We want to make two plasmids:
Plasmid #1 - A constitutive promoter with LacZ and LacY.
Plasmid #2 - A lactose inducible promoter with lysozyme and holin.
The general idea is to overexpress LacY (Lactose transporter) with LacZ (Beta-Galactosidase). When lactose is present, the cell will uptake lactose and will flip on the Lac promoter repressed by lacI when enough lactose is present. We then express lysozyme and holin which will lyse the cell releasing beta-gal cleaving lactose into glucose and galactose. Glucose and galactose will then be taken up by the host in the colon.
We have decided to use the I732017 LacZ (1009/11H) because we could not transform I732005 LacZ (1018/6F).
To double check, we will plate our transformed I732005 LacZ with 100μL onto a kan plate, and the rest on an Amp plate.
We ordered primers for LacY, and we will need to PCR out LacY. We then need to run site directed mutagenesis on LacY by added 2H to prevent EIIA(GLUC) binding.
For our constitutive promoter, we will use various strengths to test our beta-gal concentrations.
Low expression - J23113
Medium expression - J23106
High expression - J23100
We need to cut out mRFP, which is originally in the promoter plasmid family of J23100, by doing a digest with SpeI and PstI. We will paste in LacZ and LacY, and add a double TT.
Colonies grew from I732017 LacZ transformation. They will be cultured Sunday afternoon.
LacY, LacY +H, and LacY +2H has been inputted into the registry of parts. Primers for LacY site directed mutagenesis were designed.
When primers for LacY arrive, need to PCR LacY out of E. coli genomic DNA.
After LacZ culture is grown, mini prep DNA.
Set up o/n cultures.
1. LacZ in Amp
2. pSB2K3 in Km
3. p1005 in Amp
4. p1005 in Amp + Tet (Just to test the strain is both Tet and Amp resistance)
Mini prep'd LacZ and pSB2K3, eluted in 50 μL.
LacZ conc. = 115.4 ng/μL
pSB2K3 conc. = 427.4 ng/μL
Ran a PCR to amplify LacY out of E. coli genomic DNA (hoping that E. coli genomic DNA has LacY wild type).
1 ul PFU Ultra II, 1 ul E. coli genomic DNA, 2 ul (Primer FWD), 2 ul (Primer RVS), 10 ul 10x PFU Buffer, 2 ul dNTP's, and 82 ul dH20.
PCR's did not work first time. Ran another PCR in 50 ul with 5 ul of template as the biggest revision. Annealing temp's were lowered by 4°C, and extensions were lengthened another 15 seconds.
Transformed 2 RBS's (Weak and weaker) b0031 and b0033, and LacY J22101. Will check tomorrow for grwoth.
Set up digests for LacZ and LacY.
1. Cut LacY XP
2. Cut RBS34 SP
3. Cut RBS32 SP
4. Cut High Copy Plasmid XP
5. Cut LacZ ES
6. Cut TT EX
Transformations of the 2 RBS's B0031 and B0033 was successful. The LacY J22101 did not look so good, so the plate was tossed.
Only 2,3, and 5 worked. Gel extracted them and they were put in the digest box. Concentration was not measured.
Ran another LacY PCR with template from previous LacY purification.
The gel from the PCR looked OK, so digested LacY, pSB1A2, and TT again.
Treated digest backbones with CIP, and heat inactivated.
Miniprep'd transformations from last night, B0031 and B0033 (Weak RBS's).
B0031: 113 ng/ul
B0033: 120 ng/ul
Ran last nights digests on a gel and gel extracted them.
Set up 6 ligations and transformed them.
1. LacY + RBS34
2. LacY + RBS32
3. LacY + pSB1A2
4. Control: RBS34
5. Control: RBS32
6. Control: pSB1A2
Also transformed a Control for the NEW LB+Amp plates with just D10HB. The Amp will kill off the cells if the plates were poured correctly.
The transformed plates didn't look good. On the experiment plates 1-3, there were MANY colonies on the edges of the plates, but the middle of the plate had streakyness of something. On control plates 4-6, there were no colonies, but there was the same streakyness all across the plate. A few thing could have gone wrong such as the transformation protocol, the LB + Amp plates, or the HB cells. We will see how the colony PCR turns out for plates 1-3. We took 6 colonies from plates 1 and 2, and 1 colony from plate 3.
Colony PCR Results: For plate 1, we wanted to see a band of about 1.6KB with LacY + RBS, and we saw 6 bands for the six colonies at that marker. The bands were fairly light, so we may have not have had as many cells, but it hopefully worked! For plate 2, it should show the same size band at about 1.6KB and it was very dark. We are hoping that both RBS + LacY worked, and they will be grown o/n tonight, and prep'd tomorrow.
To Do List:
1. RBS LacZ - Already have
2. Terminator - transform Fri, o/n Sat, Prep + Cut Sunday
3. B0031/33 (Weak RBS's) - Cut Sun
4. B0032/34 LacY - o/n Fri, prep Sat, cut Sun
5. pSB1A2 p1010(ccdb) - cut Sun
6. pSB1A2 LacY - retransform Fri, two LB + Amp plates from upstairs, one w/ 1/10 dilution (10ul of cells + 90LB), and the rest concentrated cells of 100ul.
7. Break glass vial with freeze dried cells - Sun... :-)
Transformation of LacY+pSB1A2 had colonies. Will colony PCR them soon and check for the band.
Miniprep of B0034/32 LacY will be done on Sunday. Forgot to save the supernatant after 10 minute spin. It will not delay us a day, which is good, since we plan to cut on Sunday anyway.
Colony PCR'd the LacY+pSB1A2, although it doesn't seem probable that it worked. Will run LacY PCR on the same gel to check if that worked too.
Made new o/n cultures of B0034/32 LacY due to error made today.
Transformed B0015 Terminator.
Set up minipreps of B0034/32-LacY, B0015 Term, and pSB1A2-LacY.
PCR purified LacY
Set up digests of:
1. B0015 EX
2. B0031 SP
3. B0033 SP
4. B0032-LacY ES
5. LacY XP
6. pSB1A2-LacY ES
Sent in sequencing for pSB1A2-LacY and B0034/32 LacY.
Gel extracted digests, set up ligations and transformed for:
1. LacZ + B0015
2. RBS32-LacY + B0015
3. LacY + B0031
4. LacY + B0033
5. pSB1A2-LacY + B0015
6. CONTROL B0015
7. CONTROL B0031
8. CONTROL B0033
Lysis cassette primers arrived.
Transformations looked good. We used the less competent cells from upstairs instead of D10HB, so I decided to spin down and plate all cells. All plates had colonies, background from controls were less than ligations, although B0031 had about the same background. Set up colony PCR's and ran on a gel.
Colony PCR's FAILED! Time to start over.
The Plan for now since we have LacY in pSB1A2:
1. LacY --> B0031/33/15
2. LacY --> SDM and primer design
3. LacZ --> B0015
4. Lysis cassette --> 1A2
PCR'd the lysis cassette out of Lambda Zap. Two bands showed, one at 1KB and 1 at about 1.3KB. The band at 1.3KB was much darker and that is what we expected our size to be.
Happy Fourth of July!
PowerPoint for 7/9/08
iGEM Meeting #3
The past few days have been failure!
Tried to re-clone LacY into B0015, B0031/33 and LacZ into B0015. It failed, and re-did the digests tonight. The analytical digest looked good, so hopefully we can gel extract clean bands. We are also working on the lysis cassette to place into pSB1A2 which I ligated and transformed tonight. Hopefully there will be little background from the control. If that looks good, we will screen with colony PCR tomorrow.
Transformation of Lysis Cassette + pSB1A2 looked beautiful. The control plate had 0 colonies while the ligation had a fair amount. Ran a colony PCR of 9 different colonies + a control of 1A2 prep. There were 4 bands at about 1.5KB (what we expect since lysis = 1.3 + VF VR = .2) and 3 bands had nothing. 1 band was VERY flooded, so something may have gone weird in that lane. We hopefully reached SUCCESS!
Will grow o/n cultures of two colonies that looked good and will send in for sequencing on Monday morning.
The digest of all backbones were gel extracted and the ligations will be carried out today.
LacY into 31/33, and LacZ into 15, for a total of 3 reactions and 3 controls.
Sucess!! The ligations had about a couple hundred fold (if not more) over the control plates. I will screen with colony PCR today and if they look good, I'll send them in for sequencing along with the lysis cassette and SDM LacY.
Colony PCR's looked beautiful! Will grow 2 cultures o/n and send 1 in for sequencing.
Sequencing for Monday:
1. SDM LacY (2 colonies)
2. Lysis Cassette + 1A2 (2 colonies)
3. LacY + 31/33 (2 colonies each)
4. LacZ + 15 (2 colonies)
Sequencing preps have been set up and will be delivered Monday morning