Blog post by Dr. Tamara Maiuri
I am still busily collecting cells to be sent for mass spec for our goal of obtaining a list of proteins that interact with huntingtin upon oxidative DNA damage. Unfortunately I’ve run into a few road blocks, which I will blog about in the coming weeks (hopefully with a resolution!).
Meanwhile, I’ve been working on methods to assess the hit proteins for their physiological relevance as potential drug targets. Last time I described one such approach: the GFP reactivation assay. Since then, data from 3 experiments have been combined and look promising. While repair efficiency varies from experiment to experiment, mouse HD cells consistently show approximately half the repair efficiency of normal cells (an average of 44.8% over 3 experiments). This is a readout we can use to test the effects of manipulating our hit proteins.
Another approach involves measuring how long huntingtin hangs around at sites of DNA damage, and whether expanded huntingtin lingers too long. We know expanded huntingtin has no trouble reaching damaged DNA, so maybe the problem is that it can’t get off, inappropriately gluing down all the proteins it is scaffolding.
To test this hypothesis, we first need a way to visualize huntingtin protein at sites of DNA damage. While most researchers use overexpression (getting cells to generate protein from externally supplied DNA) to visualize their protein of interest, this is very difficult with huntingtin because of its huge size. To get around this, many HD researchers express small fragments of the huntingtin protein. Overexpression of any protein can have pitfalls because it’s impossible to know if the overexpressed protein is behaving the way it would under normal expression levels in the cell. This is especially true if you’re only using a fragment of the protein—what if the fragment doesn’t fold into the same shape that it would as a whole? What if the missing parts of the protein interact with other important proteins?
Exciting new technologies now allow us to track the behaviour of endogenous huntingtin protein (the huntingtin existing naturally in the cell). We put to use two different intracellular antibodies, or “intrabodies” that recognize and bind the huntingtin protein. We tagged these intrabodies with yellow fluorescent protein (YFP) to generate “chromobodies”. This allowed us to follow their interaction with endogenous huntingtin in live cells. Indeed, we could watch the endogenous huntingtin protein being recruited to sites of DNA damage.
This tool is not without its drawbacks. While it doesn’t seem to interfere with huntingtin’s recruitment to damaged DNA, it must interfere with its role in cell proliferation. We know this because we can’t stably express the chromobody in cells over time. When we watch cells expressing the chromobody try to divide, they just die (unlike the cells expressing only YFP, which happily multiply).
This roadblock can be hurdled using an inducible system: the cells carry the DNA expressing the chromobody, but it isn’t turned on to generate protein until you add a drug called doxycycline. So I first cloned the chromobody into an inducible vector (cloning experiment deposited to Zenodo). When co-transfected with the doxycycline-responsive Tet3G transcriptional activator, it showed beautiful induction by doxycycline in mouse striatal cells (induction experiment deposited to Zenodo).
But we want to work with cells from HD patients. It’s harder to get DNA into these cells, but we can do it with electroporation. To avoid this labour-intensive process every time I want to do an experiment, I’m making HD patient cell lines that stably express the inducible chromobody and doxycycline-responsive Tet3G activator. The Tet3G vector carries a drug resistance gene, so I can select the cells with the drug G418. A simple experiment (deposited to Zenodo) showed that the optimal concentration for G418 selection in fibroblasts is 50 ug/mL.
At this point, my luck ran out. The beautiful induction I saw in mouse striatal cells did not happen in HD patient fibroblasts. From the first few failed attempts, I learned the following:
- If cells become too sparse during the G418 selection process, they die. Need to transfect a larger number of cells so that they can be downsized during the selection process and still maintain confluency >50% for cell health.
- Transfection of fibroblasts is an issue. Need to use electroporation, and co-transfect H2B-mCherry to identify transfected cells
- Transfection of pTRE-nucHCB2 (inducible chromobody), pEF1a-Tet3G (doxycycline-responsive transcriptional activator), and H2B-mCherry is far more toxic than the equivalent microgram quantity of sonicated salmon sperm DNA. Need to use pTRE-nucHCB2, sssDNA, and H2B-mCherry as the untransfected control (that is, -Tet3G) in order to compare rates of selection by G418 in untransfected versus transfected cells
- In contrast to striatal cells, fibroblasts don’t seem to be inducing expression of nucHCB2 with doxycycline
After ruling out protein turnover, FBS concentration in the media, and different preps of DNA, the only difference between the nice result in mouse striatal cells and the confusing result in human fibroblasts is the method of transfection (the easier, polymeric method for striatal cells versus the more tedious—and expensive—electroporation method for fibroblasts). But this couldn’t possibly be the problem… could it? Only one way to find out: I set up a direct comparison experiment. To my great surprise, the striatal cells induced when transfected by the polymeric method, but not by electroporation! The experiment is posted on Zenodo.
At this point I recalled a suggestion made a few weeks prior, by Claudia Hung, a student in the lab: she asked whether the size of the plasmids could explain the results. I really didn’t think so at the time, but now that idea might make sense! The Tet3G vector is pretty large (7.9 kb), and sure enough, difficulty transfecting large vectors by electroporation is well documented (once you look for it!). This study by Lesueur et al explains that simply giving the cells a chance to recover from the electroporation before plating them can greatly enhance cell viability and transfection efficiency. This was my next move. There was a glimmer of hope in the results: the longer recovery time resulted in induction in a few cells. After taking a closer look at the Lesueur et al study, in which they used much larger amounts of DNA, I tried increasing the amount of DNA.
Eureka! Finally, after months of trouble shooting, I found conditions in which we can induce expression of a huntingtin-specific chromobody in cells from an HD patient (see the results on Zenodo). Next week I will be electroporating cells from HD patients who have different CAG lengths in their huntingtin genes, and selecting them in G418 to get stable cell lines. The result will be a panel of cell lines with different-sized huntingtin expansions, in which we can visualize the natural huntingtin protein by dropping in doxycycline—a great tool for our lab and HD researchers around the world.
If you’ve made it this far through this tedious blog post, thanks for reading. You now have a sense of the tiny incremental steps it takes to move a project forward. This is only one facet of a much larger goal, and each facet has its own set of obstacles. But with careful, calculated perseverance we can get through each road block and move our understanding of HD forward. This work is funded by the HDSA Berman/Topper HD Career Development Fellowship.