Team 2 Notes

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Contents

Summary

Our goal of this assay was to test the activity of Transposase, but due to complications with our composite parts we didn’t get that far.

We amended the Transposase Activity Assay so that we could test its activity with pre-existing and simpler parts, but we had an issue with the plates we used for the co-transformation and didn’t have time to do it again.

We ended up running several tests to see what concentration of atc was necessary to induce self-lysis and were successful with a 100x dilution. However, it is important to note that self-lysis only occurred in LB media with that concentration of atc, and not in NEB buffer, which is probably more similar to the conditions inside the vacuole.

We also performed a Degradation Assay, which is a test to determine the rate of degradation of DNA in the lysate. It involved adding plasmid DNA to the lysate, removing samples at certain time intervals, and then using transformation and colony counting to measure how the amount of plasmid DNA changed over time. Our results showed that this degradation rate must be fairly slow, since all of our samples from 0-30 minutes produced the same number of colonies.

4/28/10 Wetlab Notes: Disappointing co-transformation results, Degradation Assay with 100x atc dilution

Cotransformation update

We planned on performing the entire transposition activity assay today, but there was a problem with our co-transformation. The kan and spec weren't evenly distributed on the plate, so we couldn't perform an accurate selection.

Degradation Assay

Dilutions:

  • 1mL of 1X NEB buffer: add 100uL 10X NEB2 to 900uL water
  • 1mL of 100x anhydrotetracycline (atc) dilution: add 10uL 2mg/mL atc to 1mL solvent (1X NEB or LB)

Protocol:

  • Prepare 1X NEB buffer (see above)
  • Pellet 1uL of cell suspension in 2 tubes.
  • Discard supernatant
  • Add 1mL of 1X NEB to each tube and resuspend
  • Transfer 1mL of cell suspension (already in LB) to 2 other tubes
  • Add 10uL of stock atc to one of the NEB tubes and one of the LB tubes and start the clock
  • Incubate all tubes in shaker for one hour.
  • Transfer 250uL of the lysate from the LB+atc and the NEB+atc vials into tubes.
  • Add 3 uL of pUC-ampR plasmid to each tube.
  • Remove 10 uL of cells at different intervals (0, 5min, 10min, 15, 20, 30)and immediately add 180 uL ADB buffer (Zymo clean-up protocol)
  • After all samples have been taken, spin down the cell junk and transfer the supernatant, which contains the DNA) to a zymo column.
  • Perform a zymo clean-up. Elute with 10uL of water.
  • Transform the 6 samples into TG1 cells according to the Standard protocol (divided by 4):
    • 7.5uL KCM
    • 12.5uL water
    • 50uL cells (TG1 cells)
    • 10uL of eluted plasmid
    • plate on ampicillin

Results: Cells in the LB + 100x diluted atc lysed after 1hr as expected. Cells resuspended in NEB2 with 100x diluted atc appear to have failed to lyse over in 1hr. Hence we only drew samples at various time points from the lysed cells in LB.

The plates made for various sampling time points all appeared to have a similar number of colonies. It looks like the plasmid did not degrade much over our sampling time range.

4/26/10 Wetlab Notes: Self-Lysis test with 100x atc dilution

Protocol:

  • Took 1mL of mid-log self-lysis plasmid and added 10uL of stock atc (100x dilution)at 2:36pm. Put in incubator at 37deg in shaker. Also used a control without added atc.

Results:

  • The mixture looks clearer at 55min (they lysed!).

Analysis:

  • Centrifuged the cells from the "ATC" and "No ATC" vials and ran the supernatant in a gel to look for genomic DNA. If self-lysis had occured, there would be more genomic DNA in the supernatant of the "ATC" vial. This was the case as shown below:

Gel:

Image: team2apr26.jpg

  1. Ladder
  2. self-lysis transformed cells LB - No atc (control)
  3. self-lysis transformed cells LB - with atc

4/21/10 Wetlab Notes: Miniprep of pRL27, Co-transformation, Self Lysis Test at 100x atc

Protocols:

  • Self lysis test 3 (100x atc dilution): Added 10uL of 1x atc to 1mL of cell at 2:58pm. After letting the cells incubate for about an hour, the cell solution looked somewhat clearer, but we didn't have a control vial to compare them to due to complications, so we should perform the test again.
  • Miniprepped the Tr-Kan-Tr--Transposase--R6K (pRL27) plasmid using the standard protocol.
  • Co-transformed Rep+/Pir+ cells (strain jtk094) with the pRL27 plasmid and the 2741-pBjk2294 plasmid (Ptet-Self-Lysis--Spec--ColE2) by following this protocol:

Standard protocol divided by 20:

  • 1.5uL KCM
  • 2.5uL water
  • 10uL cells (jtk094 strain)
  • 1 uL self-lysis plasmid
  • 1 uL pRL27 plasmid
  • plate on kanamycin


Transformed jtk094 cells with just 1 uL of the self-lysis device so we can do another test to determine how much anhydrotetracycline is needed to induce self-lysis.

4/19/10 Wetlab Notes: Degradation Assay with 1000x atc dilution

Same protocol as below, but change the dilution of atc to the following 1000x dilution:

  • 10x dilution: Add 1uL of stock atc (2mg/mL) to 9uL of ddH20.
  • Add 2.4uL of that 10x dilution to the 240uL of 1X NEB2. (10uL removed for t=0 sample)

Results:

Visual inspection suggested that self-lysis failed. We did not plate samples since we would likely see no degradation over time.

4/14/10 Wetlab Notes: Degradation Assay with 10^5X atc dilution

The goal of today's assay was to see if plasmid in the buffer would degrade when exposed to lysate (from induced self-lysis). We had to re-suspend self-lysis plasmid transformed cells in NEB2 buffer, add AmpR plasmid, then expose the sample to atc. We took samples of this mixture at various time points and added ADB to each 10uL sample to halt degradation. These samples were then zymo cleaned, transformed into TG1, and plated on Amp.

Dilutions:

  • To get 250uL 1X NEB buffer, add 25uL 10X NEB2 to 225uL water
  • To get 2ug/mL anhydrotetracycline (atc), add 1uL 2mg/mL atc to 1mL water

The protocol we followed:

  • Pellet 250uL of cell suspension
  • Discard supernatant
  • Add 250 mL 1X NEB buffer to cells and resuspend
  • Add 3 uL of pUC-ampR-sbb23 plasmid
  • Add 2.5uL of (2ug/mL) atc and start the clock
    • 2.5uL of 2ug/mL atc in 250mL will give another 100-fold dilution to give 20ng/mL atc
  • Remove 10 uL of cells at different intervals (0, 5min, 10min, 20, 30, 45)and immediately add 180 uL ADB buffer (Zymo clean-up protocol)
  • After all samples have been taken, spin down the cell junk and transfer the supernatant to a zymo column(it contains the DNA).
  • Perform a zymo clean-up. Elute with 10uL of water.

Transform the 6 samples into TG1 cells according to the following protocol:


UPDATE: A similar number of colonies were observed on all of the six plates mentioned above. The cells most likely failed to lyse, hence the plasmid was not degraded over time.

4/12/10 Wetlab Notes: Self-lysis device transformation

We were given self-lysis plasmid. We had to transform it into Lefty cells so we could carry on with our later assays.

Standard transformation protocol divided by 20:

  • 1.5uL KCM
  • 2.5uL water
  • 10uL cells (Lefty cells... pretty close to MC1061 pir+)
  • 0.5uL self-lysis plasmid
  • plate on kanamycin

4/7/10 Presentation Feedback

  • Use MC1061(pir). This strain doesn't degrade arabinose due to mutations in some of its metabolic pathways.
  • Variable: Time at which we add arabinose. Could try adding arabinose at the same time as anhydratetracycline (aTc) or an hour before aTc.
  • Could take real cells and generate buffer by grinding them up.
  • Don't use PBS since it doesn't have Mg.
  • Self-lysis won't take ~3.5 hours. It will probably take a few minutes. NOTE: incubating too long might cause DNA in the buffer to be degraded by nucleases.
  • IMPORTANT: determine rate of DNA degradation in the lysate. Could transform cells with lysate samples taken at different times and count colonies.
  • Zymo might be fine instead of miniprepping lysate.
  • MC1061, DH10B, TG1, all choices for (pir -)
  • DPN1 fingerprinting could be used for mapping. Will create ~10-20 cuts in the average plasmid. Do wild-type digest side by side with integrand, compare bands.
  • How much plasmid will we add to the lysate?

4/5/10 To do

ASAP

  • Transform cells w/ self-lysis device (kanR)
    • How much plasmid to use? Usually, 1uL of the standard concentration of pre-digested vector is used for ligation, and then the entire ligation mix is added to the 70uL of cells. Then if the self-lysis device is at a similar concentration, 1uL should be enough to add to our cells. (**Edit: GSI's say to use 0.5uL**)
  • Figure out how much plasmid to add to the lysate
    • The pUC-ampR plasmid we add will ultimately be the plasmid we want to do the second transformation with in our actual assay. So we need to add enough plasmid so that 600uL of the lysate will produce enough plasmid to perform a transformation after the 600uL of lysate has been miniprepped.
  • Degradation Assay (a day long transformation procedure)
  • Figure out what buffer to use (research online)
    • Paper on cytoplasm mimicry "Mimicking the Escherichia coli cytoplasmic environment activates long-lived and efficient cell-free protein synthesis"

Once the composite parts are complete

  • Transform genR-selflysis-transposase composite part into pir+ strain
  • Arabinose assay: test when we should add arabinose and how much we should add to optimize transposase expression (time: an hour before, when we add anhydrotetracycline)-> this will require a day long transformation procedure.
  • The actual assay


Next time: Count colonies

Materials: -ampR plasmid -cells w/ self-lysis device? Or will we have to transform them ourselves?

3/31/10 TO DO - Completed

  1. Find Data about Pbad. How much arabinose is required for expression?
  2. Find Data about Ptet. How much tetrocycline is required for expression?
  3. Decide on a buffer. Similar to vacuole? Similar to Cytoplasm?
    • PBS?
    • transposition buffer: 100 mM Tris pH 7.5, 150 mM MgCl2, 100 mM KCl, 10 mM DTT
  4. Finish powerpoint.
  5. Test cell lysis device?


3/29/10: Protocol planning

Assumed starting materials:

  • composite part (CP) w/ genR flanked by terminal repeats, self lysis device, and prepro-transposase under conditional origin of replication (R6K)
  • arabinose (how much?)
  • "vacuole" buffer (VB)
  • pUC-ampR plasmid (how much?)
  • tetracycline (how much?)

Steps:

  1. Transform a Pir strain with CP, following standard protocol. Pir necessary for plasmid to replicate because it contains a R6K origin of replication. Plate cells on gen, and select colonies. Colonies should be grown in a media containing ___uL arabinose, as the transposase is under a pBad promoter.
  2. Create lysis buffer by adding ___mL of VB, ___uL of pUC-ampR plasmid, ____uL of tetracycline (self-lysis device is under pTet promotor), and ___uL of arabinose (transposase is under pBad promoter)
    • Research promoters to find optimal amount of inducer necessary and research transposases to find out how much plasmid is necessary.
  3. Add cells to lysis buffer and let incubate for ___ min. At what temperature? Agitate? (Need to find conditions that optimize cell lysis and the transposase reaction)
    • (Note: from 2008 Berkely iGEM: The cultures were then incubated at 37 degrees again for 3.5 hours, and the absorbance at 600nm was measured with a Tecan Xfluor4 Safire2 in a Corning Inc. Costar 3603 plate. The data plotted on a log scale is shown below.)
    • Find self lysis protocol and transposase info to determine these conditions
  4. Lysate buffer (at this point it contains VB, cell junk, desired plasmid, CP, chromosomal DNA, transposase, tetracyclin, arabinose) needs to be mini-prepped. Use standard mini-prep protocol.
  5. Now we have our desired plasmid and original CP in water. Use this solution to transform cells using the basic protocol. Make sure you DON'T use a strain of Pir cells! We want the CP to fade out.
  6. Plate transformed cells on amp/gen plates.
  7. Select colonies.
  8. Mini-prep and map the plasmids so determine if and where the cassette was inserted.
    • Need to know sequence of DNA that the zinc-fingered transposase binds to and where it inserts transposons

Controls:

  1. Testing the R6K origin of replication (To ensure that genR expression in tranformed non-Pir cells is due to transposase activity and NOT successful transformation and replication of the CP)
    • Transform the non-Pir strain with only genR. Use media with arabinose, as in the previous protocol (don't want to introduce another variable).
    • Plate on gen.
    • There should be NO colonies. If there are colonies, the conditional origin of rep is not functioning properly.


Useful links

General

  • Zinc finger paper zf+ comes from Lwt, zf- comes from Rwt, binding sites are shown in Figure 4


Buffer Research

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