Team:UCSC/Results

Results

  • Results

  • Overview
  • LipLox
  • Yeast Mediated Cloning
  • Riboswitch
  • Quantification
  • Project Achievements

    • OVERVIEW

    We constructed the biosynthetic pathway for progesterone, consisting of 5 genes ( Δ7 Red, P450scc, FDX1, ADR, 3Β-HSD). In addition, we created a riboswitch that will allow us to measure the concentration of progesterone.

    LIPLOX

    Homology Arms

    To amplify out the HAs of Y. lipolytica str. FKP393, we lysed the cells and extracted the gDNA. We then performed touchdown PCR with the gDNA and confirmed the amplification of the HAs with gel electrophoresis. The UHA is 1014 bases long, and the DHA is 1000 bases long, so we expected bands at ~1 kb for both PCR samples. Figure 1, shows the expected bands just above 1 kb. It is likely that the bands are above 1 kb because the loaded samples contain Q5 reaction buffer and ran slower than the 2-log DNA ladder. We concluded that the bands produced were from the HAs. We sent the PCR products to be sequenced with primers that would identify regions of the homology arms and were able to confirm that our product were the HAs.

    Figure 1: Gel of Upstream and Downstream HAs, 1% TBE gel electrophoresis of the upstream and downstream HA PCR amplifications. We dyed each 5 µL sample with 1 µL of 6X loading dye and loaded 5 µL of this mix into each well. Lane 1 contains the 2-log DNA ladder, lane 2 contains the amplified UHA, and lane 3 contains the amplified DHA.

    Enter lox sites and ura3 into Y. lipolytica str. FKP393 genome

    YEAST MEDIATED CLONING

    We concluded the successful removal of loxP from pXRL2 via site directed mutagenesis. The results of our colony PCR suggested that the product of the reaction was slightly smaller than the original plasmid. However, the positive control had a strange band higher than we expected. We sequenced to confirm our success and found that the product was missing the desired region in an alignment with the original plasmid. We also conducted attempts of yeast mediated cloning with successful growth in leucine deficient media. This result would suggest that we had a successful transformation and possible assembly of our gene cassette on to the plasmid backbone.

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    We conducted a colony PCR to confirm the successful site directed mutagenesis of loxP from pXRL2. We used a pair of primers: cloxP F and cloxP R to amplify the region that we removed and ran the samples on a gel. A successful trial would be a band 300 bp or lower since the fragment including the lox site would be 343 bp. Our results were puzzling due to the band being higher than 300 bp but we thought that this had occurred because the sample ran slower than the other lanes evident in the arc of the dye. We conducted another colony PCR with monoclonal colonies. Strangely, our positive control appeared to be much larger than the other samples but we concluded that it was due to issues with gel electrophoresis. We resolved to use the samples with the lowest bands or smallest size which were 5C, 5D, 21B and 23A. Upon further investigation by sequencing, we concluded that we had succeeded due to the region being missing in an alignment with the original plasmid. However, in sample A, more bases were deleted than expected so we did not use it because of risk of a frameshift. We, then relinearized the four sequenced samples for gene assembly via yeast mediated cloning. The results of this suggested a success since the bands resulting from gel electrophoresis were at the expected 8-10 kb.

    GIBSON

    The Riboswitch team has been using a plasmid called pD17, generously gifted from Cory Schwartz at UC Irvine, which contains homology arms for the pseudogene d17 in Y. lipolytica. We decided to try and insert loxP-URA3-lox71 into that plasmid. If successful, we would then amplify out to obtain a linear construct including the homology arms to d17, which we could perform homologous recombination into the Y. lipolytica genome with. We PCR amplified pD17, cutting out plasmid backbone between the homology arms to create a construct with homology arms at each end of the linear product. We ordered primers with Gibson Overhangs (GO) to the pD17 plasmid to amplify loxP-URA3-lox71 with. The PCR results were unsuccessful every try. Due to these issues, we decided to use KLD to insert the loxP-URA3-lox71 gene block into pD17. We performed KLD treatment with linearized pD17 backbone, and the loxP-URA3-lox71 gene block and transformed NEB 5-alpha E.coli. We saw many colonies, and we picked 15 for colony PCR to check if integration occurred.

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    The gel shows in lane 2 that we achieved a clean, single band at ~5kb. The primers bound to the pD17 plasmid at the beginning and end of the d17 homologous arms, including the arms and our loxP-URA3-lox71 insert. The expected size of our product is 3.6 kb. The band is slightly higher than this size, which we believe is due to the salts in the cell causing it to run a bit slower than it should. Because of this we are confident that this band is our desired product. We have grown a liquid culture of Colony 2, miniprepped it, and performed PCR using Q5 polymerase and the same primers used for colony PCR to attempt to amplify out loxP-URA3-lox71 sandwiched between the d17 homology arms. The results have not yet shown any bands from this.

    Once we are able to amplify loxP-URA3-lox71 with d17 homology arms, we will transform Y. lipolytica with this linear product and confirm success by growing the colonies on uracil-deficient media. Once the loxP and lox71 sites are integrated into the genome we can perform Recombinase mediated cassette exchange with the plasmid created by YMC with all pathway genes using the corresponding lox sites.

    RIBOSWITCH

    For our part of the project, we intended to create a one-gene reporter system for progesterone concentration consisting of: a Renilla GFP gene (hrgfp), a cyc1 terminator, TEF(136) promoter, and a hammerhead ribozyme expression platform attached to a progesterone-specific aptamer.

    Our first task in this project was to create a large stock of our pHR_D17_hrGFP plasmid (from here on referred to as D17), which we did by transforming a portion of our IDT D17 stock into chemicompetent E. coli, running a selection on an LB agar plate with a 100 ug/L concentration of ampicillin, and then growing out selected colonies in liquid LB medium. We then isolated our samples of D17 from the bacterial culture by using a Zymo plasmid miniprep kit.

    In order to confirm the sequence of our hrgfp gene as well as the cyc1 terminator, we also designed a series of primers meant to be used in Sanger sequencing of our D17 plasmid. These primers were labeled Rib(f)-1 through Rib(f)-3 and Rib(r)-1 through Rib(r)-3 and encompassed the entire hrgfp gene as well as the cyc1 terminator with several hundred base pairs of overhang on either side. Sequencing data from the UC Berkeley DNA sequencing facility showed a successful primer design and an intact DNA sequence in our D17 plasmid.

    Figure 1: Agarose gel confirming the presence of our linearized D17 plasmid at around the 9kb mark.

    In order to carry out Gibson Assembly, we also had to amplify our stock of riboswitch inserts. To do this, we designed the Apt(f) and Apt(r) primers to anneal to the overhangs of all 5 different inserts. Running a series of PCR trials and imaging on an agarose gel in Figure 2 showed strong bands at the expected bp length, indicating successful amplification of our inserts.

    Figure 2: Agarose gel confirming the amplification of riboswitch oligomers on lanes 2 through 5.

    We then performed a Gibson assembly trial with our newly amplified D17 vector and riboswitch inserts. We attempted a confirmation PCR to check if our inserts were successfully integrated into the D17 backbone with the ampGB(f) and ampGB(r) primers. A successful integration event would result in amplicons approximately 4.2 kb in length, while a circular D17 template would create an amplicon 4.0 kp in length. Imaging of our gel in Figure 3 suggested the continued presence of D17 circular template along with our new pOPPY_GFP plasmids, even after digestion with >500-fold excess of DpnI.

    Figure 3: Agarose gel comparing the presence of linear vs. circular D17 template amplification after Gibson assembly.

    To remedy this, we performed a series of transformations and selective platings with our Gibson assembly product on 100 ug/mL LB-ampicillin medium and selected a total of 52 colonies--10 from each Gibson transformation and 2 from a D17 control plate--for colony PCR and spot plating on an index plate. The imaged gels in Figure 4 from the colony PCR trials allowed us to select single colonies we believed to be carrying our riboswitch insert.

    Figure 4: Agarose gels of index plate colonies 1-52, 1-10 of pOPPY_GFP 1, 11-20 of pOPPY_GFP 2, and so on. Colonies 51 was our negative D17 control. We looked for a 200 base pair difference in each pOPPY_GFP variant to confirm presence of our riboswitch insert.

    From these gels we selected colonies 2, 16, 27, 32, 43, as likely carriers of pOPPY_GFP(1)--pOPPY_GFP(5), respectively, and colony 51 as a control carrier of D17. We then used our previously grown index plate to create monoclonal liquid cultures of these colonies in LB medium. After overnight incubation, we then performed a plasmid isolation with all 6 cultures and transformed the isolated plasmids into our competent Y. lipolytica strain, which we created using the Zymo Frozen EZ-yeast transformation protocol.

    QUANTIFICATION

    We constructed the biosynthetic pathway for progesterone, consisting of 5 genes ( Δ7 Red, P450scc, FDX1, ADR, 3Β-HSD). In addition we created a riboswitch that will allow us to measure the concentration of progesterone.

    PROJECT ACHIEVEMENTS





    Designing PCR Primers

    Our teaching assistant, McKenna Hicks, taught us the following methods for primer design.

    1. Identify the DNA sequence to amplify or stitch together with overlap extension PCR (OE-PCR). During PCR, primers anneal to their complementary sequence and recruit DNA polymerase to extend the sequence in the 5' to 3' Direction.
    2. Design a forward (5'- 3') and a reverse primer (3'- 5') to amplify both the top and bottom DNA strands. The ideal primer length is between 18 and 23 bp, but this can be changed to accommodate other constraints.
    3. For the forward primer, select a sequence of base pairs (bp) of optimal length.
    4. Repeat Step 3 for the reverse primer and generate the reverse complement of the reverse primer sequence.
    5. Analyze each sequence using a computational primer tool (such as IDT's Oligo Analyzer) and record the GC content and the melting temperature (Tm) for each primer.
    6. The optimal GC content is between 40-60%, and the optimal Tm range is between 52-62℃. Ideally, the difference in Tm for the forward and reverse primer pair should not exceed 5℃.
    7. If the GC content exceeds 60% for either primer, add High GC Enhancer to the PCR reaction mix.
    8. Check that the free energy (ΔG) value for self-dimers and hetero-dimers on each primer is above -10 kcal/mol to avoid primers spontaneously self-annealing or annealing to each other.
    9. Check that hairpin formations do not exceed 30℃ in each primer. PCR uses different temperatures for each thermocycler step; the lowest temperature used during the replication step for primer annealing is 58-60℃. If hairpins form around this temperature, the primer binding efficiency may be reduced and yield less product. However, if the primer Tm is above the hairpin formation temperature, and if the potential hairpin leaves the 3' end of the primer free for annealing, the primer should still anneal properly without reducing PCR Efficiency.

    E. Coli Miniprep

    The following protocol was adapted from the ZyppyTM Plasmid Miniprep Kit procedure from Zymo Research.

    1. First prepare your cell samples: centrifuge the cell cultures (which should be 5 mL in conical tubes) for 15 minutes at 3,750 rpm and 30℃.
    2. During waiting time, grab a 250 mL beaker and add roughly 50 mL of 10% bleach. Grab some foil, label waste, and cover the top of the beaker. Use this beaker to discard biohazardous supernatants and flow-throughs throughout your miniprep.
    3. Collect Zyppy columns, Zyppy collection tubes, and 2 sets of 1.5 mL microcentrifuge tubes to match the number of samples you centrifuged. Place the columns over the collection tubes. Be sure to label all vessels accordingly. Place a kimwipe over the open columns.
    4. After centrifugation, discard the supernatants into your waste beaker. Be sure to replace the foil lid afterwards.
    5. Add 600 uL Milli-Q water into each of the conical tubes containing the cell pellets. Resuspend each by slowly pipetting up and down; use that same pipette to transfer the resuspended mixture to its respective clean microcentrifuge tube. Change tips between samples.
    6. Add 100 uL of 7X Lysis Buffer (blue) and mix by inverting the tube 4-6 times. Proceed to next step within 2 minutes. After addition of 7X Lysis Buffer, the solution should change from opaque to clear blue, indicating complete lysis.
    7. Add 350 uL of cold Neutralization Buffer and mix thoroughly. The sample will turn yellow when the neutralization is complete and a yellowish precipitate will form. Invert the sample an additional 2-3 times to ensure complete neutralization.
    8. Centrifuge at 13,000 rpm for 3 minutes.
    9. Transfer the supernatant into the provided Zymo-SpinTM IIN column, which should be on top of a collection tube. Avoid disturbing the cell debris pellet.
    10. Centrifuge the column + collection tube for 15 seconds at 13,000 rpm (stop the centrifuge when it reaches 15 sec).
    11. Discard the flow-through and place the column back into the same Collection Tube.
    12. Add 200 uL of Endo-Wash Buffer to the column. Centrifuge for 30 seconds at 13,000 rpm. Discard the flow-through.
    13. Add 400 uL of ZyppyTM Wash Buffer to the column. Centrifuge for 1 minute at 13,000 rpm. Discard flow-through. Be sure to close cap tightly on wash buffer so that ethanol does not evaporate.
    14. Transfer the column into a clean 1.5 mL microcentrifuge tube then add 50 uL of Milli-Q water in place of Elution Buffer (EB) and add it as close to the filter as possible, being careful not to touch it (use Milli-Q instead of EB to prevent contamination from salts and EDTA). Let stand for at least 10 minutes at room temperature. Incubate longer if needed.
    15. Centrifuge for 30 seconds at 13,000 rpm to elute the plasmid DNA.