, as tolerances between the new part and the existing hardware had to be tested. The first model was also able to be used and reconfigured in order to determine what modifications would make the system run smoothly.
After installation the new design worked, and was able to extrude prepared hydrogels using existing printer gcode. However, the extrusion rate used for the plastic based system pushed a much smaller diameter feed using a gear with a smaller diameter, two factors that both contributed to an extrusion rate that was initially nearly two orders of magnitude too high. After reducing this speed, the mechanisms behind the printer head were safe from self destruction and extrusion testing could begin.
Modifications were made to the syringe diameter, nozzle, and racking system in order to improve the print quality. A video camera was also added to the printer in order to record video and allow for a livestream to be connected. Achieving a quality print required honing in on sufficient balance between print speed and volume, nozzle geometry, and ink composition. The bacterial hydrogel mimetic used was gelatin, which presented issues due to forming clumps and forming weak bonds after printing in its hydrated state. With the thixotropic bacterially produced hydrogels, they will have the ability to reconnect and form strong bridges and 3D structures.
Software: There are two major software components relating to the 3D printer. The first part is the conversion of the submitted images into a 3D printer compatible matrix, and the second portion is the adjustment to the 3D printing code itself.
Our project aims to make bioart accessible to large audiences, and part of that goal is allowing publicly submitted art to be created on our 3D bioprinter. In order to make these submitted art pieces compatible with our 3D printer, we need to convert the image format (either .png or .jpg) file into a 3D printer compatible matrix format. We will be accomplishing this through our app and translation into matrices using MATLAB. Our app will accept the image submission, and then feed the image data to a MATLAB script, which will use image analysis to convert the image into a 3D matrix of values, using 4 layers (1 layer for each color channel and the 4th layer corresponding to the 3D height of each pixel). This converted image will then be fed to our 3D printer where it will then be printed.
In addition to the conversion of the image type, we will be adjusting our 3D printing code to make our 3D printer biocompatible by editing gcode in the Cura slicing software. Traditional 3D printers which extrude plastic through the use of a heating element in the head must control the temperature of the heating element in the base code. Since our 3D printer is adjusted to remove the heating element, we must also remove the code controlling heating in order to enable the 3D printer to begin printing without issues, as most 3D printing codes will not run if there is no heating element [3]. In addition, we must also make changes to enable multiple color channels, since most 3D printers are only designed to print in a single color. We will also be adjusting the 3D printing extrusion code to be compatible with our hydrogel extrusion. Since we will be using a combination of stepper motors and pumps to extrude our hydrogels, we must adjust the extrusion code to activate our motor/pump proportional to the layer height of our 3D printed image. All changes will be made directly to the 3D printing gcode made after the processing of our image into an stl, then slicing into the final print code. The changes will be made using Cura slicing software, and gcode changes will be logged for future use.
Handheld Bioprinter
Overview: As an alternative to a large 3D printing machine, we decided to create a 3D handheld bioprinter to provide a mobile and user-friendly device for artists. Previous IGEM teams such as Warwick and LMU-Munich have developed 3D bioprinters to aid in tissue and/or bone repair, yet none of them have created a handheld device. This handheld bioprinter of 145 mm height x 32.5 mm width x mm length will contain two 3 mm syringes with extrusion nozzles to allow printing of two colors. This device will allow for easy access to bioprinting in the art industry and will act as a substitute for paint brushes and/or colored pencils.
Hardware: The hardware is similar to the gradient machine implemented in Cornell Collatrix and based off other sources [1,3]. It operates using two stepper motors powered by motor drivers connected to an Arduino Uno. The rotation of the stepper motor will push a threaded rod through a syringe pusher into a syringe to initiate the extrusion process. However, we plan on extruding through both syringes as opposed to Collatrix’s gradient mixer, which uses two syringes to extrude from one nozzle. Therefore, a push was integrated into the breadboard to control stepper motors and drivers to alternate between syringes and colors.
Bioprinter Casing: When designing the bioprinter casing, we prioritized minimizing space while still being able to hold two syringes. The casing includes a base that protrudes from the back to maintain syringe pusher position and prevent rotation along with the threaded rod. The syringe pusher will move along the base to initiate extrusion.
Reloading: When considering reloading, our initial idea was to use the counterclockwise rotation of the stepper motors and threaded rods to move the syringe pusher upstream of the base. The syringe pusher was designed to reload the syringe as it was moved up the base. However, this mechanism failed since the stepper motor’s maximum speed of 15 rpm was excruciatingly slow and ineffective for reloading. Thus, we came up with an alternative solution by redesigning the casing to include a cover attached to a friction freeze hinge and a sliding rod to fixate the cover when closed. This would allow users to open the casing and remove syringes when printing finished. Second iteration of the casing can be seen here:
Testing: We designed the handheld bioprinter based on the comfortability and ease of use. This was tested based on how comfortable it was to hold the bioprinter and by printing an entire syringe of hydrogel. Based on our extrusion tests, the bioprinter could be held comfortably and printed successfully.
Testing was conducted using gelatin hydrogel. We tested extrusion by determining if it could extrude the hydrogel smoothly and at a consistent rate throughout the nozzle. Speed could be adjusted through the code by changing the rate of revolution of the stepper motor. Tested at the fastest setting of 15 rpm, extrusion rate was 0.882 mm/s. However, this also revealed inconsistencies in printing. To fix this, we added a 1 mml syringe tip and tested. Testing failed due to pressure buildup and we concluded it was more effective to maintain the 3 mm syringe tips. Extrusion can be seen here:
Accuracy and precision of the bioprinter was tested by printing sketches of sample bioart and comparing expected results to printed products. From this, resolution, precision, and accuracy were determined through user testing seen in Figure 9.
In the above video, one can see that in the middle of the printing process, the hydrogel pauses for a second or two before continuing to print. This is due to the fact that when filling the syringe, there were air bubbles that did not allow for further movement of the gel. All in all, the smiley face drawing showed us that our resolution and accuracy can improve with further iterations of the handheld device, in which we can shrink the syringe size for precision and use a different type of hydrogel that does not contain gelatin.
Cyclodextrin (CD) Nanofiber Incorporation
After testing gelatin and glutaraldehyde hydrogel, we attempted to enhance the mechanical properties of the gel by combining CD nanofiber into it, which have also been used for VOC-uptake applications [**]. Professor Tamer Uyar from Cornell Human Ecology Department kindly offered us 0.6 g of electrospun CD nanofiber from his lab to experiment on.
Sources
Wetlab
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[2] T7 and DH5 alpha. Protocol Online. (n.d.). Retrieved from http://www.protocol-online.org/biology-forums-2/posts/12993.html.
[3] Act II: Experimental and Results. iGEM Manchester 2019 - Cutiful. (n.d.) Retrieved from https://2019.igem.org/Team:Manchester/Experiments.
Product Development
Bioreactor
[1] “Hardware.” 2021 iGEM Team:Cornell, 2021, https://2021.igem.org/Team:Cornell/Hardware
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3D Printer[1] Ioannidis, K., Danalatos, R. I., Champeris Tsaniras, S., Kaplani, K., Lokka, G., Kanellou, A., Papachristou, D. J., Bokias, G., Lygerou, Z., & Taraviras, S. (2019). A Custom Ultra-Low-Cost 3D Bioprinter Supports Cell Growth and Differentiation. Frontiers in Bioengineering and Biotechnology. https://doi.org/10.3389/fbioe.2020.580889
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Handheld Bioprinter[1] Ying, G., Manríquez, J., Wu, D., Zhang, J., Jiang, N., Maharjan, S., Medina, D. H. H., & Zhang, Y. S. (2020, August 21). An open-source handheld extruder loaded with pore-forming bioink for in situ wound dressing. Materials Today Bio. Retrieved September 30, 2022, from https://www.sciencedirect.com/science/article/pii/S259000642030034X
[2] Duraj-Thatte, A. M., Manjula-Basavanna, A., Rutledge, J., Xia, J., Hassan, S., Sourlis, A., Rubio, A. G., Lesha, A., Zenkl, M., Kan, A., Weitz, D. A., Zhang, Y. S., & Joshi, N. S. (2021, November 23). Programmable microbial ink for 3D printing of living materials produced from genetically engineered protein nanofibers. Nature News. Retrieved September 30, 2022, from https://www.nature.com/articles/s41467-021-26791-x
[3] Hakimi, N., Cheng, R., Leng, L., Sotoudehfar, M., Ba, P. Q., Bakhtyar, N., Amini-Nik, S., Jeschke, M. G., & Günther, A. (2018, April 11). Handheld skin printer: In situ formation of planar biomaterials and tissues. Lab on a Chip. Retrieved October 6, 2022, from https://pubs.rsc.org/en/content/articlelanding/2018/lc/c7lc01236e
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