User:GeorgeXu

Me

Important Stuff that was done (in reverse chronological order)

7/12/07

My First Clonewell

1. Get iBase+Clonewell gel. Remove gel (leave the comb!) and place into iBase.
2. Pre-run (Mode 0) for two minutes while the comb is still in the gel
3. Take out comb, load each well in top row with 20 µL samples, the middle well with ~10 µL ladder, and the bottom row wells with 20 µL nuclease-free water
4. Run the gel with (Mode 5: Run Clonewell), watch the bands and stop the gel when your desired band hits the dashed line above the bottom row of wells
5. With the gel stopped, fill the bottom row wells with nuclease-free water.
6. Run the gel again. When you see the band starting to enter the well, take out the sample in the well and put into clean labeled centrifuge tube. Then refill the well with 20 µL nuclease-free water and continue running the gel.
7. Repeat this procedure until you get enough sample. If the band runs past the well, you can use Mode 6: Reverse Clonewell to make the band move back into the well

So basically, I did this for the stuff that was digested (see immediately below) and I ended up with a lot of water, so I am going to vacufuge the results tomorrow.

Digests of P0140, P0340, R0011, and R0051

• XbaI and PstI digests
  • P0340
  • P0140
• SpeI and PstI digest
  • R0011
  • R0051
• Mixture
  • 42.5 µL DNA
  • 1 µL enzyme 1
  • 1 µL enzyme 2
  • 0.5 µL BSA
  • 5 µL Buffer 2

Protocol

Perry's E-gel

I ran an E-gel for Perry's colony PCR.

1. Take out E-gel (leave comb in) and put in one of the bases. NOTE: One of the red bases has a faulty contact which requires covering by aluminum foil to work correctly.
2. Pre-run the gel for 2 minutes. (Press and hold either 15min or 30min on the red bases or Mode 0 on the gray bases)
3. When the pre-run is finished, press any button to stop the beeping.
4. Load 10 µL of nuclease-free water and 10 µL of ladder into the first well. Then 10 µL nuclease-free water + 10 µL sample into every other well. If there are empty wells, put in 20 µL of nuclease-free water into the empty wells. NOTE: Nick mentioned this and it is a pretty good idea. Put the ladder in the middle well or if there are empty wells, put ladder in the end empty wells. This way, it is easier to compare, especially if the bands bend.
5. Run the E-gel for 30-min (Press 30min on the red bases, or run Mode 1 - changing the time if necessary)
6. When the run is done, press any button to stop the beeping. Use the UV machine in the small room to visualize the gel.

It confirmed that P0140 and P0340 were correct. However, I don't think his GFPmeks turned out very well.

Results

7/10/07

Colony PCRs

7/9/07

Growing Liquid Cultures of Tomorrow's Plate Reader Samples

Protocol

Stephanie and I grew liquid cultures of the following parts:

• I13263
• I13273
• I5311
• I13522
• B0015
• T9002 (+)
• T9002 (-)

Also, Stephanie left the T9002 (+) sample incubating with 1000 nM OHHL to serve as a comparison to J37015.

Colony PCR of Ligated Parts

Protocol

Stephanie, Perry, and I colony PCR'ed the parts that they ligated the week before.

See Stephanie's notebook for results.

6/29/07 FACS DAY (I13263)!

FACS-related

• 8:30 AM - Stephanie and I took the OD of the overnight cultures.
  • Results - 1.785, 1.793, 1.780
• 9:15 AM - Stephanie and I performed first dilution (growing the 4 hr induction cells). Unfortunately, we didn't dilute correctly...
  • 1:40 dilution: 50 µL liquid culture + 148 µL LB broth + 2 µL ampicillin
  • 1:80 dilution: 25 µL liquid culture + 173 µL LB broth + 2 µL ampicillin
  • 1:100 dilution: 20 µL liquid culture + 178 µL LB broth + 2 µL ampicillin
• 10:30 AM - Oopsie, we realized we didn't dilute 1:40, etc.; instead, we had actually diluted 1:4, etc. We measured the OD's of these incorrectly diluted samples and rediluted the samples. Note that in order to measure the OD, we diluted samples 1:100, measured the OD, and multiplied the measured OD by 100 to give us our measurement for the actual sample.
  • 1:4 sample OD = 3, redilute 1:100 giving us a sample of about OD=0.03
  • 1:8 sample OD = 1.8, redilute 1:30 giving us a sample of about OD=0.05
  • 1:10 sample OD = 1.3, redilute 1:100 giving us a sample of about OD=0.01
  • Also, we diluted 1:40, 1:80, and 1:100 2 mL samples from the overnight stock to grow as our 2 hr induction cells
• 11:00 AM - I diluted the stock solution 1:10,000 and 1:1,000,000 in 100 µL 200-proof ethanol to give a 97 µM and 97 nM solution.
• 11:30 AM - Stephanie and I diluted 1:40, 1:80, and 1:100 2 mL samples from the overnight stock to grow as our 1 hr induction cells
• 12 noon - Stephanie and I took the OD of the 4 hr induction cells
  • 4 hr induction cells, 1:40: 0.2
  • 4 hr induction cells, 1:80: 0.2
  • 4 hr induction cells, 1:80: 0.1

Since each of these were about 2 mL samples, we didn't have enough sample to have 2 mL cuvette samples. Instead, we used 1mL of cells with 10.5 uL HSL. 2 samples were taken from the 1:40 first dilution (8:30am) - used for the 10 and 100nM HSL runs - and 1 from the 1:80, used with 1nM HSL.

• 1:30 pm - Took OD of the 1hr and 2hr samples + Induce the 2 hr samples

See Stephanie's entry for the rest

Transformation and Liquid Culture of Assorted BioBricks

Transformation Protocol

Liquid Cultures Protocol

Perry and I transformed and prepared liquid cultures of the following parts:

• S03623
• S03608
• J37034
• J23039
• R0011
• R0140
• T9002
• B0015
• R0051

6/28/07

Resuspending the HSL stock

I took the 25 mg of N-Butyryl-DL-homoserine lactone and resuspended in 1.5 mL of 200-proof ethanol. I then put the tube in the Quorum sensing box in the freezer at Stephanie's bench.

This was done in preparation for inducing the FACS samples on 6/29/07.

Digestion of F1610: Part Two

Stephanie ran another F1610 digest with

1. 2.5 µL Buffer 2
2. 0.5 µL BSA
3. 0.5 µL XbaI
4. 0.5 µL PstI
5. 21 µL F1610

Basically it is the same as the previous digestion except we now have 21 µL vs. 15 or 6 µL of DNA. This was done because the Clonewell of the F1610 that Perry and I ran had an almost invisibly faint band corresponding to the F1610. Where did all the DNA go?

Grow Liquid Cultures of I13263

Protocol

Stephanie and I grew I13263 in preparation for the FACS on 6/29/07.

Sequences

Perry checked the sequences that we got back from Genewiz
Results

• F1610 (something is very wrong...)
  • The actual part is 798 bp, we got back a sequence ~300 bp
  • The sequence contains the terminator and some random junk after it
• I13263 (w00t)
  • The first and last ~650 bp are perfect.
• I13272
  • !!!!add something here

!!!!upload the sequences?

Clonewell of F1610

!!!!Protocol Results
F1610 is about 800 bp. There is an almost invisibly faint band at 800 bp. What happened to all the DNA? We need to run it again

Transformation of Additional Parts

Protocol

Perry and I transformed the following parts
!!!!descriptions

• R0040
  • Promoter
• R0051
  • Promoter
• R0011
  • Promoter
• E0240
  • RBS+GFP+Terminator
• B0015
  • Terminator
• T9002
  • Complete luxR+GFP system

Digestion of F1610 (XbaI, PstI)

Mix 1:

1. 2.5 µL Buffer 2
2. 0.5 µL BSA
3. 0.5 µL XbaI
4. 0.5 µL PstI
5. 15 µL F1610 (168 ng/µL)
6. 6 µL Nuclease-free water

Mix 2:

1. 2.5 µL Buffer 2
2. 0.5 µL BSA
3. 0.5 µL XbaI
4. 0.5 µL PstI
5. 15 µL Nuclease-free water
6. 6 µL F1610 (168 ng/µL)

Perry and I ran two digests of F1610 with different amounts of DNA. The plan is to gel extract and ligate with the promoters that Perry has transformed.

Protocol

6/27/07

Sequencing the Midiprepped QS parts

Protocol

Just checking if the Midiprepped sequences are correct. Details of the order:

Tube Label	DNA Name	DNA Length	Primer
AV01	F1610	3987	VF2
AV02	F1610	3987	VR
AV03	I13263	4123	VF2
AV04	I13263	4123	VR
AV05	I13272	4162	VF2
AV06	I13272	4162	VR

Transformation of I13263

Protocol

Stephanie and I transformed 1 µL of the I3263 that we midiprepped into 30 µL of BL21. We then plated them onto agar+ampicillin plates. We eventually want to carry out a test experiment by inducing the I3263 with different concentrations of HSL to test and possibly start characterizing the part.

Hispeed! Midiprepping of the QS parts

Protocol

Perry and I Midiprepped the QS parts in order to get DNA that we could sequence and transform.

6/26/07

Growth of QS parts in Liquid Cultures

Protocol

We grew the QS parts in a liquid LB+ampicillin broth.

Nanodrop

Protocol

Results
10mer: 389.8 ng/µL 15mer: 278.6 ng/µL 20mer: 168.7 ng/µL

Nucleotide Removal of the 10mer, 15mer, and 20mer library

Again, this was done in preparation for the ligation reaction that we will do with the massive amounts of DNA.

Protocol

PCR Extension of the 10mer, 15mer, and 20mer library

Basically, this was done in preparation for the ligation reaction that we will do with the massive amounts of DNA.

Protocol

6/25/07

Transforming and Plating the QS parts

Protocol

Perry transformed the parts F1610, I13263, and I13272 into Top10 cells and plated them on LB+ampicillin plates. We want to sequence the parts, test them, and eventually use them in our BL21 cells.

6/22/07

Counting colonies: Overnight vs. Short Ligation

We counted the number of colonies in the Overnight vs. Short Ligation plates.
Results

Nanodrop of Ligated Colonies

Protocol

Results
OmpA1+random library+extension: 50.8 ng/µL OmpA2+random library+extension: 50.7 ng/µL The curves looked smooth. %

6/21/07

Protein Gel of OmpA+His/OmpA+Strep Cells

Protocol

We ran this protein gel to see if the OmpA1+His, OmpA1+Strep, OmpA2+His, and OmpA2+Strep was actually expressed. Also, we wanted to see if there was any difference between the immediate induction, delayed induction, and no induction.

Results

Bacterial Innoculation and Induction

Inocculation and Induction Protocol

Measuring Optical Density Protocol

We took the OmpA1+His, OmpA1+Strep, OmpA2+His, and OmpA2+strep that were grown in liquid culture overnight starting on 6/20/07 and split them into three sets. The first set was not induced, the second set was immediately induced with 10 µL IPTG¸ and the final set was induced once it hit log phase, which we determined by measuring the optical density.

Optical Density Results

6/20/07

Gel Extraction

Protocol

We extracted the DNA from the gel we ran on 6/19/07

Gel after Excision

6/19/07

Running the OmpA1/OmpA2 gel

Protocol

We poured a gel and ran the DNA from the dephosphorylated samples.

Results

Dephosphorylating the OmpA1/OmpA2

Protocol

Six times this amount of mix was made as a Master Mix in order to dephosphorylate the OmpA1 and OmpA2. There were five groups total:

1. Group 1: OmpA1, Nhe1, Pst1
2. Group 2: OmpA1, Nhe1, Pst1
3. Group 3: OmpA2, Nhe1, Pst1
4. Group 4: OmpA2, Nhe1, Pst1
5. Group 5: Perry...