Janet B. Matsen:Closed Lab Questions

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Revision as of 18:37, 17 November 2012 by Janet B. Matsen (talk | contribs) (Why do we disinfect with 70% EtOH? Isn't 100% more potent?)
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I define "closed" as "not bothering me incessantly." One can always learn more but we have to stop at a point otherwise we would never move forward with our research!

If you have any input/corrections/references please drop me a line.

I'm sorry for the low quality pictures. I have a very old phone, and those pics were decently high tech in its day!

Lab Basics:

Why do we disinfect with 70% EtOH? Isn't 100% more potent?

  • A: I have read that you don't want to denature the proteins that allow substances like water and ethanol into the cell. If you do, the EtOH can't get into the cytosol. By using 70%, you denature some of the transporter proteins but leave enough to allow more to enter the cell. Also, 70% is cheaper than 100%!
  • I still use 100% for everything.
  • 70% is still concentrated enough to be very flamable!

Wow.. these 2 batches of TB media are very different in color... did I mess up?

  • A:Maybe, but not necessarily. Different batches of yeast and tryptone can have very different compositions. This is one of the purposes of defined media.
    these two TB batches were made with the same recipe but different bottles of one of the yeast-derived reagents. I forget whether it was yeast extract or tryptone.

How does colony size vary with colony density?

  • As the number of colonies per unit area increases, nutrient diffusion becomes limiting. New molecular biologists are sometimes worried that variation in colony sizes on a plate indicates contamination, but it can be caused by tight packing.
    colony size varies with colony density

Colony PCR

The gel is dark where I loaded it -- what does this mean?

  • A: It probably means you overloaded the PCR with too many cells.


Should I pay extra to further purify large bp oligos?

  • Usually no. The sequencing companies just want your money. Do sequence your creations to confirm that you made what you expected and remember you can always screen a new colony if the first one looks bad.

My primers are yellow!

normal oligos have a yellow color that is visible with large synthesis scales
  • Don't worry -- this is normal! Moreover, it is extra visible when you order a lot of a primer (large synthesis scale) because there is more of it to see.


How do I decide whether to gel purify after digestions (before ligations)

  • The BioBrick/Ginkgo manual doesn't instruct you to. Besty always does.
  • Um... I don't do much restriction enzyme cloning at this point... Gibson rules! 7/2012


Is it ok to re-use electroporation cuvettes after they arc and turn black?

normal and "burnt" electroporation cuvettes
  • Electroporation cuvettes are aluminum. The "black stuff" that appears after arcing is aluminum oxide. Though this usually appears low in the cuvette away from the pocket that holds the cells, it correlates to oxidation & non-uniformity in the metal that does contact the cells. This non-uniformity makes arcing more likely when you electroporate a future sample. You will notice that re-used cuvettes are more likely to arc, and that ones with visible oxidation are even more likely to arc.
  • To reduce arcing, reduce the resistance (ohms) setting. Though you do need some current to flow through the cells for electroporation, reduced current leads to less arcing. I set my minimum to 200 ohms & maximum to 500 ohms 2012/07/31.
  • To avoid arcing completely, use chemically competent cells instead. Though it is more work to prepare the special growth media, you will save time/money/DNA reletive to electroporation.
  • added 7/31/2012

How does the density of electrocompetent cells affect transformation efficiency?

electroporation: number of colonies versus competent cell density
  • It matters a LOT! The undiluted samples (dilution factor = 0) were harvested ad mid-exponential phase and resuspended in 1/500ths of the culture volume. Details of the experiment are here.

What is the ballpark efficiency of electroporation?

  • In the experiment described just above, I calculated that for 0.1 ng of plasmid, the maximum efficiency was 5*10-6. I haven't looked up literature values to see how this compares to published statistics. I find this amazing, because it makes Gibson cloning seem even more amazing. Even if you get only a few colonies, there were likely many orders of magnitudes more viable plasmids in the transformation product than that!

Gene Expression & Plasmids

Is a two plasmid system where the two plasmids have different antibiotic resistance genes but the same origin of replication stable?

  • A collaborator implied use of a different antibiotic is sufficient. However, I suppose this could lead to unpredictability in the number of each plasmid. For example, suppose the copy origin they share yields 100 plasmid copies. It could be that 10 copies of the antibiotic A & 90 copies of the antibiotic B plasmids is equally favorable as 90 copies of the antibiotic A plasmid & 10 of the antibiotic B plasmid. Presuming the two plasmids encode different proteins, the relative ratios of the expression levels would be drastically important.
  • If allowed to compete in a flask, the burden of each plasmid will affect the distribution of the plasmids.
  • added 7/16/2012 after talking to BioRad customer service


The parts in the registry have sequence that don't include the prefix & suffix... do they come with it?

  • Yes, they come with a prefix/suffix. You can find which BioBrick method a part is compatable with under the part's "Assembly Compatibility" listing. Most are RFC 10 (http://partsregistry.org/partsdb/scars.cgi), the standard format.


What is the different between phosphate buffer & PBS (phosphate buffered sailine)?

  • Phosphate buffered saline has MUCH lower concentration of phospate buffer. It is mostly salt.

How stable is HEPES?

  • Not stable. Make it the day you use it.

How do I decide the spacing between a promoter and ribosome binding site? A ribosome binding site & the start codon?

  • Recall that a promoter starts making a transcript at the +1 site, no matter the sequence, and that the promoter is only for transcription and the RBS is only for translation. The -10 and -35 locations in the promoter that are significant for binding are numbered relative to this +1 site. You can have any arbitrary sequence before the RBS, and people often include operators for genetic regulation. Or, you can put the ribosome binding site next. This is more standard. The ribosome binding site needs to be correctly spaced from the start codon, and you have to look at the individual RBS to determine it. For some, a BioBrick scar is the perfect distance between the end of the RBS and the start codon; this is useful for making an expression vector like UW's 2010 iGEM vectors.