Baylor Team Develops Dual-Color Reporter To Study Alternative Splicing Regulation

Name: Thomas Cooper

 

Position: Professor, pathology/molecular and cellular biology, Baylor College of Medicine, Houston

 

Background: Assistant/associate professor, pathology/molecular and cellular biology, Baylor, since 1989; Assistant research anatomist, University of California, San Francisco, 1986-1989; Postdoc, anatomy, UCSF, 1982-1986; MD, Temple University School of Medicine, 1982

 

 

Researchers led by Thomas Cooper from the Baylor College of Medicine in Houston recently published a paper in Nucleic Acids Research describing a bichromatic fluorescent reporter that expresses two different fluorescent proteins from one construct. They also demonstrated how the new reporter protein improves on existing fluorescence-based methods by quantitatively analyzing alternative splicing in single cells.

 

Cooper and colleagues believe that the method may help researchers better understand the regulation of splicing mechanisms, and may enable high-throughput screening of compounds that effect splicing regulation in mixed cell cultures and tissues of transgenic animals.

 

Cooper took a few moments this week to discuss his work with CBA News.

 

 

Alternative splicing starts with the idea that there are 20,000 to 25,000 genes in the human genome, but hundreds of thousands of proteins. Where did all those proteins come from, and why are there so few genes? It turns out that individual genes can make more than one protein, and they do that by this process of alternative splicing.

 

What we’re interested in is not the fact that you can get all of these different proteins – it’s that these proteins will have different functions; different isoforms from the same gene will have different functions; and the expression of those isoforms is regulated. There are tissue-specific forms; developmentally regulated forms, which is something we’re particularly interested in, where one isoform is expressed in an embryo and another in the adult.

 

It also turns out that there are some diseases – one in particular, called myotonic dystrophy – in which the transition from an embryonic splicing pattern to an adult splicing pattern is messed up. That’s a main feature of that particular disease, and there are probably other diseases where there is a screw-up in alternative splicing and the regulation of alternative splicing that’s causing the disease.

 

 

There are two main approaches. One is just to use a construct that expressed one fluorescent protein – one construct giving one fluorescent protein, and this is most commonly GFP or eGFP. This is basically an on-off assay. If the transgene spliced one way, it would put GFP in frame and you’d get a GFP readout. If it spliced the other way, you didn’t get a GFP readout – it was out of frame. That’s great, but the issue is that if you get a GFP readout, you don’t know whether only one percent of the RNAs were spliced in a certain way, and 99 percent did not; or it’s 99-percent spliced to give the GFP, and the rest is not. So you can’t quantify, you don’t get a ratio. And you really can’t look at two different cells that might have different regulation going on and quantify it very well.

 

The second method, which people have done to get around that, is to have some sort of internal standard. So they still have that same construct, where if it splices one way, you get GFP, and if it splices another way, you don’t. But then they put in another plasmid or transgene that expresses another fluorescent protein, commonly dsRed. You’re always doing a ratio of constant dsRed expression to GFP expression. That gives you kind of an internal standard, but there still could be variability between expressions of the different plasmids.

 

The way that we set it up, so that you’re expressing both proteins – dsRed and GFP – from one plasmid, is more sensitive, because if you are pushing a splicing pattern towards one, you’re reducing it from the other. There is a bigger change in the ratio of the two fluorescent proteins, whereas if you have a constant level of expression of one and you’re changing the other, you can see there is a change in the ratio, but it’s not as strong of a change.

 

 

Yes. You can take one plasmid, and have both dsRed and GFP in it, and use alternative splicing to change the reading frame, which is what we do. One splicing pattern is giving RNAs that only give dsRed, or they’re giving the other RNA which gives GFP. If you regulate an alternative splicing event, not only will it start making more GFP, it will make less dsRed. Also, you don’t have to worry about the variation in expression from two different plasmids. You know that this one plasmid is expressing two RNAs by alternative splicing, and one of those RNAs gives dsRed and the other gives GFP.

 

 

Right. And I think [fluorescence-activated cell sorting] analysis is important here, too. I think if people wanted to have a population of cells that are undergoing alternative splicing, you can put in whatever gene you want into this vector, and design it so it will be regulated under some signaling event. You can starve cells or add insulin and cause a change in splicing. And if you want to detect those cells that really have a very strong effect on splicing, you should be able to do that by FACS analysis.

 

 

No. The idea was out there. GFP and dsRed are out there. But here’s the trick: If you look at the open reading frame of dsRed, it starts with ATG and ends with a stop codon. But if you look at the other two reading frames, there is one reading frame that has absolutely no start codons and no stop codons. That was the key. All we noticed was that there was this complete read-through reading frame of dsRed. And it turns out that eGFP has it, as well. It’s weird. Usually, if you look at a 700- or 800-nucleotide segment, you’re going to find stop codons in all three reading frames. I think that the organisms that [naturally] express these proteins – they’re doing something with that. For some reason they don’t want stop codons. What it allowed us to do is to clone dsRed upstream of GFP, and when we switch the reading frame, we can either express the dsRed reading frame, or we can shift it by one nucleotide, put it into that “no-stop codon” reading frame, and it just reads through into the GFP. It’s just a way for us to express both proteins from the same construct. That’s basically what we did, to make that observation.

 

 

We didn’t play around with plate readers, but I think it would work. And actually, that’s what we’re thinking about trying. Baylor actually has a very nice setup where they do robotic screening based on plates, using bifluorescence microscopy and other methods. This would work great for that. That’s more along the lines of what we were thinking about using it for.

 

The other thing that would be interesting to use this for is transgenic animals. We’re so much at the tip of the iceberg at understanding not just how alternative splicing is regulated, but how much regulation there is out there. It looks like there is a lot more than we initially thought – that a lot of alternative splicing events are regulated. And there are people, including us, who are doing splicing microarrays, and you can see all kinds of changes that are taking place on a genome-wide scale. I think it’s neat to think about putting it into mice, but we haven’t thought about a specific question to ask. I think we would find that a splicing event that you assume is heart-specific or brain-specific – if you put it into a mouse, you would see, cell by cell, very specific differences. With one transgene, you can look at a cell-by-cell basis that two different cells beside each other of different cell types have different splicing patterns, and you can quantify that ratio.

 

 

One thing is that you can use it to identify things that regulate splicing. I think you can also use it to identify small molecules that prevent or enhance a particular splicing event. In addition, there are a lot of mutations that cause disease, and do so by not messing up the trans factors that regulate splicing, but instead it’s a mutation within the exon or the splice site [next to] the exon that causes the exon to not be recognized. This isn’t even alternative splicing – this could be a constitutive exon. It would be very straightforward to put that genomic segment containing that exon and some of the flanking introns into this reporter, and then to screen for small molecules that would enhance inclusion of that exon. So this might be useful as a method for gene therapy. The example we used in the paper was spinal muscular atrophy. There are a number of labs that are trying to address this, because it’s actually like the case I just described – there is a mutation in the exon that causes it to be skipped, and it’s a devastating disease. The exon isn’t completely skipped – it’s like 10-percent included – it’s just not enough to make enough protein to ameliorate the problems. If you could find something that could cause that exon to be included a little more, you could really help these people.

 

 

I would think so. Everything came from already available technologies. I tried to see if I could get some licensing rights or something, but Baylor didn’t think so. If somebody commercializes this, then that would be great – I just hope they reference the paper, because that’s about all I’m going to get out of it. [Laughs]. There is not a huge number of labs that work on the regulation of alternative splicing, although it’s growing. But I would think that more human geneticists might be interested in it as a way to see if they can reverse a mutation that causes a splicing defect.