The Stewart Shuman Lab: Research Overview

The goal of my research is to understand the mechanisms and structures of enzymes that perform and regulate essential nucleic acid transactions. My research integrates diverse experimental approaches (including virology, biochemistry, structural biology, and genetics) and applies them to model systems ranging from viruses to bacteria to fungi to mammalian cells. An explicit aim is to identify novel enzymatic targets for treatment of human diseases.

My interest in infectious pathogens as model systems derives from an early appreciation from my Ph.D. studies of the central role of viruses and bacteria in understanding gene expression and replication strategies, plus a fascination with infectious diseases acquired during my clinical training at Mass General. After completing my residency in Internal Medicine, I did a fellowship in the Laboratory of Viral Diseases at NIH. I continued to focus on problems in virus gene expression and replication when I first established my own lab at Sloan-Kettering in 1988. My research interests have expanded since then to embrace other model systems and biological problems. I have especially tried to translate insights gained from studies of viral RNA/DNA metabolism to analogous (and often more complex) regulatory problems in cellular biology. In particular, I have become interested in the evolution of enzyme systems and pathways and the ancestral connections between RNA and DNA metabolism. For example, we have explored the nature of a key eukaryote-specific RNA processing event (mRNA capping) in a wide variety of species and viruses. Our findings have yielded insights to eukaryal phylogeny and highlighted capping as a promising target for antiviral, antifungal, and antiprotozoal drug discovery. Our studies of the central role of the RNA polymerase II (Pol2) carboxyl-terminal domain (CTD) and CTD phosphorylation in directing the capping enzymes to nascent pre-mRNAs helped establish the concept of a CTD code.

Our studies of DNA modification enzymes have revealed the mechanism and evolutionary history of topoisomerase IB. Our work on DNA repair, centered on DNA ligases, has broad evolutionary and mechanistic significance, as well as clear implications for targeting DNA ligases for treatment of bacterial infections.

We have extended these studies to embrace RNA damage and RNA repair reactions involving site-specific endoribonucleases and RNA ligases.

I have a strong commitment to, and extensive experience in, the teaching and training of graduate students and postdocs. I have mentored 33 graduate students who received their doctorates for thesis research conducted in my lab. Many of my former trainees are now in faculty positions at academic institutions in the US, Canada, the UK, and France; other alumni are working as research group leaders in the US pharma and biotech sector.

Five areas of inquiry in my laboratory are summarized briefly below.

1. DNA topoisomerase mechanism and structure: We’ve developed vaccinia topoisomerase as a highly instructive model for the eukaryal DNA topoisomerase IB (TopIB) family. We’ve shown that the TopIB catalytic repertoire is broader than suspected and have uncovered novel aspects of its mechanism via ingenious DNA chemistry and protein mutagenesis. We leveraged these discoveries to invent a clever TOPO-cloning technology that is used widely.

Shuman, S. (1994) Novel approach to molecular cloning and polynucleotide synthesis using vaccinia DNA topoisomerase. J. Biol. Chem. 269, 32678-32684.
Sekiguchi, J., and Shuman, S. (1997) Site-specific ribonuclease activity of eukaryotic topoisomerase I. Molecular Cell 1, 89-97.
Cheng, C., Kussie, P., Pavletich, N., and Shuman, S. (1998) Conservation of structure and mechanism between eukaryotic topoisomerase I and site-specific recombinases. Cell 92, 841-850.
Krogh, B.O. and Shuman, S. (2000) Catalytic mechanism of DNA topoisomerase IB. Molecular Cell 5, 1035-1041.

2. mRNA capping: The 5’ m⁷GpppN cap is a signature feature of eukaryal mRNA that is required for mRNA stability and efficient translation. We have illuminated the structures, mechanisms, and evolutionary histories of the triphosphatase, guanylyltransferase, and methyltransferase enzymes that perform mRNA capping. We discovered the distinctive properties of viral/fungal RNA triphosphatases and (in collaboration with Chris Lima) the novel structure of yeast RTPase, the founder of a “triphosphate tunnel metalloenzyme” superfamily. We showed (in collaboration with Alfonso Mondragon) that mammalian RTPase has an entirely different structure and mechanism. Our work recommends capping as a target for antiviral, antifungal, antiprotozoal drug development.

Shuman, S., and Hurwitz, J. (1981) Mechanism of mRNA capping by vaccinia virus guanylyltransferase: characterization of an enzyme-guanylate intermediate. Proc. Natl. Acad. Sci. USA 78, 187-191. PMCID: PMC319016
Shuman, S., Liu, Y., and Schwer, B. (1994) Covalent catalysis in nucleotidyl transfer reactions: essential motifs in Saccharomyces cerevisiae RNA capping enzyme are conserved in Schizosaccharomyces pombe and viral capping enzymes and among polynucleotide ligases. Proc. Natl. Acad. Sci. USA 91, 12046-12050. PMCID: PMC45373
Lima, C.D., Wang, L.K., and Shuman, S. (1999) Structure and mechanism of yeast RNA triphosphatase: an essential component of the mRNA capping apparatus. Cell 99, 533-543.
Changela, A., Ho, C.K., Martins, A., Shuman, S., and Mondragon, A. (2001) Structure and mechanism of the RNA triphosphatase component of mammalian mRNA capping enzyme. EMBO J. 20, 2575-2586. PMCID: PMC125469

3. DNA repair: DNA ligases are the sine qua non of genome integrity. They are essential for DNA replication and repair in all organisms. Our biochemical and crystallographic studies of bacterial and viral DNA ligases revealed the structural basis of nick recognition and the chemical and kinetic mechanisms of 3’-OH/5’-PO₄ end sealing. Our elucidation of the mycobacterial NHEJ pathway (in collaboration with Mike Glickman) and its central catalyst LigD has instated a new paradigm of microbial DNA repair/recombination. In the area of DNA double-strand breaks repair, we discovered a new mycobacterial helicase-nuclease that catalyzes end-resection during homologous recombination.

Nandakumar, J., Nair, P.A., and Shuman, S. (2007) Last stop on the road to repair: structure of E. coli DNA ligase bound to nicked DNA-adenylate. Molecular Cell 26, 257-271.
Nair, P.A., Nandakumar, J., Smith, P., Odell, M., Lima, C.D., and Shuman, S. (2007) Structural basis for nick recognition by a minimal pluripotent DNA ligase. Nature Struct. Mol. Biol. 14, 770-778.
Aniukwu, J. Glickman, M.S., and Shuman, S. (2008) The pathways and outcomes of mycobacterial NHEJ depend on the structure of the broken DNA ends. Genes & Development 22, 512-527. PMCID: PMC2238672
Sinha, K.M., Unciuleac, M.C., Glickman, M.S., and Shuman, S. (2009) AdnAB: a new DSB-resecting motor-nuclease from Mycobacteria. Genes & Development 23, 1423-1437. PMCID: PMC2701575

4. RNA repair: Our work is driving an emerging appreciation of the existence of “RNA repair” pathways that rely on RNA ligases to maintain or manipulate RNA structure in response to biologically purposeful breakage events. Our longstanding hypothesis is that many flavors of RNA repair enzymes exist and that they can be uncovered by integrating biochemical and structural insights to ligase mechanism with phylogenetic analyses of cellular and viral proteomes. This has been a fruitful effort in our hands that has resulted in the discovery of many new RNA repair enzymes and the delineation of novel RNA repair pathways. In particular, our discovery of bacterial RtcB as a 3’-PO₄/5’-OH ligase that acts via a covalently activated RNA(3’)pp(5’)G intermediate has overturned the textbook view of how nucleic acid phosphodiesters are synthesized by enzymes.

Nandakumar, J., and Shuman, S. (2004) How an RNA ligase discriminates RNA damage versus DNA damage. Molecular Cell 16, 211-221.
Nandakumar, J., Shuman, S., and Lima, C.D. (2006) RNA ligase structures reveal the basis for RNA specificity and conformational changes that drive ligation forward. Cell 127, 71-84.
Nandakumar, J., Schwer, B., Schaffrath, R., and Shuman, S. (2008) RNA repair: an antidote to cytotoxic eukaryal RNA damage. Molecular Cell 31, 278-286. PMCID: PMC3289999
Chakravarty, A.K., Subbotin, R., Chait, B.T., and Shuman, S. (2012) RNA ligase RtcB splices 3’-phosphate and 5’-OH ends via covalent RtcB-(histidinyl)-GMP and polynucleotide-(3’)pp(5’)G intermediates. Proc. Natl. Acad. Sci. USA 109, 6072-6077. PMCID: PMC3341019

5. The RNA polymerase II CTD Code: The carboxyl-terminal domain (CTD) of the Rpb1 subunit of RNA polymerase II (Pol2) consists of tandemly repeated heptapeptides of consensus sequence Y¹S²P³T⁴S⁵P⁶S⁷. The CTD is essential for cell viability because it recruits proteins that regulate transcription, modify chromatin structure, and catalyze or regulate mRNA capping, splicing, and polyadenylation. The inherently plastic CTD structure is modulated by phosphorylation of the heptad serine, threonine, and tyrosine residues. The combinatorial complexity of the CTD serine-2,5,7, threonine-4, and tyrosine-1 phosphorylation array is enormous. The instantaneous primary structure of the CTD provides informational cues about the state of the transcription machinery – a CTD code – that is “read” by CTD receptor proteins, including the mRNA capping apparatus. We have focused on the role of the CTD in directing m⁷G capping to nascent Pol2 transcripts via the binding of RNA guanylyltransferase (GTase; capping enzyme) to the Ser5-phosphorylated CTD. In collaboration with Chris Lima, we elucidated the structural principles underpinning capping enzyme•CTD interactions by co-crystallizing budding yeast, fission yeast, and mammalian GTases bound to Ser5-phosphorylated Pol2 CTD peptide ligands. A salient (and surprising) insight was that capping enzymes from three different taxa have evolved three distinct strategies to read the Ser5-PO₄ “letter” of the CTD code. In collaboration with Beate Schwer, we are currently endeavoring to decipher the informational rules for the CTD code by genetically manipulating the composition and structure of the fission yeast Pol2 CTD. By introducing alanines and conservative mutations, singly or pairwise in all heptad repeats, we showed that: (i) the Tyr1, Pro3, Ser5, and Pro6 letters are essential for viability, whereas Ser2, Thr4, and Ser7 are not; (ii) Phe is functional in lieu of Tyr1; and (iii) Ser5 is the only strictly essential phosphorylation site. Our studies are revealing how CTD coding letters are assembled into “words” (i.e., a vocabulary) and how CTD coding cues govern specific cellular gene expression programs.

Schwer, B., and Shuman, S. (2011) Deciphering the RNA polymerase II CTD code in fission yeast. Molecular Cell 43, 311-318. PMCID: PMC3142328
Schwer, B., Sanchez, A.M., and Shuman, S. (2012) Punctuation and syntax of the RNA polymerase II CTD code in fission yeast. Proc. Natl. Acad. Sci. USA 109, 18024-18029. PMCID: PMC3497752
Schwer, B., Bitton, D.A., Sanchez, A.M., Bähler, J., and Shuman, S. (2014) Individual letters of the RNA polymerase II CTD code govern distinct gene expression programs in fission yeast. Proc. Natl. Acad. Sci. USA 111, 4185-4190. PMCID: PMC3964054
Doamekpor, S.K., Sanchez, A.M., Schwer, B., Shuman, S., and Lima, C.D. (2014) How an mRNA capping enzyme reads distinct RNA polymerase II and Spt5 CTD phosphorylation codes. Genes & Development 28, 1323-1336. PMCID: PMC4066402