Materials:
Equipment:
Protocol:
Colony PCR played a big role in troubleshooting and determining the efficacy of our golden gate assembly reactions following transformation. NEB Robust Colony PCR Protocol (accessed 7/13/23) to fit our project needs, namely to accommodate liquid culture samples. Culture plate:
1. Transform ligation mix or other plasmid-con- taining reaction mixture into the desired bacterial strain, and incubate agar plates overnight at the appropriate temperature.
2. Set up 50 µl NEB OneTaq reactions consisting of 25μl 2x OneTaq master mix, 200nM primers, and add diH2O to top off to 50μl.
3. Use a sterile pipette tip to poke or pick up individual colonies and dip into each reaction tube. Generally speaking, the more the better (up to a colony or pellet with 2mm diameter), however merely poking colonies works when you do not wish to disrupt the colony, have good primers, and use touch-down PCR.
4. As soon as the solution looks cloudy, remove the pipette tip. To create a stock of each individual colony either: a.) Dip the pipette tip into 3 ml growth media with appropriate antibiotics and culture overnight. Or b.) Streak the pipette tip onto another agar plate containing the appropriate antibiotics and grow overnight.
5. Transfer reactions to a PCR cycler, and perform touch-down PCR within the NEB OneTaq guidelines.
6. Load 4–6 µl of each PCR reaction directly onto an agarose gel, alongside an appropriate DNA ladder.
We made frequent use of the Touch-Down PCR methodology in nearly every PCR experiment, and achieved significantly better results than with the traditional PCR methodology. Touch-Down PCR can be applied to many PCR kits, making sure to stay within the prescribed thermocycle parameters.
Touch-Down is a method for increasing specificity of PCR reactions and can be applied to many different PCR kits and is particularly useful when dealing with complex DNA samples or when you want to minimize nonspecific amplification. Touchdown PCR uses a cycling program where the annealing temperature is gradually reduced (e.g. 0.5-2°C /every second cycle). The initial annealing temperature should be several degrees above the estimated Tm of the primers. The annealing temperature is then gradually decreased until it reaches the calculated annealing temperature of the primers or some degrees below. Amplification is then continued using this annealing temperature.
Initial Denaturation: This step is the same as in standard PCR, where the DNA is denatured at a high temperature.
Touchdown Annealing: In the initial 10 cycles, the annealing temperature is set 3°C higher than the actual Tm (melting temperature) of the primers, and stepped down 0.5°C each cycle reaching a final temp of 2 °C below the calculated annealing temp. This ensures that only highly specific binding occurs, reducing nonspecific binding or primer-dimer formation. In each subsequent cycle, the annealing temperature is lowered by a few degrees.
Extension and Amplification: The PCR reaction continues with primer extension and DNA amplification, as in standard PCR.
Annealing: In the remaining ~20 cycles, annealing is continued at the lowest temp reached during touchdown (eg. 2°C below predicted annealing temperature)
Extension and Amplification: The PCR reaction continues with primer extension and DNA amplification, as in standard PCR.
Final Annealing and Extension: In the later cycles, the annealing temperature stabilizes at a temperature closer to the actual Tm of the primers, allowing for more efficient amplification of the target DNA.
The idea behind touchdown PCR is to start with stringent annealing conditions to enhance specificity and gradually transition to more permissive conditions to promote amplification of the target DNA. This helps reduce background noise and enhance the yield of the desired PCR product. Touchdown PCR can be particularly useful when dealing with templates that have a high degree of sequence similarity with non-target sequences or when you want to maximize the formation of difficult to amplify amplicons.
Protocol adapted from NEB Touch Down PCR (accessed 7/20/23)
This protocol has been adapted from iGEM team HK SSC (2019) and Choi’s 2021 transformation protocols for cyanobacteria. While the protocol could benefit from larger culture volumes, we made use of smaller volumes to maximize variations of the protocol that we could attempt. Our target organism, Microcystis aeruginosa, is known to be both naturally and electro-competent, hence the pre-pulse incubation period with DNA.
Materials:
Equipment:
1. Culturing Conditions: Grow Microcystis aeruginosa cells at 25C under a 12/12 light cycle until they reach early log growth phase.
2. Cell Collection: Aliquot 10 mL of Microcystis aeruginosa culture into a centrifuge tube.
3. Centrifugation: Using a 4°C centrifuge, spin at 1000g for 12 minutes or until the supernatant becomes clear.
4. Wash Steps: Gently resuspend the cell pellets in 2.5 mL of 0.1 mM HEPES. Centrifuge at 1000g for 12 minutes or until the supernatant becomes clear. Repeat steps 3-4 for a total of three washes to reduce the concentration of extracellular salts and materials.
5. Viability Check: To assess cell viability, transfer 50uL of the cell suspension to a 0.2 mm electrotransformation cuvette. Perform electroporation with a pulse duration to achieve a time constant of ~5 msec. Follow the manufacturer's instructions for the specific electroporation system (e.g., Bio-Rad E. Coli Pulser). Repeat wash steps (3-4) if arcing occurs during electroporation.
6. DNA Gradient: Aliquot 50 uL of the cell suspension into micro-centrifuge tubes. Add 1 ng to 5 pg of the genetic construct in a gradient. This gradient approach allows for optimized DNA concentration to be utilized, maximizing transformation efficiency due to challenges in precise cyanobacteria cell count quantification.
7. Pre-pulse Contact: Transfer the tubes containing the cell suspension and DNA to ice and allow them to incubate for 1-5 hours.
8. Electroporation: Transfer 50 uL of each cell preparation to a sterile, chilled 0.2 mm electrotransformation cuvette. Deliver a 2.5 kV pulse according to the manufacturer's instructions (e.g., Bio-Rad E. Coli Pulser). Immediately after pulsing, add 1 mL of BG11 + 5mM Glucose medium to the cuvette and transfer the mixture to a culture tube (https://doi.org/10.3390/biom11020214, accessed 10/1/23).
9. Incubation: Transfer the culture tubes with cell suspension back to the original growth conditions (25C, 12/12 light cycle). Incubate for 24 hours. Note* it is optimal for this step to be timed when growth lights are on.
10. Antibiotic Selection: Add 50 ug/mL chloramphenicol to the culture tubes. Incubate for approximately 2 days to allow antibiotic selection to take place.
11. Transformation Efficiency Assay:Assay transformation success and efficiencies as needed, considering the specific parameters and techniques relevant to your project.
Purpose: Test and analyze a plasmid's capability to transform our selected strain of M. aeruginosa through its capability of naturally uptaking foreign DNA.
Materials:
Equipment:
1. The natural transformation protocol was adapted from the 2019 HK_SSC iGEM team. [24].
2. Pipet a sample of 30 mL of M. aeruginosa culture and centrifuge for 10 minutes at 2500 x g
3. Decant the supernatant and wash cells using 10 mM NaCl
4. Centrifuge again and a tenth of the original volume of BG-11 to resuspend the pellet
5. Pipet 2 µg of pSHDY into the sample and mix by inverting
6. Incubate the sample in the dark overnight at 20°C
7. The next day, add the culture to 25 mm diameter, 0.45 µM pore size millipore filters placed on BG-11 plates
8. Incubate the plates for 20 hours at 20°C in a 12:12 light dark cycle with a light lux level < 3200 and consistent gas exchange
9. Transfer the filters to 2% agar BG-11 plates containing antibiotics to select for transformants. The antibiotics used were spectinomycin, kanamycin and chloramphenicol at a 15µg/mL concentration.
10. Incubate the plates for 7-10 days at 20°C
Purpose: BG11 media was made for the culturing and growth of M. aeruginosa. This media was selected due to its established efficacy in cultivating freshwater microalgae, marine microalgae, and cyanobacteria. The creation of this media as opposed to others is simpler and only requires materials more commonly found in the laboratory, relative to other options.
Note: *KOD vitamins can be substituted for other supplements such as a vitamin B12+ biotin solution.
Materials:
Equipment:
Protocol:
1. Fill the glass bottle or flask to 900 mL with milliQ water.
2. Add 10 mL of each reagent listed sequentially:
3. Add 1 mL of 1X Trace Metals solution to the flask.
4. Measure and adjust the pH of the solution to 7.5 using a pH meter and 5M HCl respectively.
5. To sterilize, place the bottle/flask in an autoclave and run a 30 minute liquid cycle.
6. After autoclaving, allow the solution to cool, add 10 µL 1000X KOD vitamins.
7. Store at 4°C.
Purpose: Golden Gate Assembly will be used to create stealthed and unstealthed versions of pSPDY with a promoter, fluorescence, and conjugation insert.
Materials:
Equipment:
The plasmid was assembled according to the PaqC1, T4 DNA ligase NEB protocol
1. Create a 20 µL reaction by combining the following in a PCR tube: 2 µL Hi T4 DNA Ligase 2 µL 10X T4 DNA Ligase Buffer 2 µL PaqC1 0.5 µL PaqC1 activator 75 ng of vector with 150 ng of each insert MilliQ water up to 20 µL
2. Run the reaction in the thermocycler under the following cycle parameters: 60 cycles at 37°C for 5 minutes, the 16°C for 5 minutes enzyme inactivation at 60°C for 5 minutes held at 4°C
3. The Golden Gate product was transformed into TOP10 E. Coli for replication, extracted with miniprep, and verified with PCR and gel electrophoresis to determine the successful plasmid assembly
Purpose: Gel electrophoresis is employed to separate and visualize DNA fragments, enabling the assessment of PCR reaction success and the precise determination of fragment sizes. We utilized 0.8% agarose gel.
Materials:
Equipment:
Protocol:
1. Measure 0.8 g of agarose and add it to 100mL of 1X TBE Buffer.
2. Microwave solution in increments of 30 seconds until agar is fully dissolved. Swirl in between rounds.
3. Let the solution cool down to about 5°C.
4. Add 5 µL of 20,000X Apex DNA Safe Stain to the agar and swirl to mix.
5. Pour agar into casting tray with well comb, let it solidify.6. Make the following sample mixes:
7. Load sample and DNA ladder into wells.
8. Run gel at a voltage of 120V.
9. Stop the gel run when the loading dye is around 80% down the gel.
10. Visualize with gel imaging system.
Note: Protocol adapted from Zymo Research's Quick DNA HMW MagBead Kit
Appendix A: Microbial Lysis
1. Pipette 800 µL of M.aeruginosa culture, and centrifuge at 7000 rcf for 6-8 minutes or until the pellet forms.
2. Remove supernatant and set aside on ice.
3. Add 100 µL 1X PBS buffer to the pellet and resuspend with a vortexer.
4. Centrifuge at 5000 rcf for 1 minute.
5. Remove supernatant and combine supernatant with leftover supernatant from step 2
6. Add 1 mL PBS to the pellet and resuspend.
7. Centrifuge at 5000 rcf for 1 minute, save the pellet and discard the supernatant. Add 100 µL of TE buffer to the pellet (centrifuge) and resuspend with TE buffer.
8. Add 20 µL of 10% SDS to the pellet.
9. Incubate between 60°C - 80°C in a heat block for 10 minutes.
10. Add saved supernatant from step 2 to the incubated sample.
11. Add 10 µL of Proteinase K (pipette mix 10X) to the sample.
12. Incubate at 55°C for 10 minutes on a heat block.
13. Centrifuge at 5000 rcf for 1 minute or until a pellet forms. Record the supernatant volume and discard the supernatant.
14. Add equivalent amounts of MilliQ water to replace the supernatant.
15. Add 800 µL of MagBinding Buffer and mix well.
Appendix B: DNA Binding and Extraction
1. Add 400 µL of Quick-DNA MagBinding Buffer to 400 µL of Microcystis aeruginosa sample. Mix 5 times by pipetting up and down.
2. Add 33 µL of MagBinding Beads to the solution. Mix 5 times by pipetting up and down.
3. Secure the sample on a vortexer and vortex for 10 minutes at a medium-intensity setting.
4. Transfer to a magstand, remove and discard the supernatant.
5. Add 500 µL MagBinding Buffer. Mix by pipetting up and down 5 times to resuspend.
6, Vortex at setting 2 for 5 minutes.
7. Transfer to magstand, remove and discard supernatant.
8. Add 500 µL of DNA pre-wash buffer. Pipette up and down 10 times to mix.
9. Transfer to the magstand, remove the supernatant and discard.
10. Add 900 µL of gDNA wash buffer, and mix by pipetting up and down 10 times.
11. Transfer all liquid and beats to a new microcentrifuge tube.
12. Transfer to a magstand, remove and discard supernatant.
13. Repeat steps 10-12.
14. Dry beads on a heat block set to 55°C for 10 minutes.
15. Add 58 µL DNA elution buffer. Place on the vortex for 5 minutes at room temperature.
16. Transfer for magstand, remove supernatant, and place into a new microcentrifuge tube.
17. Store at -20°C.
Purpose: This process was used to prepare the genomic DNA of M. aeruginosa UTEX 2385 for genomic sequencing with Oxford Nanopore MinION. The results were then utilized by the Stealth program to find underrepresented sequence motifs.
Materials:
Equipment:
DNA Library Preparation
1. Thaw the DNA Control Sample (DCS) at room temperature, spin it down, mix by pipetting, and place it on ice.
2. Prepare the NEBNext FFPE DNA Repair Mix and NEBNext Ultra II End Repair / dA-tailing Module reagents in accordance with the manufacturer’s instructions, and place them on ice.
3. Prepare the DNA in nuclease-free water:
4. In a 0.2 ml thin-walled PCR tube, mix the following:
Between each addition, pipette mic 10-20 times
5. Ensure the reaction is thoroughly mixed by gently pipetting and spin down briefly.
6. Using a thermal cycler, incubate at 20°C for 5 minutes and 65°C for 5 minutes.
7. Resuspend the AMPure XP Beads (AXP) by vortexing.
8. Transfer the DNA sample to clean 1.5ml Eppendorf DNA LoBind tube.
9. Add 60 µL of resuspended AMPure XP Beads (AXP) to the end-prep reaction and mix by flicking the tube.
10. Incubate on a Hula mixer (rotator mixer) for 5 minutes at room temperature.
11. Prepare 500 µL of fresh 70% ethanol in nuclease-free water.
12. Spin down the sample and pellet on a magnet until the supernatant is clear and colorless. Keep the tube on the magnet, and pipette off the supernatant.
13. Keep the tube on the magnet and wash the beads with 200 μl of freshly prepared 70% ethanol without disturbing the pellet. Remove the ethanol using a pipette and discard.
14. Repeat the previous step.
15. Spin down and place the tube back on the magnet. Pipette off any residual ethanol. Allow to dry for ~30 seconds, but do not dry the pellet to the point of cracking.
16. Remove the tube from the magnetic rack and resuspend the pellet in 61 μl nuclease-free water. Incubate for 2 minutes at room temperature.
17. Pellet the beads on a magnet until the eluate is clear and colorless, for at least 1 minute.
18. Remove and retain 61 μl of eluate into a clean 1.5 ml Eppendorf DNA LoBind tube.
Adpater Ligation and Clean-Up
1. Spin down the Adapter Mix H (AMX H) and Quick T4 Ligase, and place on ice.
2. Thaw Ligation Buffer (LNB) at room temperature, spin down and mix by pipetting. Due to viscosity, vortexing this buffer is ineffective. Place on ice immediately after thawing and mixing.
3. Thaw the Elution Buffer (EB) at room temperature, mix by vortexing, spin down, and place on ice.
4. To enrich DNA fragments of 3 kb or longer, thaw one tube of Long Fragment Buffer (LFB) at room temperature, mix by vortexing, spin down, and place on ice.
5. To retain DNA fragments of all sizes, thaw one tube of Short Fragment Buffer (SFB) at room temperature, mix by vortexing, spin down, and place on ice.
6. In a 1.5 ml Eppendorf DNA LoBind tube, mix in the following order:
7. Ensure the reaction is thoroughly mixed by gently pipetting and spin down briefly.
8. Incubate the reaction for 10 minutes at room temperature.
9. Resuspend the AMPure XP Beads (AXP) by vortexing.
10. Add 40 μl of resuspended AMPure XP Beads (AXP) to the reaction and mix by flicking the tube.
11. Incubate on a Hula mixer (rotator mixer) for 5 minutes at room temperature.
12. Spin down the sample and pellet on a magnet. Keep the tube on the magnet, and pipette off the supernatant.
13. Wash the beads by adding either 250 μl Long Fragment Buffer (LFB) or 250 μl Short Fragment Buffer (SFB). Flick the beads to resuspend, spin down, then return the tube to the magnetic rack and allow the beads to pellet. Remove the supernatant using a pipette and discard.
14. Repeat the previous step.
15. Spin down and place the tube back on the magnet. Pipette off any residual supernatant. Allow to dry for ~30 seconds, but do not dry the pellet to the point of cracking.
16. Remove the tube from the magnetic rack and resuspend the pellet in 15 μl Elution Buffer (EB). Spin down and incubate for 10 minutes at room temperature. For high molecular weight DNA, incubating at 37°C can improve the recovery of long fragments.
17. Pellet the beads on a magnet until the eluate is clear and colorless, for at least 1 minute.
18. Remove and retain 15 µl of eluate containing the DNA library in a clean 1.5 ml Eppendorf DNA LoBind tube. Dispose of the pelleted beads. Quantify 1 µl of eluted sample using a Qubit fluorometer.
The prepared library is used for loading into the flow cell. Store the library on ice until ready to load.
Materials:
Equipment:
Protocol:
1. Inoculate 300 uL of microcystis culture into 30 mL of BG11 media.*
2. Incubate at 25° C for optimal growth or at 20° C for optimal microcystin production at <3200 lux light, a 12:12 hour light/dark cycle, and consistent shaking and gas exchange.
3. *Transfer of microcystis culture should be done in a BSL-2 cabinet or setting.
Purpose: Assembling and amplifying gene fragments of unmodified and modified pSPDY plasmids.
Materials:
Equipment:
Protocol:
1. Create a 50 µL reaction by combining the following in a PCR tube:
2. Place the PCR reaction in the thermocycler and run a touchdown PCR protocol:
*Other methods and reagents for PCR may be used in place of Q5.
Protocol:
1. Thaw the Sequencing Buffer II (SBII), Loading Beads II (LBII) or Loading Solution (LS, if using), Flush Tether (FLT), and one tube of Flush Buffer (FB) at room temperature before mixing the reagents by vortexing and spin down at room temperature.
2. To prepare the flow cell priming mix, add 30 µl of thawed and mixed Flush Tether (FLT) directly to the tube of thawed and mixed Flush Buffer (FB), and mix by vortexing at room temperature.
3. Open the MinION lid and slide the flow cell under the clip. Press down firmly on the flow cell to ensure correct thermal and electrical contact.
4. Slide the flow cell priming port cover clockwise to open the priming port.
5. After opening the priming port, check for a small air bubble under the cover. Drawback a small volume to remove any bubbles:
a. Set a P1000 to 200 µL
b. Insert the tip into the priming port.
c. Turn the wheel until the dial shows 220-230 ul, to draw back 20-30 ul, or until you can see a small volume of buffer entering the pipette tip.
6. Load 800 µl of the priming mix into the flow cell via the priming port, avoiding the introduction of air bubbles. Wait for 5 minutes. During this time, prepare the library for loading by following the steps below.
7. Thoroughly mix the contents of the Loading Beads II (LBII) by pipetting. The Loading Beads II (LBII) tube contains a suspension of beads. These beads settle very quickly. It is vital that they are mixed immediately before use.
8. In a new tube, prepare the library for loading as follows:
Total: 75 µL
9. Complete the flow cell priming:
a. Gently lift the SpotON sample port cover to make the SpotON sample port accessible.
b. Load 200 µl of the priming mix into the flow cell priming port (not the SpotON sample port), avoiding the introduction of air bubbles.
10. Mix the prepared library gently by pipetting up and down just prior to loading.
11. Add 75 μl of the prepared library to the flow cell via the SpotON sample port in a dropwise fashion. Ensure each drop flows into the port before adding the next.
12. Add 75 μl of the prepared library to the flow cell via the SpotON sample port in a dropwise fashion. Ensure each drop flows into the port before adding the next.