Purpose:

Design a putrescine-responsive circuit in E. coli

Methods:

We designed the genetic circuit shown below. This circuit constitutively expresses PuuR, a protein that binds to the pHub(1B) inducible promoter. When putrescine is present, the repressor protein PuuR is stopped and eGFP is produced. mCherry is included downstream of PuuR to ensure expression of the PuuR protein.

We obtained sequences for the PuuR repressor protein and Phyb1B promoter from the literature (Selim et al., “A Synthetic Biosensor for Detecting Putrescine in Beef Samples.”). We chose to use a plasmid with mCherry (Addgene #68142) as our backbone, to shorten the length of the gBlock we would order. The sequences were all imported into SnapGene for plasmid design. The assembly was simulated using the SnapGene HiFi tool. The backbone was cut with HindIII and XbaI and the following insert was cloned. Homology regions were designed for each end of the insert, with the following specifications: melting temperature of 60 degrees C and 20-30 base pairs long. Cut sites on the backbone and exposed ends on the insert were checked for hairpins and altered such that no hairpins formed above 50 degrees C.

We also designed a similar insert, with the only change being swapping a T7 constitutive promoter instead of the pHyb(1B) inducible promoter. The purpose of this design was to observe GFP expression, and to have a measure to quantify the inducible promoter expression against.

Results:

Finalized simulated assembly of the gBlock into the mCherry backbone.

Notes:

We initially ran into challenges choosing cut sites on the backbone, as hairpins were predicted to form above 50 degrees C. This caused us to pivot to using different enzymes than we initially planned.

Purpose:

Order gBlocks

Methods:

We finalized the insert designs and ensured the complexity was appropriate for IDT to synthesize.

Results:

Ordered 2 gblocks from IDT, the mCherry backbone from Addgene, and restriction enzymes from NEB

Purpose:

Prepare for plasmid assembly and cloning.

Methods:

Streaked out plates with mCherry.

Notes:

We noticed that the agar stab delivered by Addgene had both green and pink bacteria growing. The plate that was streaked out also had pink and green colonies. See images below.

Purpose:

Prepare for plasmid assembly and cloning.

Methods:

We made overnights of the mCherry backbone, miniprepped, stocked, and sent for sequencing. We miniprepped using the “low-copy” protocol for better results.

Results:

Initially the concentration of DNA after miniprepping was less than 100 ng/uL, which was lower than we expected for a high-copy plasmid. The 260/280 ratio was within the normal range of 180-2, so we didn’t think contamination was an issue. After switching to the low-copy miniprep protocol we obtained eluted DNA with a concentration above 120 ng/uL.

Notes:

The sequencing results were perfect except for a 1 bp mutation in the origin of replication.

Purpose:

Clone the inserts into the backbone.

Methods:

We performed a restriction digest on the isolated mCherry backbone DNA. After rehydrating the gblocks with our DNA inserts, we performed HiFi assembly and transformed the pHyb(1B)-plasmid into NEB Dh5-α.

Results:

Correct sequencing results for the pHyb(1B)-plasmid in NEB Dh5-α.

HiFi Assembly

Purpose:

Clone the constitutive-insert into the mCherry backbone.

Methods:

We performed HiFi assembly and transformed the constitutive-plasmid into NEB Dh5-α.

Results:

Correct sequencing results for the constitutive-plasmid in NEB Dh5-α.

Transformation into BL21(DE3)

Purpose:

It was necessary to transform the plasmids into BL21 (DE3) because the design includes T7 components, which require T7 RNA polymerase (only in BL21).

Methods:

We followed the protocol for transformation into ThermoFisher BL21 (DE3) competent cells.

Results:

Transformation was unsuccessful with the ThermoFisher competent cells, and the puc19 positive control showed no growth.

Purpose:

Alter the transformation protocol to obtain successful transformants.

Methods:

We switched to NEB BL21 (DE3) competent cells and followed the protocol below.

The initial transformation showed one constitutive-plasmid colony, which we cultured, miniprepped, and sequenced. In order to transform the inducible-plasmid, we altered the NEB transformation protocol by heat-shocking for 30 seconds at 40 degrees C.

Results:

The sequencing of both plasmids came back with perfect results.

Purpose:

Gain insight into the growth mechanics of our plasmid.

Methods:

We performed a growth assay on both the plasmids, as detailed below

1. Grow overnights in culture tubes with 5mL LB and 5uL of ampicillin. Incubate at 37 C for 15-18 hours.
2. Measure OD600.
3. Spin down culture such that OD = 1 in a volume of 1mL. Resuspend the pelleted cells in 1mL of LB + ampicillin.
4. Dilute the resuspended cells at 1:100 in LB + amp, for a starting OD of 0.01
5. In a 96 well plate, add 200uL of diluted resuspended cells per well in triplicate.
6. Measure OD600 every hour, with the plate reader at 37C, shaking 800rpm, for 22 hours.

Results:

There were differences in growth between the three plasmids. This could be due to differing metabolic burdens on the cells.

Dose Response: IPTG

Purpose:

Determine the optimal concentration of IPTG for induction.

Methods:

1. Grow overnights in culture tubes with 5mL LB and 5uL of ampicillin. Incubate at 37 C for 15-18 hours.
2. Measure OD600.
3. Spin down culture such that OD = 1 in a volume of 13mL. Resuspend the pelleted cells in 1mL of LB + ampicillin.
4. Incubate at 37C until it reaches an OD of ~1 (~4hours). Measure OD each hour.
5. In a 96-well black-wall clear-bottom plate, induce cultures with varying concentrations of IPTG {0, 0.1, 0.3, 0.5, 0.7, 1} mM.
6. Incubate for 18 hours shaking at 37C in a plate reader. Measure OD600, GFP, and RFP fluorescence every 20 min.

Results:

We determined 0.5mM to be the optimal concentration of IPTG to use for future testing. See the experiments page for more details.

Dose Response: Putrescine

Purpose:

Determine the response of the inducible promoter to exogenous putrescine.

Methods:

1. Grow overnights in culture tubes with 5mL LB and 5uL of ampicillin. Incubate at 37 C for 15-18 hours.
2. Measure OD600.
3. Spin down culture such that OD = 1 in a volume of 13mL. Resuspend the pelleted cells in 1mL of LB + ampicillin.
4. Incubate at 37C until it reaches an OD of ~1 (~4hours). Measure OD each hour.
5. In a 96-well black-wall clear-bottom plate, induce cultures with 0.5mM IPTG and varying concentrations of putrescine {0, 10, 100, 500} mM.
6. Incubate for 18 hours shaking at 37C in a plate reader. Measure OD600, GFP, and RFP fluorescence every 20 min.

Results:

Notes:

There was significant clumping of cells observed with high concentrations of putrescine.

Putrescine Dose Response 2.0

Purpose:

Avoid clumping of cells during the dose response assay.

Methods:

Same protocol as above, but use new varying concentrations of putrescine {0, 1, 10, 50} mM.

Results:

Notes:

There was much less, but still some visible clumping of cells observed with higher concentrations of putrescine.

Validation Test 1.0

We performed the Validation Test, first version, as described on the Experiments page.

Validation Test 2.0

We performed the Validation Test, second version, as described on the Experiments page.

Genetic Design

We designed an alternative gblock that replaces the T7 promoter with a constitutive, non-inducible promoter (pVeg1). Based on our experimental results, we hypothesized that IPTG was having an unwanted influence on our biosensor. By removing the T7 promoter, we eliminate the need to induce the cells with IPTG, thus simplifying the genetic system. Additionally, this new design will always repress GFP, ensuring that GFP is only turned on when putrescine is present. This will allow for tighter control of GFP, and thus stronger and more reliable quantification of putrescine.

Validation Test 3.0

We performed the Validation Test, third version, as described on the Experiments page.

IPTG Dose Response in M9 Media

We performed an IPTG dose response in M9 media, as described on the Experiments page.

Group Meeting:

During our group meeting this week, we presented confusing experimental data we collected to Professor Jimenez. He noticed that our plasmid included two additional promoters that could be influencing our system unintentionally: a LacUV5 promoter upstream of the PuuO synthetic promoter, and a reverse T7 promoter downstream of the mCherry CDS.

We developed a plan to cut our plasmid to eliminate these additional promoters.

1. Fix V002 (mCherry backbone with V004 design in DH5a).
1. To do this, we first need to cut out the lacUV5 promoter with NspI enzyme. This will result in the plasmid shown below.
2. Second, we need to remove the reserve T7 promoter. To do this, we need a few things: NotI enzyme, DraIII enzyme, and Quick Blunt kit.
2. Clone the new gblock into the backbone, but without the bad promoters.
1. We will cut out the LavUV5 promoter and reverse T7 promoter with the same method as above. This will result in a “clean” backbone to clone into.
2. Use HiFi assembly to clone the new gblock (with pVeg1) into the “clean” backbone.

After these steps are complete, we plan to perform more experimental testing to determine if changing the promoters will improve the response of our biosensor.

Plates:

We made plates of our backbone (mCherry), V002 (PuuR/PuuO system in Dh5a), and V004 (PuuR/PuuO system in BL21).

From these plates, we made overnights to begin the process of cutting out the promoters.

Miniprep:

Colonies we miniprepped:

• V004.1
• V004.2
• V002.1
• V002.3
• V002.. → from a good colony
• V002 big → from a big colony
• V001.1
• V001.2

We obtained high yield from V002 and V004 colonies. V001 did not produce high enough yield to continue through the process of digestion.

Digest with NspI:

We followed the lab protocol for restriction digestion.
Notes:

• Digest V001.2 and V002.big and V004.2.
• Run at 37 C for 30 minutes.

After the digestion, we added loading dye and ran it on a gel to separate the linearized backbone DNA from the insert that was cut out.

Preparative Gel

After the digestion, we added loading dye and ran it on a gel to separate the linearized backbone DNA from the insert that was excised.

DNA Cleanup

We performed a cleanup on the DNA to extract the DNA from the gel. We selected the highest-yield sample to continue with for a ligation. We performed the ligation twice in parallel, one with Hi-T4 quick ligase, and the other with T4 ligase, which requires overnight incubation. We hope to compare them and select the best result.

1. Quick
1. Ligate with-Hi T4 Quick Ligase using the NEB protocol for Hi-T4™ DNA Ligase (NEB #M2622)
2. Add 50ng of DNA for a final concentration of 2.5ng/uL
   1. Concentration of DNA from cleanup: 137.87 ng/uL
   2. 2.5 ng/uL * 20 uL = 137ng/uL * x → 0.36uL
3. Put on thermocycler
1. 10 min 25 C
2. 10 min at 65C
3. Hold at 4
4. Transform

Components to combine:

1. 2uL buffer
2. 0.36uL DNA
3. 16.64 NFW
4. 1 uL Hi-Ligase

2. Slow
1. Ligate with T4 Ligase using the NEB protocol for Ligation Protocol with T4 DNA Ligase (M0202)

Components to combine:

1. 2uL buffer
2. 0.36uL DNA
3. 16.64 NFW
4. 1 uL (normal) Ligase

Transformation of Quick Ligase

1. Only transform Hi-T4 Quick Ligated DNA
2. Need 4 tubes of comp cells (DH5a): C2987
• One for pUC19
• One for V002_qx outgrowth
• One for V002 outgrowth
• One for V002_minusLigase outgrowth
3. Samples to plate: total of 13 plates
• Puc19 : transformation control
• V002_qx (1x) 100uL plated
• V002_qx (3x) 100uL plated
• V002_qx (3x) 100uL plated
• V002_qx (3x) 100uL plated
• V002 (1x) 100uL plated
• V002 (3x) 100uL plated
• V002 (3x) 100uL plated
• V002 (3x) 100uL plated
• V002_minusLigase (1x) 100uL plated
• V002_minusLigase (3x) 100uL plated
• V002_minusLigase (3x) 100uL plated
• V002_minusLigase (3x) 100uL plated

Steps for making 3X DNA

1. Plate 100uL of 1X (normal DNA)
2. Spin down for 1 min the remaining 900uL of DNA in SOC media
3. Discard 600uL of media
4. Resuspend
5. Plate 100uL on 3 plates

Quick Ligase Transformation Results

We transformed V002_2 (which is V004 in Dh5alpha, missing the LacUV5 promoter). We then picked 8 samples to send for sequencing, and 6 out of 8 came back with correct sequences.

In the upcoming weeks, we will continue with the plan listed above to alter our genetic design with the aim of increasing the desired response of our biosensor.

We plan to push two genetic design plans forward:

1. Edit our current biosensor. This will include the following steps:
   1. Digest with NotI and DraIII
   2. Use blunting kit
   3. Ligation
   4. Transformation into DH5α
   5. Miniprep
   6. Transformation into BL21

1. Insert the new gBlock into the original backbone. This will include the following steps:
   1. Miniprep successfully
   2. Digest with NotI
   3. Gel
   4. Cleanup
   5. Sticky end ligation
   6. Transformation into DH5α
   7. Miniprep
   8. Digest with NotI and DraIII
   9. Blunting kit
   10. Ligation
   11. Transformation into DH5α
   12. Miniprep
   13. Transformation into BL21

Once these plans have been accomplished, we will perform testing on our new plasmid designs and compare to our previous results.