(Difference between revisions)

May 2007 Su M Tu W Th F Sa 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31

June 2007 Su M Tu W Th F Sa       1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30

July 2007 Su M Tu W Th F Sa 1 2 3 4 5 6 17 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31

August 2007 Su M Tu W Th F Sa       1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31

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September 2007
Sa	Su	M	Tu	W	Th	F
						1
2	3	4	5	6	7	8
9	10	11	12	13	14	15
16	17	18	19	20	21	22
23	24	25	26	27	28	29
30

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October 2007
Su	M	Tu	W	Th	F	Sa
	1	2	3	4	5	6
7	8	9	10	11	12	13
14	15	16	17	18	19	20
21	22	23	24	25	26	27
28	29	30	31

Contents 1 May 2007 1.1 May 14 1.2 May 15 1.3 May 16 1.4 May 30 2 June 2007 2.1 June 4 2.2 June 20 2.3 June 21 2.4 June 22 2.5 June 26 2.6 June 28 2.7 June 29 3 July 2007 3.1 July 5 3.2 July 6 3.3 July 11 3.4 July 12 3.5 July 17 3.6 July 18 3.7 July 19 3.8 July 20 3.9 July 21 3.10 July 22 3.11 July 24 3.12 July 25 3.13 July 27 3.14 July 31 4 August 2007 4.1 August 1 4.2 August 2 4.3 August 3 4.4 August 4 4.5 August 8 4.6 August 9 4.7 August 10 4.8 August 11 4.9 August 12 4.10 August 13

May 2007

May 14

1. Made the same as May 15, but did not use Type 1 (milliQ) H2O, so the solutions were discarded.

May 15

1. Made 1M CaCl2, and glycerol/CaCl2 solutions
   1. 1M CaCl2 Solution: 100mL was made
      1. CaCl2 Dihydrate = 146.986 g/mol
      2. 100mL of 1M CaCl2 requires 14.699g CaCl2 Dihydrate
   2. Glycerol/CaCl2 Solution: 30mL was made
      1. 5.00mL of 60% glycerol solution
      2. 3.00mL of 1M CaCl2
      3. 22.00mL of Type 1 (milliQ) H2O

May 16

1. Transformation of Cells:
   1. Transformed 1 vial of cells with Jun vector, and another vial with Fos vector (1uL of vector each); Sterile
   2. Used 400uL of LB (sterile) for each vial
   3. Incubated for 1 hour in shaker at 37C, then incubate overnight at 37C

May 30

1. Test PCR Machine

June 2007

June 4

1. Made Kanamycin plates
   1. 200mL of Agar
   2. 200mL of 2x LB
   3. 2mL of Kanamycin

June 20

Today: extract DNA from plates, transform, plate on amp plates.

I - Extraction of DNA from Plate (5M and 9G)

1. Puncture foil with pippette tip
2. Add 15uL of Type 1 water
3. Remove all liquid (with DNA in solution)

II - Transformation of extracted DNA

1. Chill cells on ice for 10 min
2. Heat shock cells at 42°C for 30 sec
3. Add DNA for cells
4. Add 500uL of 42°C SOC medium to each vial
5. Incubate on ice for 1 min
6. Place in 37°C shaker incubator for 1h 30min
7. Plate onto 4 separate amp plates
   1. 25uL R0062
   2. 25uL C0060
   3. 400uL R0062
   4. 300uL C0060
8. Place into 37°C incubator at 3:55PM

Next: seed, miniprep and screen the DNA.

June 21

Checked colonies in the 37°C incubator. Well formed colonies were on each plate, so plates (all 4) were transferred to the small fridge (middle shelf on the right).

June 22

Today: miniprep DNA from July 20th transformation

I - Miniprep of Colonies from R25, R400, C25, C300 (indication the biobrick and the amount in uL that was plated)

1. Transfer 1.5mL of cells suspended in LB into a microcentrifuge tube.
2. Centrifuge for 2 min at max speed (14000 rpm)
3. Remove supernatant with pipette and discard.
4. Add 250uL of buffer P1 and 250uL of buffer P2
5. Close and invert tubes 6 times
6. Add 350 uL of buffer N3 (within 5 min of adding buffer P2)
7. Close and invert 6 times
8. Centrifuge for 10 min at 13000 rpm
9. Carefully pipette off the supernatant into a Qiaprep spin column. NB: for sample R400 a large amount of supernatant was discarded. No reason, stupid move by Tim.
10. Centrifuge spin column for 60 sec
11. Discard flow-through
12. Add 750uL of buffer PE
13. Centifuge for 60 sec (max speed)
14. Add 50uL of sterile water (of EB buffer could be used)
15. Let stand for 1 min then centrifuge for 1 min
16. Transfer flow-through to a new micro-centrifuge tube (one with a cap) with a pipette.
17. Stored in -20°C freezer (green box labelled "Team 2")
18. Remaining cells in LB stored in small fridge

Next: Screen the DNA.

June 26

Today: Screen of R0062 and C0060 from July 20th transformation

I - For R0062 add:
1.5uL of buffer 2
1.5uL of 10X BSA
3uL mini-prepped DNA
1uL of XmnI
1uL of XbaI
7uL of sterile water
Total volume = 15uL

II - For C0060
Same as above but no XbaI and 8uL water

Let both sit in 37°C water bath for 1h 40 min (From 3:30PM to 5:10PM)

III - Gel electrophoresis

1. Add 12uL of water to each R25, R400, C25, C300
2. Add 3uL of respective digested DNA
3. Add 1.66uL of loading dye
4. Gel contains:
   1. Ladder (5uL)
   2. R25 uncut
   3. R25 cut
   4. R400 uncut
   5. R400 cut
   6. C25 uncut
   7. C25 cut
   8. C300 uncut
   9. C300 cut
5. The remaining uncut mini-prepped DNA was stored in the pink "Team 1" box.

Next: Interpret the screen and proceed to midi-prep or reseed.

June 28

Today: Reseeding of R0062

I - Reseeding of R0062

1. Prepare 6 culture tubes with 2.5mL of sterile water and 2.5 mL of LB medium
2. Add 5uL of 1000X Amp to 5 of the tubes (no antibiotic added to the control)
3. 3 seedings from R25, 2 seedings from R400
4. At 4:00PM the seedings were placed in the 37°C shaker incubator (in styrofoam pieces wedged inbetween clamps.
5. Plates returned to fridge

Next: Miniprep and screen the re-seed.

June 29

Today: Digest and screen the re-seed from June 28th

I - Digest using same procedure as June 26th

1. DNA was removed from the water bath at 4:35PM

II - Gel electrophoresis

1. Gel contains
   1. Ladder (5uL)
   2. 400A
   3. 400B
   4. 25A
   5. 25B
   6. 25C
2. Gel turned on at 5:02PM. 100V set for 40 min with variable current.
3. Picture saved as "June 29 R400A R400B R25ABC"

Next: Interpret the gel and plan appropriate course of action.

July 2007

July 5

Today: Digest and screen R and C from July 4th midiprep

I - Digest Volumes
1.5uL 10X buffer
0.5uL suspended DNA (less than usual b/c midiprep is more conc.)
1uL EcoRI
12uL water
Total volume - 15uL

At 11:52AM tubes were placed in a 26.2°C water bath that was on its way up to 37°C.

II - Spectrophotometric DNA quantification

1. Dilute DNA by a factor of 1000 (i.e. 1uL of DNA in 999uL of water). Mix by tapping.
2. Reference (at 260 nm) a quartz cuvette that has been rinsed and filled with water.
3. Rinse the cuvette with DNA solution.
4. fill the cuvette and take an absorbance reading at 260 nm.

Results: A(R@260nm) = 0.022;    a(C@260nm) = 0.75

Calculation: [DNA] = 50 x (dilution factor) x (Absorbance at 260nm)
Therefore [R] = 1100 ng/uL; [C] = 4250 ng/uL

Next: Gel quantification and ligation

July 6

Today: Gel quatification of "I" and "R"

I - Gel quantification
Wells contain:
7uL water
1uL loading dye
2uL DNA (either the vector "R" or the insert "I")
Order of wells:

1. Ladder (5uL)
2. Insert
3. Vector

The gel was run for 45 minutes.

July 11

Today: Transformation of ligation product (R+C)

I - Transformation

1. Same procedure as June 20 except:
   1. All of ligation rpoduct was put into cell capsules
   2. Heat Shock was 2 min b/c they are Top 10 cells
2. Plates were stored overnight in 37°C incubator.

Next: Seed and miniprep

July 12

Checked plates and there was no growth, so we have to set up a the ligation again.

July 17

Today: Plating of ligation and extra plasmid (to check the efficiency of transformation)

I - Plating

1. Went to take cultures out at 2:30PM but the shaker had been turned off. So the shaker was turned back on and left to run for an additional 15 min.
2. Tubes were removed from SI at 2:45PM and cells were plated on Amp plates.
   1. Control (R400) 25uL
   2. Control (R400) 300uL
   3. Ligation 25uL
   4. Ligation 250uL
3. Stored in 37°C incubator over night

Next: Seed and miniprep

July 18

Today: Seeding of ligation plate (R0062 from July 17)

1. Prepare 10 culture tubes
   1. 2.5mL sterile water
   2. 2.5mL LB medium
   3. 5uL Amp
2. Seeds were only taken from the 25uL ligation plate. The 250uL plate was too full to distinguish colonies.
3. Prepared two controls as well
   1. water and LB only
   2. Water + LB + Amp
4. All tubes were placed in 37°C incubator at 4:05PM

Next: Miniprep and screen

July 19

Today: Miniprep of Ligated DNA (R0062 from July 17)

NOTE: After starting, Alex pointed out that due to the lack of ribosome binding site in our construct, what 
 we are miniprepping is useless.  We will keep going though to make sure that our ligations are working.

I - Miniprep

1. Follow usual mini-prep protocol as per June 22.
2. We had to wait before proceeding to the screening digest because there were no small tips autoclaved. Miniprepped DNA was left on benchtop for the 20 min that tips were autoclaving.

II - Screening Digest 5uL of mini-prepped DNA
1.5uL of buffer 2
1.5ul of 10X BSA
1uL of EcoRI
1uL SpeI
5uL sterile water

Gel contents:

1. Ladder
2. Culture A
3. Culture B
4. Culture C
5. Culture D
6. Culture E
7. Culture F
8. Culture G
   

Cultures H through J, we ran out of SpeI and so were not digested or run on the gel.

Results: We know how to ligate. YAY! Now we restart.

Next: Incorporate a RBS

July 20

Today: Extraction of B0034. I - Extraction of B0034.

1. Pierce the foil top with a pipette.
2. Extract the DNA with 15ul of sterile water.
3. Transfer to a microcentrifuge labeled [B0034; 20/07/07; 15ul] and stored in pink "Team 1" box.

II - Tansformation

1. Allow Top 10 cells to thaw on ice for 10 min. DNA was also placed in the ice bucket.
2. Add 2uL of DNA to Top 10 cells.
3. Incubate on ice for 30 min then heat shock at 42°C for 30 sec.
4. Add 500uL of LB, 250uL SOC and ice for 1 min.
5. Place in 37°C shaker incubator for 1 hour (started at 3:12PM)
6. Plates put into 37°C incubator at 4:50PM
7. Remaining cells placed in Team 1 freezer box; labelled "B0034 cells"

Next: Seeding

July 21

Today: Seeding of B0034. Hanmo and Allan performed the seeding. Standard protocol. Two tubes per concentration plate (i.e. 25 and 250)
Placed into 37°C SI at 1:20PM

Next: Mini-prep

July 22

Today: Dilution of seeding. Justin and Hanmo diluted the seeded cells by a factor of two. They divided the original cultures in liquid media into 2.5ml batched and then diluted them all to 5ml with 1X LB (with amp)
Went into the 37°C SI at 10:45AM.

Next: Mini-prep.

July 24

Today: Quantification of Midi-prepped B0034 NB: On July 5th we diluted by a factor of 1:1000, but this was because we expected the DNA to be very concentrated (and it was). Depending on the volume of DNA solution (and therefore the supposed concentration)the dilution might only have to be 1:100. But it turns out that due to the barely visible pellet we will proceed with a 1:1000 dilution.

Results: A(B1@260nm)= 0.086 A(B2@260nm)=0.087
[DNA]= 50 x (duilution factor) x (absrobance @ 260nm)
= 150 ng/ul for B1
= 200 ng/ul for B2
In the future we might want to use the 1:100 dilution Next: Sceen the midi-prep and ligate

July 25

Today: Sceening of Midi-prepped B0034 Only 0.5ul of Ladder was added because we were using syber-green. the load in other wells was: 1ul DNA, 1.67ul loading dye, 14ul water (total: 16.67ul)
The gel was loaded as:

1. Ladder
2. uncut B0034
3. cut with XbaI
4. cut with XmnI + BseRI
5. cut with BseRI
6. cut with XmnI
7. cut with XmnI + XbaI

The gel was started at 3:36PM and run for 50 min. The gel picture was saved as "July 25 B0034 5 enz".

Next: Proceed with the next ligation.

July 27

Today: Turn off water incubator (37C) and placed in the freezer to stop the digest at 12:56PM. THe tube is in the pink Team 1 freezer box. (It's the only tube with tape on it.)

Next:Transform the ligation.

July 31

Today: Transformation of B0034+C0060 ligation.

August 2007

August 1

Today: Seeding of the B0034+C0060 ligation I-Seeding
5 colonies from the 70ul plate and 5 from the 215ul plate (+2 controls)
Each tube contains:

1. 2.5ml of 2X LB
2. 2.5ml of distilled water
3. 5ul Amp

Culture tubes placed in the 37C shaker incubator at 4:58PM. Plates were put back in the fridge (top shelf right hand side). NB: the 215ul had a cracked lid; the crack has been sealed with tape.

Next: Mini-prep and screen.

August 2

Today: Mini-prep of seeding from Aug 1 Digest carried out with EcoRI and SpeI in EcoRI buffer. The mini=prepped DNA was diluted by an extra 5ul of water during the digest so more DNA and less water was used when loading the gel (to ensure a decent resolution).

From the results of the gel we diluted 250ul of cells in suspension from tube 6f (215ul batch, #5) in 25 ml of LB and 25ul Amp, for Midi-prep.
The flask went into the SI (37C) at 6:15PM.

Next: Midi-prep.

August 3

Today: Midi-prep and screen Note that the SI was turned off when Steph checked this morning, but there was growth, so we will proceed nonetheless. The Midi-prepped DNA was digested in two batches to ensure that everything worked properly.
Digest 1: BsrGI and SpeI with expected bands at 2455bp and 425bp.
Digest 2: BseRI and SpeI with expected bands at 2096bp and 784bp.
Digests were carried out in the 37C water bath starting at 12:23PM

We obtained unpromising results from the second digest. We suspect that the SpeI may be faulty, so we will try to cut it with BseRI and PstI. In buffer 2 PstI has only 75% efficiency so we will leave it overnight. Digest was started at 4:09PM.

After speaking with Jay we are going to try to regrow some cells left over from the original seeding (the 1 out of 10 that worked). There wa about 300ul left. We diluted 250ul in 25ml of LB. Cells went into the incubator at 5:26PM.

Next: Check on new digest and on cells in incubator.

August 4

Today: Check on new digest and on cells in incubator Digesting DNA was taken out of the 37Cincubator at 11:08AM. BAD NEWS from the gel. Only one band b/w 3kb and 2kb and it wasn't a short band that ran off the gel, the smallest section of ladder was halfway down the gel. Picture is saved as "Aug 4 (BseRI + PstI)"

Removed

Also the bleach bottle is cracked and I didn't have time to switch it. Next: Check on new digest and on cells in incubator.

August 8

Today: Plating of B0034+C0060 (the second ligation attempt)

1. Put Amp plates in the 37C incibator to warm them up at 2:25PM
2. Plates went into the incubator at 3:23PM. Right Hand wall middle shelf

Next: Seed, mini-prep and screen

August 9

Today: Seeding and prepping reagents
I - [Preparation of 500ml of 2X LB]x2
II - Preparation of 200ml of agar

1. 6g in 200 ml of water

III - 3 x 500ml of sterile water (type 2)
All went into the autoclave on setting 1 at 1:15PM

IV - Seeding of ligation: 250ul plate only

1. Ten cultures (2.5ml LB, 2.5ml water, 5ul amp)
2. Put into 37C shaker at 2:30PM

Note: the 25 ul plate had very abnormal growth so it was not seeded.

Next:mini-prep

August 10

Today: Mini-prep of 10 overnight cultures of ligated DNA
I - Cut with BseRI + SpeI and BseRI (if it needs to be redone, use PstI instead of SpeI). All 20 digests went into the 37C at 2:10PM

II - Gel electrophoresis

1. We used 3ul of mini-prepped DNA, 12ul water, 1.66ul of loading dye
2. In the way of results: 6 showed well in both, 3 showed well for the double digest but not the single cut, 8 showed well for the single but not the double
3. All the same, we prepared midi-prep dilutions of all three (25ml LB, 250ul cells, 25ul amp) and we will screen the regrown cells before continuing.
4. The dilutions went into the incubator at 6:40PM

Next:Midi-prep

August 11

Today: Midi-prep of dilutions of cultures 3, 6 and 8.
After midiprep the DNA was quantified spectrophotometrically.
3: 108 ng/ul
6: 30 ng/ul
8: 990 ng/ul
Due to the small amount of DNA from #6 we prepared another dilution in case it needs to be re-midi-prepped

I - Gel electrophoresis

1. 8 cut with BsrGI one band at 3000bp
2. 3 cut with BsrGI one band at 1300bp
3. 8 cut with EcoRI + SpeI faint band at 3kbp others at 2kbp and 700bp
4. 3 cut with EcoRI + SpeI one band 2kbp

File saved as "August 11 C+B midi screen digest"

Next: Large scale digest for ligation.

August 12

Today: Large scale digest of #8
I - Conditions

1. Midi-prepped DNA (1.5ul)
2. Xba (1ul)
3. PstI (1ul)
4. BSA (5ul)
5. Buffer 3 (5ul)

Place in 37C water bath at 4:32PM. Will be left overnight because of the large amount of DNA to cut and Xba has only 75% efficiency in buffer 3. There is no star activity, so it shouldn't be a problem.

Next: Gel extraction.

August 13

Today: Gel extraction
I - Running the Gel

1. Result: 8 was bad, 6 was cut properly

II - Gel extraction

1. Weigh excised gel pieces
2. Add 3ul of QG buffer for every 1mg of gel
3. Put in 50C water bath for 10 min (vortex a bit every 2-3 min)
4. Solution should be yellow
5. Add 1 ul of isopropanol for every 1 mg gel
6. Load into a spin column, spin for 1 min, discard flow-through
7. Add 0.5ml QG wash, spin 1 min, discard flow-through.
8. Add 0.75ml PE wash, spin 1 min, discard flow-through.
9. Spin for 1 more min,

Next: