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Team:Tuebingen/Protocols

PacMn

Protocols

To prepare fragments for Gibson Assembly® (Protocol 11), complementary overhangs had to be added to them. For this, the UCP HiFidelity PCR Kit (QIAGEN) was mostly used following the standard protocol (Table 1 and Table 2) provided in the kit. Fragments were either first analyzed by agarose gel electrophoresis (Protocol 2) or directly purified and concentrated (Protocol 3) for Gibson Assembly® reactions.

Table 1: 25 µl reaction setup for UCP HiFidelity PCR Kit
Component Volume
UCP PCR Master Mix, 2x 12.5 µl
Forward primer 0.625 µl = 0.25 µM
Reverse primer 0.625 µl = 0.25 µM
H2ONF 10.25 µl
Template DNA 1 µl = 10 ng
Table 2: UCP HiFidelity PCR Kit standard cycling conditions for amplicons ≤1 kb
Step Temperature Time
Initial PCR activation 98 °C 30s
3-step cycling (35 or 40 cycles)    
Denaturation 98 °C 10s
Annealing 3 °C above primer Tm (63 °C) 10s
Extension 72 °C 15s
Final extension 72 °C 2min

General:

Agarose gel electrophoresis typically served the analysis of PCR products or digested plasmid DNA (Protocol 6). Only 1% (w/v) agarose gels were prepared. Electrophoresis ran between 45-60 min at 100 V. The 1X Tris Acetate EDTA (TAE) buffer used for preparing the gels, for running electrophoresis, and for the DNA staining solutions, was a 1:50 dilution of self-made 50X TAE buffer (Table 1) in Milli-Q® water.

Table 3: Composition of 50X TAE buffer
Component Concentration
EDTA 50 mM
Tris base 2 M
Acetic acid 1 M

Staining:

During the first half of the wetlab work, gels were stained after electrophoresis in a solution containing 5 µL of SYBR® Gold Nucleic Acid Gel Stain (10,000X) (Invitrogen, California) from an old stock and 50 ml TAE buffer (1X).

In the second half, gels were stained before electrophoresis by adding SYBR® Safe DNA Gel Stain (10,000X) directly to the boiled agarose solution. For analytical gels, SYBR® Safe was added at a 6:100,000 ratio. For gel extraction (Protocol 10), SYBR® Safe was added at a 1:10,000 ratio.

DNA laddyers:

As a reference for DNA length, two different ladders were routinely used: BenchTop 1kb DNA Ladder (Promega, Wisconsin) and GeneRuler® 1kb DNA Ladder (Thermo Scientific).

Amplicons and other DNA products were purified (and concentrated) with the DNA Clean & Concentrator-5 kit (Zymo Research, Freiburg). The provided protocol was used, and elution volumes varied between 10-20 µl (case-specific).

According to Sambrook and Russel, 2001

For transformation, NEB® 5-alpha Competent E. coli (High Efficiency) cells were used. Because of the high transformation efficiency of these cells, volumes were split into 25 µl aliquots. The steps for transformation of 25 µl chemically competent cells were the following:

  1. Thaw chemically competent cells on ice
  2. Add DNA (1-100 ng; amount was case-specific) and mix gently by swirling the pipette tip and flicking the reaction tube. Incubate on ice for 20-40 min. While waiting, take out the required selective LB agar plates and leave them in the sterile bench (pre-warming), and pre-heat a heat block to 42 °C.
  3. After the incubation on ice, heat the tube at 42 °C for 45 s (heat shock) in the heat block. Place the tube back on ice to cool it down while you walk over to the sterile bench.
  4. In the sterile bench, add 900 μl of sterile SOC Outgrowth Medium (NEB), invert the tube 5 times, and incubate for 1 h at 37 °C (static).
  5. After incubation, take 100 µl of the cell suspension and spread it on a pre-warmed selective plate.
  6. Centrifuge the rest of the cell suspension for 3 min at 13000 rpm with a tabletop centrifuge (equal to ca. 17000 rcf). Discard the supernatant by tipping over the reaction tube and resuspend cells in the supernatant which remained in the tube (this is approximately 80-100 μL) .
  7. Spread the remaining cell suspension on a pre-warmed selective plate.
  8. Incubate the plates at 37 °C overnight.

For cloning and sequencing, DNA was isolated with the Monarch® Plasmid Miniprep Kit (New England Biolabs® Inc., Massachusetts), using the standard protocol provided.

This protocol is intended for screening of potential transformants, and not to open vectors or digest linear DNA for cloning experiments. For this reason, only one restriction enzyme (RE) is used per reaction, so that plasmid DNA is linearized and visualization in gel can be precise and consistent. The RE should be a unique cutter.

  1. Mix the following components (Table 4) together for one reaction (order: top → bottom):
    Note: For multiple samples, prepare a single “master mix” and then divide it into individual PCR tubes. Always calculate for one extra sample, so that you have enough volume in your master mix. The extra volume can always be used as a negative control (H2ONF)
    Table 4: Restriction digestion reaction for plasmid analysis
    Component Volume
    H2ONF 12.5 µl
    CutSmart® Buffer 2 µl
    RE 0.5 µl
  2. Add 5 µl DNA from a MiniPrep (Protocol 3) to a reaction, which is approximately 1-2 µg DNA
  3. Incubate (static) at 37 °C for 30 min (High Fidelity RE) or 1 h (normal RE)
  4. After digestion, visualize the DNA by gel electrophoresis (Protocol 2). For this, mix 4 µl DNA Gel Loading Dye (6X) (Thermo Fisher Scientific) with the 20 µl sample and load 7 µl of this mixture on the gel

For cloning experiments, pUC19 was isolated from E. coli transformants containing the vector in large amounts with the QIAGEN® Plasmid Plus Maxi Kit, using the standard protocol provided.

For colony PCR (Protocol 12) and addition of Gibson overhangs to fragment F, the AllTaq® PCR Core Kit (QIAGEN) was used, following the protocol provided in the kit (Table 5 and Table 6). This kit, although having a lower fidelity than the UCP HiFidelity PCR Kit, is more reliable when amplifying complicated fragments (F).

Table 5: 20 µl Reaction Setup for AllTaq Master Mix Kit
Component Volume
AllTaq Master Mix (4X) 5 µl
Primer forward 0.5 µl = 0.25 µM
Primer reverse 0.5 µl = 0.25 µM
H2ONF 13 µl
Template DNA 1 µl = 10 ng
Table 6: AllTaq cycling conditions for amplicons 1–9 kbp
Step Temperature Time
Initial PCR activation 93 °C 3 min
3-step cycling (40 cycles)    
Denaturation 93 °c 15s
Annealing Tm of primers 10s
Extension 68 °C 1 min/kb

This protocol is designed for opening a vector backbone isolated from E. coli for Gibson Assembly. In our case, pUC19 had to be opened with EcoRI and HindIII (NEB). To avoid re-ligation of pUC19, Calf Intestinal Alkaline Phosphatase (CIP) was added to the reaction mix.

  1. Mix the following components (Table 7) for a 50 µl reaction in order from top to bottom:
    Table 7: components
    Component Volume (µl)
    H2ONF 32
    CutSmart® Buffer 5
    Template DNA 10 (ca. 3 µg of DNA)
    Restriction Enzyme A 1
    Restriction Enzyme B 1
    CIP 1
  2. Incubate the mixture at 37 °C for 1 h and 30 min
  3. Purify the mixture (Protocol 3)
  4. Analyze results by agarose gel electrophoresis (Protocol 2)

Because most PCR products contained unspecific bands and intense primer clouds, gel extraction of the correct bands was necessary to improve Gibson Assembly® efficiency. Only purified and concentrated DNA samples were loaded for gel extraction. To create more defined bands, at least two teeth of the comb were taped together in such a manner, that at least two gel pockets formed a single larger pocket.

Once a gel piece was carefully cut and separated from the rest of the gel, it was turned on its side and trimmed. This improved the DNA-to-gel ratio. The rest of the extraction proceeded according to the protocol of the QIAquick Gel Extraction Kit (QIAGEN). After elution (30-50 µl), the eluted DNA was concentrated (Protocol 3) for Gibson Assembly®.

For the assembly of our constructs, the 4X Gibson Assembly® HiFi kit (Codex DNA, California) was used. The provided protocol was used but personalized slightly:

  • 50 ng of digested pUC19 (Protocol 9) was used in every reaction as the backbone
  • Unless specified otherwise, all fragments were added in a 3:1 molar ratio to the backbone except for fragment H, which was added in a 5:1 molar ratio due to its smaller size (ca. 300 bp)
  • From the 10 µl reaction (2.5 µl Gibson Assembly® HiFi HC master mix (4X) + 7.5 DNA mix), unless specified otherwise, 5 µl were used for transformation of 25 µl aliquots of NEB® 5-alpha Competent E. coli (High Efficiency)

Colony PCR was used to screen Gibson Assembly® transformants. For the PCR, the robust AllTaq® PCR Core Kit (QIAGEN) was used (Protocol 8), but adjusted for 10 µl reactions. Colonies were poked with a pipette tip, streaked on a selective sample plate, and then dipped inside the corresponding reaction tube. The M13 primers were used, which allow amplification of an insert in the multiple cloning site of pUC19. The extension time was adjusted according to the expected insert size. After PCR, 1-2 µl of Thermo Scientific® 6X DNA Loading Dye was added to each reaction tube and 4-6 µl of this mixture were loaded on a gel for analysis (Protocol 2).

Fluorescence was measured as a qualitative and quantitative test of our constructs.

Prerequisites:

  • Calibration of plate reader with iGEM silica microspheres at Abs660 (DOI: 10.1021/acssynbio.6b00072)
  • Calibration of plate reader with a fluorescent dye with approximately equal absorbance/emission spectrum as fluorogen
  • MnCl2 stock solution of 1 M (50 ml)
  • MnCl2 stock solution of 100 mM
  • MnCl2 stock solution of 10 mM
  • TFCoral /TFLime stock solution of 5 mM (vial + 50 µl DMSO)

Steps:

  1. Prepare 5 ml overnight cultures with LB medium and ampicillin (to 100 µg/ml)
  2. The next morning, roughly measure OD600 of cell cultures by mixing 250 µl of the cell culture with 750 µl of LB medium in a cuvette, and inoculate four times in 5 ml LB medium (100 µg/ml ampicillin) to an OD600 of ca. 0.09 with the following MnCl2 concentrations:
    1. 0 µM
    2. 10 µM
    3. 100 µM
    4. 1 mM
  3. Incubate the cultures at 37 °C, shaking, and measure the OD600 after 1 h, and then after 15-60 min, until an OD600 of 0.8 is reached. OD600 is always measured the same way (250 µl + 750 µl in cuvette)
  4. Harvest the cells
    1. Fill a 1.5 ml eppi with the cell suspension of a culture and centrifuge it at 17,000 rcf for 3 min. Discard the supernatant, fill it again, and repeat (4.5 ml in total)
    2. Add 1 ml PBS to the remaining cell pellets, resuspend, centrifuge, and discard supernatant. Repeat this step
  5. Prepare a 1 ml labeling solution for every three cell pellets: 1 µl of 5 mM TFCoral/TFLime + 999 µl PBS
  6. Resuspend a cell pellet with the 0.3 ml labeling solution, wait 10 s, fill three wells with 100 µl of this suspension, measure fluorescence immediately, then measure cell density

Reverse Transcriptase (RT)-PCR was used to verify the functionality of our Manganese-inducible promoter. For this, cell cultures were grown in selective LB medium overnight. The next day, the cultures were diluted 1:10 in 5 ml fresh selective LB media containing 10 µM Manganese(II) chloride and without Manganese(II) chloride as a control. Total RNA was extracted with the PureLink® RNA Mini Kit using the protocol version for bacterial cells. This RNA was used as a template for reverse transcription with the enzyme RevertAid H Minus RT, a recombinant M-MuLV RT. As a primer for the RT, we used the FAST_mid_RV primer, which binds in the FAST region of the mRNA. The synthesized single-stranded cDNA subsequently served as template DNA for robust PCR amplification (Protocol 8). Here, we used primers FAST_mid_RV and 15 (see Notebook). The amplicon (ca. 320 bp) was analyzed by gel electrophoresis (Protocol 2).