About Our Lab
Our lab studies how macromolecular machines are assembled and function. We are primarily focused on the components of the yeast spliceosome and how spliceosomes are built. In addition, we collaborate with a number of laboratories on campus and elsewhere that are interested in RNA biochemistry and/or macromolecular machines. We use a combination of single molecule fluorescence, chemical, and biological tools to investigate the following areas:
In the pre-spliceosomal A complex, both the 5'SS and branchsite are identified by basepairing with the U1 and U2 snRNPs, respectively. In some organisms the U1 and U2 must also functionally interact with one another to form an intron-defined complex that is critical for determining which RNA sequences are removed and which are kept during alternative splicing. Using a combination of single molecule and biochemical approaches, we are investigating how the U1 and U2 snRNPs are recruited to the correct locations on a pre-mRNA. As part of this process, we are reconstituting and characterizing the spliceosomal E complex (i.e., Commitment Complex) in order to elucidate the mechanism by which this is transformed into A complex. Finally, in collaboration with Charles Query (Albert Einstein College of Medicine), we are studying how the DEAD-box ATPase Prp5 assists in docking of the U2 snRNP onto the branchsite and how ATP hydrolysis by Prp5 can be coupled to the fidelity of branchsite selection.
The tri-snRNP is assembled from the U4, U5, and U6 snRNPs and is the most dynamic subunit of the spliceosome. During each round of splicing, the spliceosome is activated and the tri-snRNP is extensively remodeled—so much so that after each turnover the tri-snRNP must be regenerated from the free U4, U5, and U6 snRNPs. ~24,000 free U4, U5, and U6 snRNPs are generated in human cells each minute as a consequence of splicing, and tri-snRNP recycling must be extremely efficient to maintain splicing activity within the nucleus. We are using single molecule colocalization, single molecule FRET, and single molecule pull-down in order to study pathways for tri-snRNP assembly and disassembly. Many of these experiments are carried out in collaboration with David Brow (UW-Madison, Biomolecular Chemistry) and Sam Butcher (UW-Madison, Biochemistry). Our tri-lab collaboration is using a combination of biochemical, single molecule, genetic, and structural techniques to study the dynamics of the U6 snRNP during tri-snRNP assembly and splicing.
Imaging RNAs by fluorescence microscopy has proven to be incredibly informative about RNA conformation, regulation, and localization. However, attaching fluorophores to specific RNA transcripts—particularly those that are heavily processed, structured, or only a few dozens of nucleotides long—is extremely challenging both in vitro and in vivo. Using a variety of approaches, we are developing methods to label RNAs in the test tube and in live cells. These methods will facilitate biophysical studies of RNA in vitro and enable single RNA molecule microscopy in live cells.
Cells rely on motor proteins such as dynein and kinesin to transport organelles, pro
teins, and RNAs from one location to another. This is particularly important in neurons where specific iomolecules must be localized to either the axon or dendrite. In collaboration with Professor Jill Wildonger (UW-Madison, Biochemistry), we are using single molecule fluorescence assays of Drosophila dynein motility to elucidate the connections between the biochemical properties of the dynein motor in vitro and Drosophila neurobiology in vivo.
Collaborators
Charles Query
David Brow
Sam Butcher
Eric Strieter
Jill Wildonger