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From 2008.igem.org

July 22nd

We have 4 tubes from last year. These strains are frozen in glycerol.

• PNZ8901 plasmid in E.coli MC1061 Strain
  • Chloroamphenicol resistant
  • "sure" vector
  • does not integrate

• B. subtilis strain 1A1
  • No resistance
  • Same as strain 168 but tryptophan deficient

• E.coli strain MC1061, no plasmid

• PP182 plasmid in E.coli strain DH5alpha
  • Ampicillin resistance
  • integrates

Checking antibiotic resistance

Purpose : Grow each of them on a plate to test their antibiotic resistance

Protocol

• Warm up frozen tubes
• Take 15uL of each and put it on plates without antibiotic
• Incubate at 37°C

July 23rd

• Take one colony from each of the plates grown from the tubes yesterday and plated them again on LA without antibiotics

1/ Purpose Check antibiotic resistance of our different strains

Strain	Plasmid	Extracted from	Antibiotic use	Concentration of antibiotic
DH5alpha	PP182	tube	Amp	100
MC1061	NONE	tube	Cm	35
MC1061	PNZ8901	tube	Cm	35
DH5alpha	PP182	plate	Amp	100
MC1061	NONE	plate	Cm	35
MC1061	PNZ8901	plate	Cm	35

2/ Purpose : Find the right concentration of antibiotic so that B. subtilis survive

- Grow 15uL of B. 1A1 (frozen tube) in 5mL of LB

- Incubate at 37°C

3/ Purpose : Grow plasmids in TOP10, transformation

4 plasmids :

• I746000
• I746100
• I746101
• I746001

1 control : PUC19

- Add 20uL of TOP10 competent cells and 0.5uL of Plasmid in an eppendorf

- 30min on ice

- Heat shock : 60s at 42°C

- 2min on ice

- Add 89.5uL of SOC

- 60min at 37°C

- Put the mix on plate with ampicillin resistance

- Incubate at 37°C

July 24th

• Test of antibiotic resistances of strains of last year

Strain	Plasmid	Extracted from	Antibiotic use	Observations	Conclusion
DH5alpha	PP182	tube	Amp	Many colonies	Amp Resistance OK
DH5alpha	PP182	plate	Amp	Many colonies	Idem
MC1061	NONE	tube	Cm	Nothing	As expecting, no Cm Resist.
MC1061	NONE	plate	Cm	Nothing	Idem
MC1061	PNZ8901	plate	Cm	Maybe a few colonies	Contamination??
MC1061	PNZ8901	tube	Cm	No colonies	no Cm Resist.

• Transformation of E.coli with different plasmids from last year

Strain	Inserted plasmid	Antibiotic	Observation	Conclusion
E.coli	I746000	Amp	No colonies	Antibiotic resistance unknown, no Amp Resist., Pb : no terminator
E.coli	I746100	Amp	No colonies	Antibiotic resistance unknown, no Amp Resist., Pb : no terminator
E.coli	I746001	Amp	Many colonies	Transformation OK
E.coli	I746101	Amp	Many colonies	Transformation OK

Antibiotic test for Bacillus resistance

We want to find the lowest concentration of antibiotic which kills Bacillus.

Dilution of Amp. [ 1:1000 means 1 part stock to 1000 part Sterile Distilled Water ]

Concentration (μg/mL)	100	75	50	25	10
100 mg/mL Stock	1:1000	3:4000	1:2000	1:4000	1:10000

Dilution of Cm. [ 1:1000 means 1 part stock to 1000 part Sterile Distilled Water ]

Concentration (μg/mL)	35	25	15	10	5
100 mg/mL Stock	1:1000	1:1400	3:7000	1:3500	1:7000

• Add Disks in antibiotic solution for 5 mins [ Protocol to sterilize tweezers : Wipe with Kimwipes, Ethanol, Flame ]
• Melt Soft Agar
• 3mL SA + 10μL Cells 1A1 from LB prepared on 23/7/08
• Pour SA+Cells over blank hard agar plates
• put disks with different concentration of Amp/Cm above SA
• Incubate at 37°C

Results [Plates were prepared before lunch, at 5pm there were visible growth]

-Amp Plates: Huge rings of no growth around 100, 75, 50, 25, and 10.

-Cm PLates: Tiny rings of no growth around 5, 10, 15 and 25. Small ring of no growth around 35 but not well defined. (No clear zones)

Salts for making Bacillus Competent

10x Medium A Base

• Yeast Extract 10g
• Casamino Acid 2g
• add Distilled water to 900mL

-Aliquot into 5 different bottles. [180mL each]

10x Bacillus Salts

• NH4)2SO4 20g
• K2PO4 anhydrous 139.66g
• KH2PO4 60g
• Tri-Sodium Citrate 10g
• MgSO4.7H2O 2g
• Add Distilled water to 1000mL

-Aliguot into 5 different bottles. [200mL each]

Medium B

• • Preparing 50mM CaCl2.2H2O
    • CaCl2.2H2O 1.470g
    • Add Distilled water to 20mL [Final conc. 500mM]
    • Take 10mL of 500mM
    • Add 90mL of distilled water [Final conc. 50mM]

• • Preparing 250nM MgCl2.6H2O
    • MgCl2.6H2O 0.508g
    • Add Distilled water to 10mL [Final conc. 250mM]
    • Take 1mL of 250mM
    • Add 99mL of Distilled water [Final conc. 250μM]
    • Take 1mL of 250μM
    • Add 99mL of Distilled water [Final conc. 250nM]

• Add 100mL of 50mM CaCl2.2H2O
• Add 100mL of 250nM MgCl2.6H2O

-Total volume 200mL

NOTE: To complete Medium B, Take 0.2mL of this solution

July 25th

Results from Yesterday

• Antibiotic test

Results for Amp

Even with a concentration of 10μg/mL, Amp kills Bacillus. Since we want to know which is the lowest concentration of Amp which kills B.S, we are going to test with some lower concentrations.

Results for Cm

Even with a concentration of 35μg/mL, ther is not a clear area around the antibiotic disk. So we have to test some higher concentration of Cm.

Wet Work

• New antibiotic tests

Dilution of Amp

Concentration (μg/mL)	10	7.5	5	2.5	1
10 μg/mL Stock (from yesterday)	1:1	3:4	1:2	1:4	1:10

Dilution of Cm

Concentration (μg/mL)	35	37.5	40	42.5	45
35 μg/mL Stock (from yesterday)	1:1	5:6 (from 45μg/mL)	1:875	17:18 (from 45μg/mL)	9:7000

- Melt Soft Agar - 3mL SA + 10μL Cells 1A1 from LB prepared on 23/7/08 - Pour SA+Cells over blank hard agar plates - Put 5 blank disks on each agar plate - Add 40μL of antibiotic on each disk - Incubate at 37°C

Results

Nothing! 1A1 cells were kept in the freedge! B.S. can not be kept in the fridge, low temperatures kill them!

July 28th

• Results of antibiotic plates from yesterday

For Cm, 35μg/mL should be enough to kill B.S.

For Amp, nothing can be concluded!

Wet Work

• Check plasmid ppL82

We had 2 samples of ppL82 in DH5α in LB solution (~4mL), one from the frozen glycerol tube and one from a colony picked on a plate. Normally plasmid and cells should not be kept in freezer. So, we want to extract this plasmid, check its size and then keep it in the fridge. We do that for the 2 different samples.

- Plasmid Miniprep (standard protocol)

- Test the concentration of DNA in each tube

1. ppL82 plate : 89.8ng/mL
2. ppL82 tube : 71ng/mL

• Preparation of different stocks of strains and plasmids

PNZ8901

- Plate a single colony (from 23/07/08) on a Cm plate

- Incubate at 37°C

- Grow the entire frozen glycerol tube in 20mL of LB without antibiotic to check if this stock is still good (we will check the plasmid on a gel tomorrow)

- Incubate at 37°C

MC1061

-Pick e single colony of MC1061 (from 22/07/08) and put it in 10mL of LB (to make glycerol stocks)

-Incubate at 37°C

1A1

- Add 100μL of LB+1A (mix of friday) and 10mL of LB

- Incubate at 37°C

• Make B.S. 1A1 competent and transform them

- Prepare Medium A

Add 81mL of SDW, 10mL of 10X Medium A base and 9mL of 10X Bacillus salts

- Make B.S. 1A1 competent

1. In 10mL of medium A and add about 10 colonies of B.S.
2. Check the OD650, you should have an OD between 0.1 and 0.2. t0 for OD = 0.1876
3. Check the OD650 every 20min and plot OD against time on semi-log paper. When the point at the culture leaves log growth, it is ok

Time (min)	0	20	40	60	70	80
OD650	0.1876	0.2074	0.3282	0.4545	0.4895	0.5040

At, t0 = 70min, log growth seems to stop.

1. Incubate at 37°C 90min after t0
2. Warn 10 Ependorf tubes with 0.45mL of Medium B
3. Add 50μL of culture in each tube of Medium B at t90
4. Incubate the diluted culture at 37°C with vigorous aeration for 90min

- Storage : we want to store 7 tubes

6 tubes in the freezer (with 60μL of glycerol)

1 tube on the bench

- Transformation

1. We make ppL82 from plate, ppL82 from tube, and a control
2. Add 0.5μg of DNA (15μL)
3. Incubate at 37°C for 30min
4. Plate on Cm plates

July 29th

• Result from the gel (29/07/2008)

• Lane8 : ppL82 (plate)
• Lane9 : ppL82 (tube)
• Lane 10 : I746001
• Lane 11 : I746101
• Lane 12 : hyperladderI

The ladder seems to be wrong. So it was really difficult to ckeck the size of our plasmid ppL82. To make, that, we assume that the size of our biobricks was ok, and we estimate the size of our plasmid. The plasmid ppL82 seems to have the right size, but we will have to check again to be able to use a ladder.

• Competence of B.S. kept on the bench

We observed B.S. with a microscope. Problem, all B.S. seems to be dead!

Possible reasons of this problem :

- Problem with the vector (it could be a wrong vector, we are not really sure)

- Quantity of DNA, we added 0.5μg. The protocol was 1μg, but normally it should be ok.

- B.S. may not be competent (we have to test motility with microscope)

- Cells could be dead : replate them

- Try to add tryptophan in the medium

Probable reason : we forgot to dilute the medium B, that's why it did not work

• Results from transformation plates (B.S.)

There are 2 colonies on a plate, but it does not look like Bacillus. Moreover, there is also a colony on our negative control plate. SO it must be contamination (resistant to CM!). We will do the transformation again.

Wet Work

• Prepare medium for transformation of B.S.

- Preparation of medium B: 10mL of medium A and 0.2mL of 50mMCaCl22(H2O) + 250mMgCl26(H2O)

- Preparation of medium A with tryptophan : 81mL of SDW, 9mL of 10X Bacillus salts, 10mL of 10X Medium A base and 0.1mL of Tryptophan (11mg/mL)

• Check plasmid PNZ8901

- Plasmid miniprep (same protocol with 60μL of elution buffer)

- Digest : with PstI and SalI from Biolabs, and Buffer 3 (add 15μL of DNA)

- Gel (17μL of sample and 3μL of dye)

Results from the gel

• Lane9 : PNZ8901
• Lane10 : HyperladderI

The sizes we expected were about 1100kb and 2100kb. The sizes should be ok!

July 30th

• Transforming Bacillus Subtilis with medium A and medium A + tryptophan

Medium A

Time (min)	0	20	40	60	73	85	95	105
OD650	0.1394	0.1370	0.1660	0.2304	0.2778	0.3138	0.3373	0.3583

For this medium, the curb stop to increase logarithmically at 100min.

Medium A + tryptophan

Time (min)	0	20	40	60	73	85	95
OD650	0.1493	0.1636	0.2080	0.2909	0.3512	0.4002	0.4165

For this medium, the curb stop to increase logarithmically at 90min.

- Incubate 90min at 37°C by shaking

- Prepare 7 tubes for medium A and 7 tubes for medium A + tryptophan : add 0.45mL of prewarmed medium B and 0.05mL of culture and incubate 90min at 37°C by shaking.

- Transform B.S. by adding DNA and incubate 30min at 37°C

We have one transformation with ppL82 (add 7.5μL of DNA), one with PNZ8901 (add 10μL of DNA) and one negative control without DNA.

- Plate 500μL on Cm plate (35μg/mL)

- For the remaining tubes, in one, add 60μL of glycerol and keep it in the freezer and keep the other one on the bench (for each medium)

• Checking our big stock of biobricks and PNZ plasmid

We did big culture of biobricks and PNZ8901 plasmid to be able to make stocks. We want to check them before making stocks.

- Plamsid miniprep (for I746001, I746101 and PNZ8901 from big flasks

- Measure DNA concentration in our samples to decide the volume of DNA we have to add in our preparation

	260/280	DNA concentration (ng/nl)
I746001	1.79	50.5
I746101	1.87	54.3
PNZ8901	1.84	87

- Digest

For I746001 and I746101	For PNZ8901 plasmid
14μL of SDW	14μL of SDW
2μL of 10X Fast Digest Buffer	2μL of Buffer 3 (Biolabs)
2μL of DNA of biobricks	2μL of DNA of PNZ8901
1μL of EcoRI	1μL of PstI
1μL of SpeI	1μL of SalI

- Gel

• Results from this gel

• Lane 9 : I746001
• Lane 10 : I746101
• Lane 11 : PNZ8901
• Lane 12 : HyperladderI

Everthing is really too big! There is a problem, either dimerization, either contamination, either a problem in our work. So we are going to run a new gel.

• New gel to check

- Miniprep plasmid from growth bottles (I746001, I746101 and PNZ8901); from plates (I746001, I746101 and PNZ8901).

- Do single digest for PNZ8901 (one with PstI, one with SalI)

- Run a gel with : PNZ8901 from friday, PNZ8901 digest with PstI, PNZ8901 digest with SalI, 3 samples from growth bottles, 3 samples from plates (to check the size of the uncut vectors)

• Result from this second gel

The ladders are really bad! But the size of our biobricks and plasmid are too big! So we can not trust these big cultures. They is a problem. With tests from the first week, we know that biobricks are right, so we are going to grow new cultures from these first cultures, to make stocks.

Concerning the vectors, they are from last year, so we are not sure of what they are. Since we ordered new well defined vectors (we should receive them on friday), we will use them in the next steps to be sure of our work.

August 1st

• Grow received vectors

Vectors	AmpR (μg/μL)	CmR (μg/μL)	KanR (μg/μL)
ECE112	100	0	0
ECE147	50	0	0
ECE149	50	0	0
ECE150	50	0	0
ECE151	50	0	0
ECE153	100	0	0
ECE162	100	0	0
ECE165	100	10	0
ECE166	100	10	0
ECE171	50	0	10
ECE172	100	10	0
ECE176	100	5	0

- Preparation of Agar plates

We have bottles of 200mL.

Type	Amp (μL)	Cm (μL)	Kan (μL)
Amp100	200	0	0
Amp50	100	0	0
Amp100 + Cm10	200	57.1	0
Amp100 + Cm5	200	28.6	0
Amp50 + Kan10	50	0	80

- Preparation of LB tubes

We prepare 10mL of LB in which we add a single colony.

Type	Amp (μL)	Cm (μL)	Kan (μL)
Amp100	10	0	0
Amp50	5	0	0
Amp100 + Cm10	10	2.9	0
Amp100 + Cm5	10	1.4	0
Amp50 + Kan10	5	0	4

August 2nd

Yesterday's Results

• All Plates grew!
  • ECE 172 seemed to have trouble growing on the plate (no single colonies, just one big lump at the start of the streak)
  • ECE 150 grew too well! (possibly hard to pick out single colony)

• Tubes:
  • ECE 176, 172, 153 seem to be quite clear!!
  • ECE 150, 151 had strange floating 'colonies' in the tube
  • All other tubes were quite cloudy

Lab Work

Fridged all plates except ECE 172 ==> Benched.

All tubes palleted and placed in freezer although ECE 176, 172, 153 doesn't seem to have any pallets

All red plates does not seem to have any extra growth despite the air conditioning had been turned off when I entered the lab this afternoon.

August 4th

Wet Work

• Check vectors

- for each vector, plasmid miniprep from frozen pellets (step 7 only once, with 60μL of Elution Buffer for ECE112, ECE147, ECE149, ECE 150 and only 30μL for the others)

- Single digest (15μL of SDW, 2μL of Buffer, 2μL of DNA, 1μL of enzyme, then incubate 10min at 37°C and 30 min for the one using HindIII with EcoRI buffer)

- Double digest (mix 7.5μL of each single digest, and then incubate 10min (or 30min for EcoRI + HindIII and EcoRI buffer) at 37°C, then heat shock 5min at 80°C

Vectors	Enzyme 1	Enzyme 2	Buffer	Entire size	Size of part1	Size of part2
ECE112	XbaI	EcoRI	Fast Digest	10156	6900	3295
ECE147	EcoRI	HindIII	EcoRI	5482	4913	600
ECE149	SpeI	PstI	Fast Digest	6192	4919	1300
ECE150	SpeI	HimdIII	Buffer 2	6166	5300	778
ECE151	SpeI	EcoRI	Fast Digest	6166	4825	1300
ECE153	SalI		Buffer 3
ECE162	SalI		Buffer 3	7600	6000	1600
ECE165	EcoRI	HindIII	EcoRI	5952	5200	793
ECE166	EcoRI	HindIII	EcoRI	7262	5100	2103
ECE171	EcoRI	PstI	Fast Digest	5444	3600	1826
ECE172	HindIII		Buffer 2	6462	3900	2540
ECE176	EcoRI	XbaI	Fast Digest	866	4250	5128

• Results

• Lane1 : HyperladderI
• Lane2 : ECE112
• Lane3 : ECE147
• Lane4 : ECE149
• Lane5 : ECE150
• Lane6 : ECE151
• Lane7 : ECE162
• Lane8 : ECE165
• Lane9 : ECE166
• Lane10 : ECE171
• Lane11 : ECE172
• Lane12 : HyperladderI

- ECE112, ECE165, ECE166, ECE171 : right bands, ok

- ECE172 : no band, not enough DNA

- ECE147, ECE150, ECE162 : only one band, maybe not enough DNA

- ECE149 : wrong size!

- ECE151 : not sure, check again

• Plates

- Streak new plates with different strains of Bacillus : 1A1, IA751 and IA771

August 5th

Checking vectors

- Plasmid miniprep for ECE166 (for stock), ECE172 and ECE153

- Digest of ECE153, ECE172 and ECE162 (with plasmid miniprep from yesterday)

- Run on a gel single digest for ECE147 (from yesterday with more DNA), ECE149 (from yesterday with more DNA), ECE150 (from yesterday with more DNA) and ECE 153, ECE162, ECE172

• Results

• Lane3 : HyperladderI
• Lane4 : ECE147
• Lane5 : ECE149
• Lane6 : ECE150
• Lane7 : ECE151
• Lane8 : ECE1162
• Lane9 : ECE172
• Lane10 : hyperladderI

Very low bands : not enough DNA!!!

Transformation of Bacillus

• Result of Nanodrop

Vector	260/280	ng/μL
ECE112	1.75	64.6
ECE166	172	138.6

- Prepare medium A with tryptophan, and medium B

- add 5mL of medium A in 3 different tubes, inoculate each tube with colonies (1A1, IA751 ans IA771)

- Check OD every 20min

- Incubate 90min

- Add 0.45mL of medium B and 0.05mL of culture in Ependorf tubes

- Incubate 90min

- Transform : add 1μg of DNA (some with ECE112, some with ECE166) (so you need to nanodrop samples before!)

- Incubate 30min

- Pipette 200μL of solution, spread it on aech plate, wait 10min, and do it again

- Incubate 24hours

- A few glycerol tubes to stock cells : add 2/7 glycerol to cell tubes

- Transformation from glycerol stock from 30/07/2008

- Spin glycerol stocks, pipette out glycerol

- Add 0.5mL of medium B, incubate for 1 hour

- Add 10μL of ECE112 (640ng)

- Incubate 2hours

- Plate 200μL, and 10min later, still 200μL

New stocks

- Do glycerol stock of I746001 and I746101 (no sterile glycerol)

- Put IA751, IA771 in 10mL LB

- Put ECE 176 in 10mL LB + antibiotic

- Reinoculate the tube of LB from yesterday with ECE166 plate

- ECE176 replated onto Amp100 + Cm5

August 6th

PCR parts of Agr operon

We received primers for Agr A,B,C, and D today - http://openwetware.org/wiki/IGEM:Cambridge/2008/Turing_Pattern_Formation/Primers

We must PCR each individual part out of the previous BioBricks (I746001 &I746101) in order to put our bacillus RBS on them.

I was concerned that the region of most homology would be the BioBrick prefix and suffix itself, so I first digested the DNA with Xbal and SpeI.

For some reason the digestion didn't work that well, so I've decided to go ahead and use the uncut BioBrick DNA as template for my reactions.

August 9th

Plans for Monday

Lab Work

• Be sure that the Spc50 plates are working, as the control for Spc50 grew and shouldn't have. We already know that things can die at Spc100 (the double crossover test from earlier this week) so it is unlikely that the stock is bad. We should test Spc50 again with regular bacillus and with E. coli at 50-100 ug/ml and hope that they die.
• Finish all the agr PCRs, but especially the RBS.
• Confirm that amyE insertion works, using the amylase/ery tests.
• Confirm that with ECE153 inserted, we get detectable fluoresence in a microscope.
• Confirm that ECE166 transformation works, using plasmid miniprepping.
  • Find the qiagen miniprep kit or adapt the protocol.
• Confirm that when ECE166 is transformed, we get detectable fluoresence in a microscope.

Book/Computer Work

• Find good promoters in vectors and order PCR primers to get them out
• Find out how to assay for/make AIP
  • Find/Contact Jim Ajioka
• Find out how to do a c-terminal GFP fusion for membrane proteins agrB and agrC
  • Can we fuse GFP and still expect them to localize to the membrane?
    • AgrB transmembrane topology (http://www.jbc.org/cgi/content/full/277/38/34736#F1). The signal peptide is not on the N terminus, and both the N and C terminus are intracellular.
    • AgrC transmembrane topology (http://openurl.ingenta.com/content?genre=article&issn=0950-382X&volume=28&issue=3&spage=655&epage=662 (see figure 2 B)), the N terminus is probably extracellular. The C terminus is probably intracellular. The article doesn't say where the signal peptide is.
  • Should we do a antibody tag instead?

Longer Term Plans

• Ligate the RBS into a BB vector so we can build a RBS/GFP construct.
• Put the RBS/GFP construct into ECE166 and ECE 153.
• Compare the 'out of the box' ECE153 with ECE153 with RBS/GFP biobrick inserts
• Compare the 'out of the box' ECE166 with ECE166 with RBS/GFP biobrick inserts

August 11th

PCR Bacillus RBS

We created 2 different Bacillus RBS sites simply by primer annealing and extension. Each product includes an 8bp RBS plus the BioBrick prefix and suffix for a total of 54bp. The products were visualized on a 3.5% Agarose gel with bioline's hyperladder V.

RBSs is the consensus RBS sequence for Bacillus subtilis, and thus we believe it to be a very strong binding site - AAAGGAGG

RBSw has a 2bp modification from RBSs, which we believe will weaken it - AGAGGTGG

The amplification of RBSs yielded only one band on the gel. The product was purified using microclean and contained 14.8ng/uL after clean-up.

RBSw, however produced 2 bands on the gel. The correct size was gel-extracted and purified. After purification the yield was 12.3 ng/uL.

August 13th

PCR Agr A, B, C, D

Doing a PCR directly from the old Biobricks has worked for all parts of the operon. No prior digestion seems to be needed.

Part	260/280	μg/mL
A	1.78	92.5
B	1.87	96.5
C	1.9	127.7
D	1.82	26.3

August 15th

PCRing out promoters

- 4 different promoters : Pxyl, Pspc, Ppac, Pupp

- Make master mix enough for 4RXNs

Digest and Ligate Agr A and C to pSB4C5

Previous PCR products of Agr A and C were digested using Fermentas Fast Digest - EcoRI and PstI according to their protocol except a longer time was given to digest.

3uL of AgrA was used and 4.4 uL of AgrC was used and incubated for 40minutes.

10uL totaling to 1ug of pSB4C5 plasmid was also digested and incubated for 10 minutes.

The digestion was visualized on a 0.8% Agarose gel and produced expected sizes:

Plasmid: approx 3bk

Agr A: approx 700bp

Agr C: approx 1300bp

August 16th

Agr A and C ligation to pSB4C5

Ligation was performed using Fermentas Rapid Ligation kit according to their protocol.

After ligation, samples were visualized on a gel, but little product of the correct size was seen.

For agrA we should have gotten a band of size 3700 (3000plasmid + 700 insert)

For agrC we should have gotten a band of size 4300 (3000plasmid + 1300insert)

The bands of appropriate sizes were extracted for transformation.

August 18th

Check promoters

On Friday, PCR out promoters, we want to check them.

- Run a gel

- Results : good size of bands!!!

Transformation of AgrA and AgrC

AgrA and C gell-extracted ligation was transformed into Top 10 cells using standard protocol. Puc9 was used as a positive control.

August 19th

Results of AgrA and C transformation

Transformation has failed. No colonies were visible for the plates of Agr A or C. The Puc9 positive control grew. We believe that the problem was in the gel-extraction between ligation and transformation. Most of our plasmid was probably lost in this step. Next time we will directly use the results from the ligation reaction to transform. Although many bands were seen on the previous gel of the ligation reaction, we will check our transformation growth by single colony PCR to confirm transformation of the plasmid with our correct insert.

Lux parts

To make the Lux Receiver, we need 4 different parts ;

• R0040, TetR repressible promoter

• SO168, luxR + double terminator

• R0062, promoter activated by luxR

• JO4630 (GFP + double terminator)

All these parts have been transformed into E.coli. We want to test them. R004, R0062 and JO4630 have already been tested, it should be fine. We received from the MIT R0040, R0062 and S068 already transformed into E.coli, so we want to check these stocks (which are certainly fine) and use them. For JO4630, we want to double check our transformation.

- Plate on antibiotic plates and do LB stocks of single colony from the MIT stock (R0040, R0062 and S0168).

- Put on Kan plates 4 different colonies from J04630 (transformation Amp plate) and also incubate these colonies into LB

August 20th

Check Lux components

- Single colony PCR for :

• R0040 (MIT stuff)

• R0062 (MIT stuff)

• 4 different colonies of S0168 (from a transformation plate from 12/08)

• 4 different colonies of J04630 (from a transformation plate from 12/08)

- Protocol : add 1μL of cells (diluted in water), 10μL of Master Mis, 7μL of SDW, 1μL of VF primer and 1μL of VR primer

- Gel PCR products

Gel 1

• Lane2 : Hyperladder1
• Lane3 : JO4630, colony 1
• Lane4 : JO4630, colony 2
• Lane5 : JO4630, colony 3
• Lane6 : JO4630, colony 4
• Lane7 : HyperladderI

Gel 2

• Lane2 : HyperladderI
• Lane3 : ECE190 double digest
• Lane4 : S0168, colony 1
• Lane5 : S0168, colony 2
• Lane6 : S0168, colony 3
• Lane7 : S0168, colony 4
• Lane8 : HyperladderI
• Lane9 : R0040
• Lane10 : R0062
• Lane11 : Ladder 100bp

- Results

• R0040 and R0062 : one big band of about 300b (expected size 293), OK!

• S0168 : one band of about 400b for the 4 different colonies (expected size 1234!), bad! This plate does not contain S0168

• J04630 (colonies 2 and 4) : one band of about 1100b (expected size 1173), OK!

• J04630 (colony 1) : one good band plus another band...

• J04630 (colony 3) : one band of about 600b, bad!

Ligation

- Materials :

• AgrA
• AgrB
• AgrC
• AgrD
• Pupp
• Pspac
• Ppac
• Pxyl
• RBS S
• RBS W
• psB4C5

- Double digest of PCR products

- Run vector, AgrA and AgrD on a gel

- DNA clean and concentrator for AgrA, B,C and D, promoters

- Microclean for both RBS

- Nanodrop

	260/280	ng/μL
AgrA	1.66	16.4
AgrB	1.91	23.5
AgrC	1.99	35.9
AgrD	2.13	4.9
Pxyl	1.54	5.6
Ppac	1.49	4.6
Pspc	1.62	9.6
Pupp	1.88	8.5
RBS S	2.44	29
RBS W	1.44	10.7

- Extract plasmid annd Agr from gel and clean

- Ligation

August 21st

Transformation of ligation products

- Spin chemically competent TOP10, add 100μL of CaCl2 solution

- Add 5μL of DNA (ligation products), and 1μL of PUC9

- Continue the protocol of transformation

J04630

- Plate good colonies from yesterday on Kan25 and put in LB (for plasmid stock for tomorrow)

Plasmid stocks

- Plasmid miniprep R0040 and R0062

August 23rd

Check promoters (after PCR from bacillus vectors)

- Gel

• Lane1 : Hyperladder 5
• Lane2 : Pxyl
• Lane3 : Pspac
• lane4 : Ppac
• Lane5 : Pupp

Size is ok.

August 27th

New transformation of ligation products

• Products : agrD, Pupp, Pspac, RBS W, RBS S (from ligation with biolabs kit)

- Competent top 10 from the freezer

- Add 5μL of ligation products, and 1.2μL of PUC9

August 28th

Results from transformation of ligation products

- nothing on plate, even on control plate

• Reasons ?

- cells non competent anymore (try with fresh competent cells)

- not enough DNA (try with 10μL of DNA)

- very short ligation products

Transformation of ligation products (new)

• Products to ligate : Pupp, Pspac, agrD, RBS S and RBS W

- new fresh competent TOP 10

- 5μL of DNA (1.5μL of PUC9)

- 2h30 in the incubator

August 29th

Results of transformation with our ligation products

• Everything grew! Better efficiency with electrop. than with chemical protocol

Single colony PCR to check our transformation

Transformed products	number of picked colonies from chemical transformation	number of picked colonies from electrop. transformation (neat)	number of picked colonies from electrop. transformation (1:10)
Pupp	2	2	0
Pspac	2	2	0
RBS S	3	0	2
RBS W	3	0	2
agrD	3	2	0

- Single colony PCR : 13μL of SDW+cells, 5μL of Master Mix, 1μL of VF and 1μL of VR (and plate each single colony)

- Load a gel (1.3% agarose) : 5μL of PCR products + 1μL of dye (only 1μL of 100b ladder)

In the death plasmid, the VF-VR size is about 280b.

Transformed products	size of the product (with cutting sites)	expected size after PCR (about)
Pupp	255	480
Pspac	125	350
RBS S	56	280
RBS W	56	280
agrD	200	430

• Result : nothing, even no ladder, problem with the gel!

- run again on a e-gel

• Lane1 :ladder 100bp
• Lane2 : Pupp colony 1
• Lane3 : Pupp colony 3
• Lane4 : Pspac colony 1
• Lane5 : Pspac colony 3
• Lane6 : RBS S colony 1
• Lane7 : RBS S colony 4
• Lane8 : RBS W colony 1
• Lane9 : RBS W colony 4
• Lane10 : agrD colony 1
• Lane11 : agrD colony 4
• Lane12 : HyperladderI

• Result : nothing, just the primers! Problem with our PCR

Plate biobricks from MIT

• E0840
• B0014
• I712007
• C0012
• B0015
• C0061
• R0063

September 2nd

Check transformation in E.coli of our ligated products

- Single colony PCR with VF and VR : add 10μL of SDW, 5μL of MM, 1μL of VF, 1μL of VR and 3μL of cells

inserted products	number of colonies to check (chemical transf.)	number of colonies to check (electrop. transf.)
RBS S	3	2
RBS W	3	2
Pspac	2	2
Pupp	2	2
agrD	3	2

- Gel (3% of agarose)

• Result

gel	lane	inserted product (name of the colony)	observation	conclusion
top	3	RBS S (1)	one band (260bp)	size of the insert, RBS too small to see on a gel
top	4	RBS S (2)	one band (260bp)	size of the insert, RBS too small to see on a gel
top	5	RBS S (4)	nothing	?
top	6	RBS S (5)	nothing	?
top	7	RBS W (1)	2 band (260bp + 350bp)	size of the insert, RBS too small to see on a gel + something else?
top	8	RBS W (2)	one band (350bp)	too big
top	9	RBS W (4)	one band (260bp)	size of the insert, RBS too small to see on a gel
top	10	RBS W (5)	one band (260bp)	size of the insert, RBS too small to see on a gel
top	11	Pspac (1)	2 band (260bp + 350bp)	maybe problem with products loaded on gel (exactly the same bands than for RBS W)
top	12	Pspac (2)	one band (450bp)	size of Pupp?
top	13	Pspac (3)	one band (450bp)	size of Pupp?
bottom	3	Pspac (4)	nothing	?
bottom	4	Pupp (1)	one band (400bp)	size of Pspac?
bottom	5	Pupp (2)	one band (400bp)	size of Pspac?
bottom	6	Pupp (3)	one band (slightly lower than 400bp)	size of Pspac?
bottom	7	Pupp (4)	one band (400bp)	size of Pspac?
bottom	8	agrD (1)	2 bands (380 and 450bp)	big band is ok
bottom	9	agrD (2)	2 bands (380 and 450bp)	big band is ok
bottom	10	agrD (3)	no bands	?
bottom	11	agrD (4)	strong band (450bp)	ok
bottom	12	P2	one band (350bp)	ok

September 3rd

Checking the insert of promoters

We ligated promoters into death vector, and transforned into Top 10. Our transformation worked, but it was hard to say with the result of yesterday if we had the good insert. Moreover, it is possible that we inverted Pupp and Pspac. So we are going to make a new single colony PCR with the primers of promoters (we used the same single colonies than yesterday).

We also want to PCR again all promoters (Pspac, Pxyl, Pupp and Ppac) from plasmids to have more.

Picture of the gel with the different promoters

promoter	colony	primers	name
Pspac	1	Pspac primers	A
Pspac	1	Pupp primers	B
Pspac	2	Pspac primers	C
Pspac	2	Pupp primers	D
Pupp	1	Pupp primers	E
Pupp	1	Pspac primers	F
Pupp	3	Pupp primers	G
Pupp	3	Pspac primers	H

- Single colony PCR : add 10μL of SDW, 5μL of MM, 1μL + 1μL of primers, 3μL of cells

- Gel

• lane 3 : hyperladderI
• lane 4 : Pspac (colony1) with Pspac primers
• lane 5 : Pspac (colony1) with Pupp primers
• lane 6 : Pspac (colony2) with Pspac primers
• lane 7 : Pspac (colony2) with Pupp primers
• lane 8 : Pupp (colony1) with Pupp primers
• lane 9 : Pupp (colony1) with Pspac primers
• lane 10 : Pupp (colony3) with Pupp primers
• lane 11 : Pupp (colony3) with Pspac primers
• lane 12 : HyperladderI

- Results

promoter	colony	primers	observation	Conclusion
Pspac	1	Pspac primers	nothing	it is in fact Pupp
Pspac	1	Pupp primers	band (about 280bp)	Pupp OK
Pspac	2	Pspac primers	nothing	it is in fact Pupp
Pspac	2	Pupp primers	band (about 280bp)	Pupp OK
Pupp	1	Pupp primers	nothing	it is in fact Pspac
Pupp	1	Pspac primers	band (about 180bp)	Pspac OK
Pupp	3	Pupp primers	nothing	it is in fact Pspac
Pupp	3	Pspac primers	band (about 180bp)	Pspac OK

Make some stock of our new biobricks

• Pupp (colony1) into the death vector, transformed into TOP10
• Pspac (colony1) into the death vector, transformed into TOP10
• agrD (colony5) into the death vector, transformed into TOP10

- Grow this single colony into 10mL of LB (Cm35)

Transformation of new "biobricks"

- transform into E.coli :

• agrA
• agrB
• agrC
• Pxyl

September 4th

Results of new "biobricks"transformation

- Nothing on our plate, except for the control plate.

New attempt of new "biobricks" transformation

- transform into E.coli these ligated products:

• agrA
• agrB
• agrC
• Pxyl

September 7th

New PCR

- New stocks of Master Mix

- PCR agrA, agrB, Pxyl, Pspac, Ppac and Pupp

September 8th

Check product of PCR from yesterday

• Lane3 : Hyperladder I
• Lane4 : agrA
• Lane5 : agrB
• Lane6 : Pxyl
• Lane7 : Pspac
• Lane8 : Ppac
• Lane9 : Pupp
• Lane10 Hyperladder IV

• Results : everything is ok

Check transformation from 03/09

After several days of incubation, we have a few colonies on our transformed plates (agrA, B and C).

• Single colony PCR

- Add 13μL of cells diluted into SDW, 5μL of MM, 1μL of VF and 1μL of VR

• Gel

September 10th

PCR of GFP+RBS and Promoter+RBS

- PCR

- Run a 1.2% agarose gel with 1μL of sample

• Lane 3 : HyperladderI
• Lane 4 : GFP + RBS 1A
• Lane 5 : GFP + RBS1B
• Lane 6 : GFP + RBS 2A
• Lane 7 : GFP + RBS 2B
• Lane 8 : Pupp + RBS 1
• Lane 9 : Pupp + RBS 2
• Lane 10: HyperladderIV

• Result

RBS screening

• 5 colonies from chemical transformation + 6 colonies from electroporation transformation for RBS S
• 5 colonies from chemical transformation + 6 colonies from electroporation transformation for RBS W

- PCR : 11μL of SDW + 5μL of MM + 1+1μL of primers (RBS detect + VR) + 2μL of cells (program iGEM34)

- Gel1

• Lane2 : RBSS1
• Lane3 : RBS S2
• Lane4 : RBS S3
• Lane5 : RBS S4
• Lane6 : RBS S5
• Lane7 : RBS S6
• Lane8 : HyperladderIV
• Lane9 : RBS S7
• Lane10 : RBS S8
• Lane11 : RBS S9
• Lane12 : RBS S10
• Lane13 : RBS S6 with VF and VR primers
• Lane14 : RBS S11

- Gel2

• Lane2 : RBSW1
• Lane3 : RBS W2
• Lane4 : RBS W3
• Lane5 : RBS W4
• Lane6 : RBS W5
• Lane7 : RBS W6
• Lane8 : HyperladderIV
• Lane9 : RBS W7
• Lane10 : RBS W8
• Lane11 : RBS W9
• Lane12 : RBS W10
• Lane13 : RBS W6 with VF and VR primers
• Lane14 : RBS W11

• Result

- Nothing, except for the PCR with VF and VR primers, either our primers are not right, either we have no insert. But the gel is not good enough to see very well, we will try to run a new gel.

September 11th

RBS screening

- We run our PCR products from yesterday on a new 1.8% agarose gel

- Gel1

• Lane2 : RBSS1
• Lane3 : RBS S2
• Lane4 : RBS S3
• Lane5 : RBS S4
• Lane6 : RBS S5
• Lane7 : RBS S6
• Lane8 : HyperladderIV
• Lane9 : RBS S7
• Lane10 : RBS S8
• Lane11 : RBS S9
• Lane12 : RBS S10
• Lane13 : RBS S6 with VF and VR primers
• Lane14 : RBS S11

- Gel2

• Lane2 : RBSW1
• Lane3 : RBS W2
• Lane4 : RBS W3
• Lane5 : RBS W4
• Lane6 : RBS W5
• Lane7 : RBS W6
• Lane8 : HyperladderIV
• Lane9 : RBS W7
• Lane10 : RBS W8
• Lane11 : RBS W9
• Lane12 : RBS W10
• Lane13 : RBS W6 with VF and VR primers
• Lane14 : RBS W11

• Result

- Nothing, except for the PCR with VF and VR primers, either our primers are not right, either we have no insert.

RBS screning but single digest

- Grow some colonies of Top10 transformed with RBS into 10mL of LB with antibiotic

We want to check our transformation with single digest. If we have self ligation in our transformation, we will loose the XbaI cutting site.

Backbone for ligation

- New pellets in 600μL of SDW

- Plasmid miniprep

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