Software to download

I recently bought myself a new computer for the office, which meant that I had to download and install all of the key software I used for work. I decided to write down what these were so future me (or future employees) could have it as a reference. All of the following links / commands are for use with a mac.

Everyone should install these:

Google Chrome

Google Drive & Sync or Google Drive File Stream

Microsoft Office


R (requisite for R Studio)

R Studio 
packages worth installing: tidyverse, shiny, ggrepel

XQuartz (requisite for Inkscape on older macs)




Sublime Text 3

FlowJo (v10)

Optional (but useful):

Box Sync Installer

Dropbox desktop app


A plasmid Editor (ApE)

Snapgene Viewer


Sim Daltonism


The Unarchiver

4 Peaks


GitHub Desktop

Zoom Desktop

Skype Desktop

OBS (Open Broadcast Software)

Useful installations from the command line


First go to the Samtools directory

$ ./configure

$ make

$ make install

$ export PATH=bin:$PATH


$ xcode-select –install

$ ruby -e “$(curl -fsSL”

$ brew doctor


$ brew install gifski


$ brew install ffmpeg


$ conda install -c conda-forge pypdf2

A note about editing your .bash_profile

Adding printers in WRB 5th floor

Note: You will not be able to set up the printer when on the “CaseGuest” wireless network. The printer can be setup while on the “CaseWireless” wireless network, though it may be hard to access it from the lab-space. In that case, a direct link to the ethernet is probably the best way to go. When connected to the land-line, you may first encounter an error saying the identity of any websites you go to cannot be confirmed. This may be because you are on a new computer, in which case you have to first register your computer with CWRU IT by typing in “” into a browser.

The most convenient printer is the black-and-white one near my office. To add this printer on a mac: 1) Go to Systems Preferences (such as through the apple icon op) 2) Go to Printers and Scanners 3) Press the “+” sign to add a printer 4) Type in the IP address “”. 5) The protocol should be “Line Printer Daemon – LPD” 5) You can rename it to something like “Nearby Black and White” to make it easier to remember. 6) It’s fine to leave Use as “Generic PostScript Printer” 7) Click on Duplex Printing (Since it’s useful). Voila!

Make a movie of a pymol structure spinning for powerpoint

UPDATE 5/18/2020: Well, for whatever reason, (at least my version of) Pymol stopped turning the camera (turn command) or molecule (rotate command) in script form, so the below no longer works. Buuutttt. I just told used the drop-down menu to do the movie > program > camera loop > y roll > 4 (or whatever) seconds. And then went to export movie in the file menu and made my movie that way. *shrug*

I’m sure there are many ways to do this, but this is the way I’ve been doing it most recently:

1) Set up your pymol session. That means importing your structure, turning the background to white, and enabling any other setting to make it the desired quality.

2) I use a custom python script to make a series of commands that make the structure turn slightly, ray-trace the structure (for a high quality image), and export the image to a PNG file. The script that I’ve linked to here makes pymol export 360 images, making the structure spin around completely.

Note: Depending on your settings, this process can take 10 minutes or 3 hours. An easy parameter to change would be the resolution (default setting in the script is 2000). Obviously the settings that are toggled during step 1 can drastically change how long it takes as well.

3) Use ffmpeg (at least, on a mac) to turn the 360 images into a video.

$ ffmpeg -framerate 20 -pattern_type glob -i '*.png' -c:v libx264 -preset slow  -profile:v high -level:v 4.0 -pix_fmt yuv420p -vf pad="width=ceil(iw/2)*2:height=ceil(ih/2)*2" -crf 22 -codec:a aac Output.mp4

I used homebrew to install ffmpeg last time I had to do it (brew install ffmpeg). I followed these instructions last time I had to install homebrew.

4) Drag into your powerpoint presentation, and voila!

VAMP-seq Tips

VAMP-seq is a valuable tool for identifying loss-of-function variants for a protein when you don’t have a specific assay for characterizing that protein’s activity. This is because almost all proteins can be “broken” by mutations that cause the protein to mis-fold, mis-traffick, or any other perturbations that may cause the protein to disappear from its normal cellular compartment. In the case of PTEN, we identified 1,138 variants that were lower than WT abundance. Notably, ~ 60 % of variants characterized as pathogenic in people were loss-of-abundance variants, confirming the importance of this property.

VAMP-seq uses a genetic fusion between your protein of interest and a fluorescent moeity (eg. EGFP) to assay its steady-state abundance. Fusions with unstable variants of your protein results in cells expressing unstable fusion proteins that don’t really fluoresce. In contrast, fusions with stable variants (such the WT protein) may fluoresce brightly. Unlike western blots which are ensemble measurements, this assay can be performed at a single-cell level. Correspondingly, this single-cell fluorescence readout makes it possible to test a large library of variants in parallel, in a multiplexed format.

Two VAMP-seq orientation discussed here

Still, when thinking about using VAMP-seq, you must first consider its limitations. While strong loss-of-abundance variants will be non-functional, variants of intermediate abundance may still be sufficiently functional in many contexts. Furthermore, while low abundance correlates with inactivity (and pathogenicity, in many clinical genetics contexts), WT-like abundance in no way indicates activity (or benignity when observed in people). For example, active site mutants often destroy protein function while having little to no effect on protein folding and abundance.

There are also proteins that are inherently incompatible with VAMP-seq. Secreted proteins won’t work because you lose the single-cell, genotype-phenotype link needed for the single-cell assay to work. Marginally stable or intrinsically disordered proteins likely won’t work due to a lack of destabilizing effect. Obligate heterodimers won’t work, though you may be able to get around it by overexpressing the protein partner, such as what I did with MLH1 for assessing PMS2 (See Fig 6). Proteins that cannot be tagged are problematic; this likely includes proteins that normally exist in crowded complexes, or that have key trafficking motifs on their termini. Proteins that are toxic to cells when overexpressed also poses problems, though one of my new landing pad platforms may help with that.

If your protein of interest passes those criteria, the rest is empirically confirming that there is enough signal over background to run the assay. A good literature search is a great place to start. You should look for 1) evidence that an N- or C-terminal tag works well (both good expression and normal protein activity), and 2) known destabilized variants that could serve as controls when performing the preliminary experiments. Ideally, there will be a clear difference in fluorescence distributions that are separable by thresholds used in FACS sorting, and that these differences are physiologically meaningful (as far as we know). Comparing /correlating the results of western blotting with the fluorescence distributions of EGFP-tagged protein is helpful (see below for PTEN). If the initial EGFP distributions between WT and the destabilized variants don’t seem super crisp, see what it looks like when you take the EGFP:mCherry ratio. As you can tell in the below figure, the EGFP:mCherry ratio is quite handy for increasing the precision of each distribution, as it divides out much of the heterogeneity in transcription / translation between cells.

Western and flow results

Regardless, I recommend to most people that they clone both N- and C- terminal fusions, and minimally look at the MFI values of the cells expressing each fusion, as low MFI will likely mean low dynamic range of the assay (from too little signal over background). Ideally, both WT and controls will be tested in both contexts. If the signal is relatively high but there’s concern that the large GFP fusion is causing problems, you could try 1-10/11 split EGFP. This only requires fusion with the ~ 15aa beta-strand 11 of EGFP (and separately co-expressing the larger fragment in the cells), though it’s not completely free of steric hindrance as it requires the spontaneous complex formation of the two subunits for fluorescence. This format also worked with PTEN (See SFig 1d), though I noticed a ~ 10-fold hit in overall fluorescence using the splitGFP format.

Fluorescence levels from different formats

Once it passes all those tests, then follow the steps in the Nature Genetics paper. Good luck!

Landing Pad Cell Line Generation

My favorite method of landing pad cell line generation is with lentiviral transduction. The original method was with site-specific knock-in using genomic disruption (Using Cas9 or TALENs), followed by homology-directed repair from a transfected circular plasmid template. It worked, but it was slow, with a fair number of false positives, including cells that had likely still expressed BFP transiently during single cell sorting, or even more annoyingly, cells where numerous landing pads had integrated per cell. I’ve since gotten some potentially helpful tips (eg. linear template supposedly degrades faster than circular plasmid), but I still don’t think this method is practical for routine transgenic knock-in. Moreover, I’m not convinced the AAVS1 locus is even that great for cell engineering.

This brings me to Lenti landing pad transduction, which I’ve developed a bunch of supported tools for. It’s easy (few steps), fast (cells can be single-cell sorted 2 – 7 days following transduction), broadly applicable (many different cell lines are easily transduced), with low false positives (low MOI transduction, due to Poisson behavior, ONLY gives single integrants). Pretty good arguments, no?

The general steps are as follows:

  1. Transfect HEK 293T cells with a mixture of lentiviral vector plasmids to produce lentivector particles. Do this in no dox media.
  2. Change media the next day, and then replace and collect supernatant over a ~ 72 hour period. Do this with no dox media. It’s fine to keep collecting into the same collection tube.
  3. After all of your collections are done, spin out any cells that may have been accidentally collected, and pass the supe through a 0.45 micron filter.
  4. Plate out your target cells, and mix with a series of volumes of the filtered supernatant. This can be in dox media.
  5. At least 2 days after switching into dox media, read out how many cells are fluorescent protein (often BFP) positive. Go with the well closest to but lower than ~5%. If the positive cells are incredibly few, treat cells with antibiotic (often blasticidin) to enrich for transductants.
  6. Using FACS, sort individual positive cells into wells of a 96 well plate. Use conditioned media if your cells may require it.
  7. Allow clonal lines to grow out. Transfect with a mixture of Bxb1 expression plasmid, attB-EGFP, and attB-mCherry to confirm no stable double positives exist. Cells that pass this test should be good to go for library work

Key viral non-coding elements are white rectangles. Coding regions are colored rounded rectangles. 2A sequences are red inverted triangles. Promoters are stemmed arrows. Terminators are thick black bars.

Now for the flavors of lenti-landing pad that I’ve developed. It’s very choose your own adventure, based on your needs. The original (LLP) is the most conservative choice to start with in non-HEK cell lines. LLP-Int-Blast can make your life easier with easier / higher recombination rates, but requires a bit more care in its use. If you want to study a potentially toxic gene, use LLP-rEF1alpha. For everyone else, use LLP-iCasp9-Blast. The selection with AP1903 is so quick and easy that it allows you to brute force recombinations quite easily. I used to chuckle whenever anyone used the phrase “game changer”, but I totally find it appropriate for this landing pad.

Gene knockouts by exon deletion with Cas9 transfection

Cas9 is a dream biotechnology, since you can use it to quite easily and specifically disrupt almost any DNA sequence you’re interested in modifying. Its been particularly helpful in work with cultured mammalian cells. In this document, I will describe how I use Cas9 to make gene knockouts in HEK 293T cells.

So although I’ve heard good things about transfecting guide-RNA loaded Cas9 ribonucleotide protein complexes into cells, I haven’t done this yet. Instead, I’ll describe a likely less optimal, but super cheap and easy method of co-transfecting Cas9 and guide-RNA expressing plasmids into cells to create knockouts. Specifically, I’ll describe my method of knocking out a gene by removing a big chunk of coding sequence, either ablating its start codon, or getting rid of an important exon to only create truncated or frameshifted non-functional proteins.

Equipment & Materials:

1) Tissue culture hood & incubator
2) Transfection reagent
3) Fluorescence Activated Cell sorter (FACS)
4) PCR machine, polymerase, Gibson master mix.
5) Agarose gel electrophoresis apparatus and some form of GelDoc.
6) Sanger sequencer (or sequencing service)

Step1: Make custom guide-RNAs for your gene of interest

As mentioned above (and shown in the figure), to create genetic knockouts, I like to delete out entire exons. The main reason is that it’s really easy to genotype with PCR. It’s also nice in that if the change is made, even if the repair is an in-frame indel, the deletion will be so large that the protein should still be loss-of-function. Lastly, exon deletion can often be achieved by targeting adjacent introns; this means that even if you stably express the Cas9 / gRNA combo, you can re-express a cDNA version and have it not be susceptible to disruption.

I first find guide RNAs that will target the DNA I want to disrupt. Addgene seems to have a pretty handy list of guide-RNA sequence design applications here: ( By default, I still tend to go to this classic website: ( Have it come up with a pair of seemingly specific guides, per the strategy above. Note: When using a U6 Pol3 promoter, I’ve read that if your guide RNA doesn’t start with a G, you should plan to include an extra G in front of the guide to increase transcription.

Addgene lists a bunch of protocols from the original labs here: ( Depending on how you like to do your molecular cloning, you may want to follow one of those. I’ll describe making a custom guide-RNA using Gibson Assembly, which is absolutely amazing.

Using a really basic U6-promoter guide-RNA expressing plasmid (like so: XXXX), design a pair of long primers that overlap in their 5′ ends with the guide-RNA sequence, but hybridize to sequences adjacent to the guide-RNA sequence but point in the opposite directions. The ~20nt guide-RNA sequence makes for a great Gibson overhang.

The steps here are:
1) Amplify plasmid, appending the guide-RNA sequence to the termini of the amplicon. I use Kapa Hifi, with 40ng template plasmid (30ul total), and ~7/8 cycles.
2) DPNI digest. I pipet 1ul of DPNI directly into the 30ul reaction tube, and incubate at 37*C for 2 hours (I suspect you don’t need all two hours).
3) “Purify” your intended plasmid. At first, I used to run my amplifications out on an agarose gel and gel extract (using a Qiagen gel extract kit). Nowdays, I just use a Zymo Clean and Concentrate kit (first setting aside some sample to run on a gel afterwards, for diagnostic purposes only). I elute in a small volume (6 ul for the Zymo kit).
4) I then make equal volume mixtures (eg. 1ul “insert”, 1ul “vector”), and mix in the corresponding amount of 2x Gibson Master Mix (So 2ul for the above example). I incubate this at 50*C for 0.5 to 1 hr. Standard protocol here.
5) I mix the sample (usually 4ul) into 50ul of chemically competent E.coli, and transform (eg. “Heatshock” transformation).
6) In the case of the Church lab plasmids, they’re Kan resistant, so don’t forget to recover before plating onto Kan plates.

Normally, I end up seeing tens to hundreds of colonies the next day. I then usually pick 2 colonies to grow up and miniprep. When I run Sanger, usually at least one colony contains plasmid DNA with the intended guide-RNA encoded. If neither colony has the intended plasmid, I normally pick two more and screen those. I don’t remember the last time this protocol failed (possibly hasn’t in my dozen attempts), but I think if it does it’s likely going to be from there being too much template, so I would then re-do the process really making sure DPNI is doing its job (and maybe gel extracting if I have to).

Step2: Transfecting the guide-RNA

I’ve only ever done this in HEK 293T cells, since that’s where I do most of my tissue culture work (since they’re so easy to handle). Next step is to transfect cells with the guide-RNA and Cas9 encoding plasmids. I tend to do this in 6-wells, using Fugene 6.

For this transfection, I either mix the guide-RNA plasmids (half plasmid for guide 1, the other half plasmid for guide 2) with PX458 (which encodes Cas9 2A-linked with EGFP), or a plasmid expressing untagged Cas9 along with a plasmid encoding a fluorescent protein. Since transfection usually results in many many copies (hundreds? thousands?) of plasmid getting in per cell, cells expressing the fluorescent protein (generally) should have been transfected with guide RNAs and Cas9 also (and thus are the cells most likely to have been edited).

Then, approximately two days after transfection, I sort for transfected cells using the fluorescent marker. I usually do this in a 96-well plate, sorting single cells into individual wells (to create clonal lines), while taking one of the corner wells and sorting 100+ cells into this one. This gives me an improved bulk population as backup in case the single clone growouts fail, and gives me a well that makes focusing onto the right focal plane a lot easier when using the light microscope (since hard to correctly focus on wells where it’s not clear if there are even any cells there).

If in a rush, I’ll check on the wells in approximately a week. By then, you can definitely see colonies of cells growing by brightfield microscope. I would then trypsinize and replate to keep them growing happily. If I’m not in a rush, I’ll sometimes let the cells grow for a couple weeks before checking; at this point, the media color starts becoming yellowish in wells containing growing cells, and clonal lines that grew out are large enough that if I transfer them to a new well, I’ll be able to do an expt with those cells in a few days.

Step3: PCR Genotyping

Once I have enough cells (say, at least half a 24-well plate well’s worth), I’ll extract genomic DNA using a Qiagen DNA easy kit. I’ll then use the primer combinations shown above in the figure. Notably, primer combo BC should only amplify if there is a wild-type copy present (DNA from unmanipulated cells should serve as a positive control). This should be a binary “amplifies or doesn’t” readout. The other primer combo also gives a slightly orthogonal perspective on what might be present at the intended locus within the genome. If unmodified, combo AC should give a large band (or no band, if these are really large introns). If modified, combo AC should give a much smaller product (I aim for a few hundred bases for easy visualization by agarose gel electrophoresis). To verify that this small band is what I think it is, I gel extract the band and sequence using Sanger. Conveniently, you can even use either primer A or primer B as the Sanger sequencing primer if you didn’t order a dedicated internal primer. In the handful of times I’ve done this, I think the predominant modified product that I’ve seen is literally a blunt-ended Non-Homologous End Joining of the two DNA fragments without any resection / removal of terminal nucleotides. Thus, if this is indeed the major modified product, I’m able to design my guides such that NHEJ ligation results in a frameshift.

Using the above PCR, I go through and screen the colonies to find one that looks to be a complete knockout (no amplification with primers BC, only small band with primers AC). Of course, actual protein knockout should be confirmed by western blot.

Voila! Knockout cells to now use in your experiments!

Some references:

I think this project / paper from super grad student Molly Gasperini in Jay Shendure’s lab is what originally made me think about using paired gRNA deletions to make knockouts.

I’ve tried to link to specific protocols in the above descriptions. For any details that are ambiguous, the methods described in my 2018 NAR paper should largely describe the basic steps / reagents used.

PS: Ed Anderson, a Postdoc at UNC Chapel Hill, made the excellent point that the molecular cloning in this post (ie. Gibson assembly steps) could be performed with In-Vitro Assembly (IVA). We certainly had some success with this method in my postdoc lab (albeit we used it for parallelized NNK library generation), though could certainly be used here for simpler and most cost-effective routine molecular biology.