Lidstrom:GeneMorph II

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These notes are assembled from Janet Matsen's 1st creation of an enzyme library, leveraging Yakov Kipnis' extensive experience.


  1. Mutazyme rxn (GeneMorph II kit)
  2. PCR clean-up
  3. Digest with NcoI/XhoI (+DpnI if desired)
  4. Gel extraction
  5. Insert self-ligation test
  6. Ligation into backbone


Mutazyme II rxn

The enzyme in this kit is more sensitive to operating conditions than most polymerases. It can be sensitive to:

  • template concentration (specifically inhibitors present)
  • adding enzyme into the 10X buffer directly (best to add it at the end & not include it in a master mix).
  • Yakov: "In my opinion Mutazyme is a little bit more finicky than other polymerases and requires more attention to make it work."
  • First steps:
    • Make sure your amplification primers work using robust polymerase
      • Phusion works well because it has the same extension temperature. OneTaq Quick Load 2X also works at 72°C despite the suggested extension temperature of 68°C.
    • nester PCR?
      • You may consider nested PCR using gel purified product as template and standard robust polymerase (it is probably not even necessary to use high-fidelity polymerase)
  • Optional controls:
    • omit template. This is useful when you are using a lot of template.
      • If you get a faint band that you think is your product, it may actually be some minor species in your template prep instead of what you think it is.
    • Include a well with OneTaq or Phusion as a + control.
  • Once you have it working:
    • Run a range of template concentrations. Though the manual suggests correlations between template concentrations and mutation rates, it is best to use a range. Errors in NanoDrop DNA concentration calculations, pipetting, and PCR effects can effect the final mutation rate.
      • Yakov: "I always prepare several libraries (with different expected level of mutagenesis) and sequence 10-20 clones from each. This allows very reliable estimation of mutation load in each library and especially frequency of wild type sequences. For actual activity screening I obviously take the library/libraries that satisfy whatever criteria I had in mind."
      • Yakov: "I have been in the situations when GeneMorph kits simply didn't work (bad production batch probably). Mutagenic reactions with lowest possible concentration of template and largest number of cycles (conditions that according to instructions should flood the gene with 20 mutations/kb) barely produced 0.5 mutations/kb. Therefore I prefer scanning range of conditions (easiest being range of template concentrations) at least first time so not wasting time on sequential optimization of conditions. I do several reactions in parallel (taking advantage of using master mixes as much as possible) and preparing as many libraries as I feel necessary or can process. Characterizing the libraries with small scale sequencing. At the next step I pool together libraries I feel similar enough to each other, discarding bad libraries and end up with very few (even single) libraries I intend to screen."
  • Re-amplify with Phusion or other high fidelity polymerase
    • This helps you have enough DNA to do the digests, gel purifications, and ligations. There is a small chance you will reduce the diversity of your transformants, but it is probably unlikely.
    • You absolutely don't need the whole Mutazyme reaction. What I would do is take 2 uL of unpurified Mutazyme rxn, dilute it 100 fold in water, and use 2-4 uL as a template for reamplification. You can keep the rest of Mutazyme reaction as an alternative route to compare results afterwards.
  • Debugging
    • Consider using a minimal amount of template to see if something inhibitory is present. Or, desalt as you would a gibson with a millipore filter.
  • Yakov: "I have seen that Mg2+ titrations may be very helpful (as in case of other polymerases) to get Mutazyme to work."

PCR cleanup

  • Don't worry about yield loss here (Yakov advice)



Gel Extraction

Ligation & controls

  • testing: re-ligate your digest to make sure the enzymes worked.
  • use T4 NEB ligase. Nothing fancy.

Ligation into backbone


  • Yakov: "Transformation efficiency you want is limited by number of clones you can screen. There is no point of getting 10^9 transformants if you can test only 10^3. Any method reliably and reproducibly giving you 1000 colonies (if you can screen 96) should be OK. Electroporation is obviously a "gold standard", but considering hands-on time with cuvette, electroporator etc I would vote for something heat-shock based. Inoue's method is fine if you feel comfortable making your own cells. Advantage is you are not limited by bacterial strains available from companies. Buying cells is also fine. Since you don't need a lot of transformants you can easily split aliquot of commercial cells into several transformations which make it even more economically feasible. Keep in mind even commercial cells can be bad (I have encountered non-transformable cells from Invitrogen; I have obtained them via UW BioChem store therefore don't know whom to blame)."

What about the GeneMorph® II EZClone Domain Mutagenesis Kit?

Yakov: "I am familiar with the kit, and funny enough you already have all the components of this kit in your possession. This is variation of QuickChange mutagenesis protocol with synthetic primers replaced by longer PCR product (containing mutations). Sometimes I use this approach to do quick cloning of mutated gene if I have plasmid with wt gene, in my hands this approach never produced lots of transformants, therefore I never use it for diverse libraries. Also to me it seems to be less amenable for troubleshooting (QuickChange step, which is all-around amplification using megaprimers, is not exponential in nature and typically produces less DNA and therefore not always obvious whether your reaction worked or not). So far, to the best of my knowledge, with all the different funny methods developed for special purposes the most straightforward and robust method to construct large library is standard restriction/ligation mediated cloning. But it doesn't preclude my attempts to find/develop simpler protocol."

What do I do with mutants that are "hits"?

If you identify a handful or even a hundred mutants that have higher activity, you need to figure out which mutations benefit the enzyme. If you had >1 mutation per gene on average, a lot of the mutations are carried along and are simply neutral or even harmful. Your options include:

  • DNA shuffling
    • Nobody in the lab has experience with this as of 12/2014, but there is a vast literature about it. You basically use sonication or an enzyme that chops up template genes into bits before reassembling.
  • Use primers that encode interesting mutation(s)
    • You can assemble whole genes by assembling oligos that have homology on the ends. Therefore, one can include some oligos that have various mutations in interesting position(s) to make a combinatorial library of known mutations.
    • Yakov's comments:
      • "The method David Baker was probably talking about is a gene assembly from overlapping oligos. I use this method to quickly assemble genes or make combinatorial libraries. You split your genes into overlapping fragments (overlaps ~15-25 bases) order oligos, do assembly PCR (technically not a polymerase chain reaction), and reamplify fragment with outer primers (standard PCR). The full length product can be ligated into plasmid and transformed. If oligomix contains oligos with degenerate codons in non-overlapping regions, assembled product will be a library of genes with one or few or multiple positions diversified. Have a look at chapter 13 in the attached book, Directed Evolution Library Creation, ISBN 978-1-4939-1052-6"
      • Chapter cited = "Assembly of Designed Oligonucleotides: A Useful Tool in Synthetic Biology for Creating High-Quality Combinatorial DNA Libraries"

Can I use Gibson Assembly to insert mutated GeneMorph colonies?

  • Janet is trying 1/2015, but am not sure whether I am limited by assembly and transformation efficiencies.

How can I increase my transformation efficiency?

  • Initially (1/2015) Janet Matsen was trying to use Gibson assembly to insert mutated genes onto my backbone. There were two problems: I wasn't getting enough colonies from transformations, and ~25% of colonies I did get lacked insert. I was looking for ways to increase efficiency, so I thought of:
    • Not transforming directly into BL21(DE3). Is this strain in general less competent than strains like Top10, as my intuition suggests? Can I transform into Top10, get orders of more colonies and miniprep them before putting them into BL21(DE3?)
    • Reverting back to restriction cloning after giving Gibson a shot.

Yakov's thoughts:

  • BL21(DE3) tends to be 1-2 orders of magnitude less efficient for transformation than cloning strains
    • "In my opinion transformation of strains engineered for DNA work is better practice. Generally these strains have 1-2 orders of magnitude higher transformation efficiencies and better plasmid prep quality than BL21(DE3) engineered for protein production. I consider transformation of freshly assembled library straight into expression strain as a quick-and-dirty shortcut, which saves a lot time, but requires some confidence. If your current plasmid is pET_something then transforming it into strain that doesn't have T7Pol is probably the best way of dealing with toxic protein product leakage (if I understand your concern about toxicity). You can safely and efficiently transform into Top10, extract plasmids and retransform nice supercoiled DNA into BL21(DE3). I do this strain exchange in cases of large naive libraries when losing diversity due to toxicity may be an issue."
  • Increase the DNA concentration, consider desalting libraries even for chemically competent cells
    • "Typically, there is a correlation between amount of DNA and number of transformants, but it is not monotonic. It looks like there is usually an optimum or plateau when further increase of DNA amount doesn't result in increased transformation efficiency. If you want to try scaling up Gibson and transforming more DNA, you should probably think about desalting and concentrating your reaction before actual transformation even if you do chem comp cells and not electroporation where desalting of enzymatic reactions is typically required."
  • Scraping colonies off of plates works fine, and may help preserve diversity relative to liquid culturing.
    • "Scraping colonies from plates is absolutely OK. This way you can potentially preserve your diversity by letting slow growers to accumulate some biomass as a separate colony without being outcompeted in the liquid culture. I have done it a lot. To take the full advantage of the technique you probably should spread your transformation reaction on several Petri dishes or use larger plates. To collect cells just add ~1 ml of resuspension buffer from your favorite miniprep kit per standard Petri dish, use scraper, bent glass pipet or simply wash colonies by pipetting up and down. Try to do it relatively quickly, but without collecting excessive amount of agar. Tilt the dish on one side aspirate remaining liquid (~500 ul, rest will be soaked in by agar, depending on how old/dry the agar layer is). Harvest cells. Do your miniprep."

My strategies to try next:

  • Compare the efficiency of electroplating into Top10 to what I have been getting in BL21(DE3).
    • If it is orders of magnitude more efficient, miniprep from Top10 and transform that supercoiled DNA.
    • I can scrape colonies off of plates and miniprep them directly if I wish.
    • Photograph the plates and make sure to under-sample the diversity miniprepped.
  • Give electroporation another shot. Requires making a fresh prep of cells because my old electrocomp BL21(DE3)s weren't amazing in recent (within a few months ago) tests.
  • Try desalting Gibsons to see if they transform better
  • Buy fresh ligase and try restriction cloning as a last resort. (I'm concerned about it being more work and requiring Phusion re-amplification of gel purified GeneMorphed products!)