\chapter{Introduction} \label{ch:1}
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\begin{quote}
``It's a magical world, Hobbes, ol' buddy \ldots \\ 
Let's go exploring!''\newline\\
-- Calvin (Calvin and Hobbes,  Bill Watterson)
\end{quote}
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\noindent A portion of this chapter originally appeared as~\cite{Janetzko:2014fy}:\\%\newline\\
``The Making of a Sweet Modification: Structure and Function of \textit{O}-GlcNAc Transferase''
\textbf{John Janetzko} and Suzanne Walker.
\textit{The Journal of Biological Chemistry}. Volume 289, Number 50: 34424--34432. 2014.
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\section{Post-translational modifications galore!}
The human proteome is vastly more complex than the human genome~\cite{Wilhelm:2014jo}: while the human genome encodes on the order of 20,000 genes~\cite{InternationalHumanGenomeSequencingConsortium:2004bm}, the human proteome has been estimated to include more than one million proteins~\cite{NorregaardJensen:2004gz}. A major source of diversity in going from genes to proteins is through protein post-translational modifications (PTMs). Two of the most widely known PTMs include phosphorylation (on serine, threonine and tyrosine) and glycosylation (both \textit{O}-linked, occurring on serine and threonine, and \textit{N}-linked, occurring on asparagine). It has been estimated that more than half of all proteins are glycosylated~\cite{Khoury:2011dd,Palaniappan:2016bs}. Given the tremendous scale and the proteome, and the diversity of protein PTMs (Figure~\ref{fig:intro-PTMs}), it should not be surprising that until recently PTMs were generally characterized on a single-protein basis~\cite{Doll:2015ev}, and often with the exact site of modification remaining unknown~\cite{Cox:2011ck}.
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.7\columnwidth]{intro/PTMs.eps}}
	\caption[Experimentally observed PTMs]{Experimentally observed PTMs. There were 87,308 experimentally observed PTMs for 530,264 proteins in the July 2011 release of the Swiss-Prot database. This figure was reproduced from reference:~\cite{Khoury:2011dd}.}
	\label{fig:intro-PTMs}
\end{figure}
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%\section{Glycosyltransferases}
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\section{\textit{O}-GlcNAc transferase and protein \textit{O}-GlcNAcylation}
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Until the mid 1980s, many believed that glycoconjugates were exclusively derived from the secretory pathway~\cite{Varki2009ab}. Thirty years ago, when Torres and Hart made the unexpected discovery that many intracellular proteins are modified with \textit{O}-GlcNAc~\cite{Holt:1986vh,Torres:1984vu}, this set in motion a change in paradigm for localization of glycoconjugates. Thirteen years later the enzyme responsible for this post-translational modification was finally cloned~\cite{Kreppel:1997va,Lubas:1997vw}. Unlike the many glycosyltransferases (Gtfs) that reside in the endoplasmic reticulum (ER) or Golgi apparatus, \textit{O}-linked-$\beta$-\textit{N}-acetylglucosamine transferase (OGT) localizes to both the nucleus and cytoplasm of metazoan cells~\cite{Janetzko:2014fy,Varki2009ab}. OGT catalyzes transfer of a GlcNAc moiety from UDP-GlcNAc to specific serine and threonine residues in its many cellular targets (Figure~\ref{fig:glcnac-rxn})(henceforth termed \textit{O}-GlcNAcylation). Remarkably, it is the sole enzyme responsible for this abundant PTM~\cite{Haltiwanger:1992vs,Kreppel:1997va,Lubas:1997vw}. In mammalian cells, OGT is essential throughout life; however, the reason for this essentially is currently unknown~\cite{Levine:2016du}.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.8\columnwidth]{intro/ogt-rxn.eps}}
	\caption[OGT transfers GlcNAc from UDP-GlcNAc to Ser/Thr residues]{OGT transfers GlcNAc from UDP-GlcNAc to Ser/Thr residues.}
	\label{fig:glcnac-rxn}
\end{figure}
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The co-substrate for protein \textit{O}-GlcNAcylation~\cite{Haltiwanger:1990tk}, UDP-GlcNAc bears intimate links to cellular metabolism~\cite{Levine:2016du,Scott:2017kb,Zibrova:2017ct}. UDP-GlcNAc is produced through the hexosamine biosynthetic pathway (HBP, Figure~\ref{fig:glcnac-cycling-hbp}a). About 2\% of cellular glucose enters the HBP, via the rate-limiting enzyme glutamine-fructose amidotransferase (GFAT)~\cite{Levine:2016du,Traxinger:1991vn}. UDP-GlcNAc combines elements of glucose, glutamine, fatty acid metabolism and nucleotide biosynthesis, and the cellular concentration of UDP-GlcNAc has been shown to vary depending on the flux through these pathways~\cite{Hart:2011kv,Marshall:2004kv,McClain:2002en,Weigert:2003ce}.\par
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A glycosidase, \textit{O}-GlcNAcase (OGA), capable of hydrolyzing the $\beta$-\textit{O}-GlcNAc linkage, is also ubiquitous within the nucleus and cytoplasm of metazoan cells~\cite{Alonso:2014eq}. As a result, protein \textit{O}-GlcNAcylation is a dynamic PTM, able to turnover multiple times over a protein's lifetime (Figure~\ref{fig:glcnac-cycling-hbp}b)~\cite{Butkinaree:2010ga,Chou:1992wt,Hart:2007dy,Kreppel:1997va,Roquemore:1996bq}. The dynamic nature of this modification in many ways makes it analogous to protein phosphorylation. However, unlike protein phosphorylation, for which there are 428 Ser/Thr kinases in the human genome~\cite{Shi:2009bh}, only OGT is responsible for all cellular \textit{O}-GlcNAc~\cite{Hart:2011kv}.\par
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\begin{figure}[b]
	\caption[The role of OGA and the HBP for protein \textit{O}-GlcNAcylation]{\textit{(On the following page)}The role of OGA and the HBP for protein \textit{O}-GlcNAcylation. (a) The hexosamine biosynthetic pathway. Glucose (Glc) is converted to glucose-6-phosphate (Glc-6-P) by hexokinase (HK), which is isomerized to fructose-6-phosphate (Fruc-6-P) by glucose-6-phosphate isomerase (G6PI). Fruc-6-P is converted to glucosamine-6-phosphate (GlcN-6-P) by glutamine:fructose-6-phosphate amiotransferase (GFAT). GlcN-6-P is converted to \textit{N}-acetylglucosamine-6-phosphate (GlcNAc-6-P) by EMeg32, using acetyl coenzyme A (CoASAc); CoASH is free coenzyme A. GlcNAc-6-P is isomerized by phosphoGlcNAc mutase (PGM) to GlcNAc-1-P. Finally, UDP-GlcNAc pyrophosphorylase (UGPi) transfers uridine diphosphate from uridine triphosphate (UTP) to produce UDP-GlcNAc and inorganic pyrophosphate (PPi). The enzyme GlcNAc kinase (GK) is capable of phosphorylating GlcNAc with adenosine triphosphate (ATP), allowing for GlcNAc to be recycled~\cite{Levine:2016du}. (b) OGA is a hydrolase that removes $\beta$-\textit{O}-GlcNAc linkages from proteins.}
	\label{fig:glcnac-cycling-hbp}
\end{figure}
\begin{figure}[p]
	\centerline{\includegraphics[width=0.8\columnwidth]{intro/ogt-cycling-1.eps}}
\end{figure}
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Perhaps it is not surprising, given the large and diverse set of protein targets modified by OGT, that protein \textit{O}-GlcNAcylation has been implicated in regulating gene expression~\cite{Gambetta:2015gt,Hanover:2012hk,Liu:2017da,Ozcan:2010fq}, cellular stress responses~\cite{Kazemi:2010dn,Zachara:2004by,Zachara:2004eg}, metabolism~\cite{Dentin:2008bw,Ruan:2014jr,Yang:2012cv,Yi:2012ju}, and proteostasis~\cite{Guinez:2004gu,Han:1997uj,Zhang:2003wl,Zhu:2015bq} in response to nutrient levels~\cite{Bond:2015iy,Hart:2011kv,Harwood:2014il,Zachara:2004kn}, to name just a few~\cite{Levine:2016du}. Furthermore, given the myriad processes affected by OGT, there have been numerous associations between OGT and human disease. Perturbations in OGT expression level and/or global protein \textit{O}-GlcNAcylation have been observed in cancer~\cite{Ma:2013bd}, diabetes~\cite{Hart:2011kv,Vaidyanathan:2014ir}, Alzheimer's disease~\cite{Yuzwa:2014gk}, and cardiovascular disease~\cite{Marsh:2014fh}. Whether OGT will prove to be a meaningful therapeutic target, however, remains to be seen~\cite{OrtizMeoz:2015jo}.\par
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\newpage
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This chapter will focus on structural and biochemical studies of OGT, including a discussion of its role in HCF-1 cleavage. Numerous comprehensive reviews of these topics have been published, including some of the earlier OGT work not discussed here. For additional reading, please see:~\cite{Gloster:2010tp,Hart:2007dy,HurtadoGuerrero:2008ku,MartinezFleites:2010cw,Vocadlo:2012gm}.\par
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\section{Structure of \textit{O}-GlcNAc transferase} \label{sec:intro-ogt-structure}
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The \textit{OGT} gene, located on the X chromosome, encodes a protein with two major domains~\cite{Hanover:2003tf}. The N-terminal domain comprises a cluster of repeating motifs called tetratricopeptide repeats (TPRs), and the C-terminal domain bears the glycosyltransferase activity~\cite{Kreppel:1997va,Lubas:1997vw} (Figure~\ref{fig:ogt-domain-tpr-structure}a). TPRs are degenerate 34-amino acid motifs that typically fold into paired anti-parallel $\alpha$-helices~\cite{Allan:2010de,Lamb:1995to,Zeytuni:2012hf}. Often found in tandem arrays, TPRs are known for their role in mediating protein-protein interactions~\cite{Allan:2010de,Blatch:1999cu,Das:1998gc}. Evidence implicated the TPRs of OGT in mediating interactions with protein substrates, as well as putative adapters and proteins shown to affect its enzymatic activity \cite{Chen:2012cc,Cheung:2008hl,Iyer:2003ge,Iyer:2003eq,Kreppel:1999vn,Lazarus:2006ke,Marz:2006ik,Pekkurnaz:2014im,Riu:2008cx,Yang:2002td,Yang:2012cv,Zhang:2014ce}. There are three physiological isoforms of OGT, differing primarily in the number of N-terminal TPRs~\cite{Hanover:2003tf,Love:2002dr,Nolte:2002vd}. The longest isoform, with 13.5 TPRs, is referred to as nuclear and cytoplasmic OGT (ncOGT). A shorter form, with 9 TPRs and bearing a mitochondrial targeting sequence, is localized to the inner mitochondrial membrane (mOGT). The smallest of the three, sOGT, has only 2.5 TPRs and is perhaps the least studied. There have been conflicting reports as to whether mOGT has catalytic activity \textit{in vivo}~\cite{Love:2002dr,Sacoman:2017ct,Trapannone:2016hx}, and it has been speculated that mOGT and sOGT may somehow be involved in apoptosis~\cite{Fletcher:2002ft,Shin:2010fv}.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.6\columnwidth]{intro/structure-figure-1.eps}}
	\caption[Schematic of human OGT domain architecture and X-ray structure of its TPRs]{Schematic of human OGT domain architecture and X-ray structure of its TPRs. (a) A schematic representation of the domain architecture of OGT. (b) The structure of the TPR domain of OGT, spanning TPRs 1 through 11.5 (PDB: 1W3B). Chains are colored individually. The C- and N-termini are labeled, as is the dimer interface.}
	\label{fig:ogt-domain-tpr-structure}
\end{figure}
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The first structure of a segment of OGT was reported in 2004. This structure comprised 11.5 TPRs from the N-terminus of the human protein and showed that the TPRs assemble into an elongated right-handed superhelix (Figure~\ref{fig:ogt-domain-tpr-structure}b)~\cite{Jinek:2004bg}. Several features of the TPR structure are noteworthy. In the asymmetric unit, two TPR molecules pack as a homodimer, with the interface centered on TPRs 6 and 7~\cite{Jinek:2004bg}. Previous studies had suggested that OGT exists as an oligomer in solution and the TPRs were implicated in mediating its oligomerization~\cite{Haltiwanger:1992vs,Kreppel:1999vn}. The TPR crystal structure suggested that OGT may exist as a dimer, and consistent with this, mutating the residues at the TPR dimer interface afforded a lower order species. Because mutations that disrupted this interface did not substantially affect enzymatic activity \textit{in vitro}, it remains unclear what role oligomerization serves for OGT function. The TPR structure also suggested a high degree of conformational flexibility, with one monomer bent by ~40$^{\circ}$ between TPRs 9 and 10~\cite{Jinek:2004bg}. While this deformation could be due to crystal-induced unfolding, the authors suggested the conformational flexibility might have implications for substrate recognition. Finally, phylogenetic comparisons showed strong conservation in solvent-exposed residues lining the interior of the TPRs, particularly several asparagine residues near the center of the cavity, and suggested an unexpected similarity to armadillo repeat-containing proteins, such as importin $\alpha$ and $\beta$-catenin~\cite{Jinek:2004bg}. These asparagine residues are highly conserved across OGT homologs, suggesting an important role in OGT function. The similarity to proteins known to recognize multiple related ligands is striking; for example: $\beta$-catenin, which binds to various cadherins~\cite{Huber:2001vd}, PEX5, which recognizes a peroxisomal targeting signal (PTS1)~\cite{Gatto:2000df}, karyopherin (importin) $\alpha$, which recognizes a nuclear localization sequence~\cite{Conti:1998uj}, and the histocompatibility protein HLA-DR1, which recognizes pathogen-derived peptides~\cite{Stern:1994ey}, all employ asparagine-mediated peptide backbone contacts as a crucial component of their ligand binding. In these cases, there are two major reasons for this; backbone-binding interactions provide sequence-independent binding energy and predictably splay the side chains of the ligand, which in turn allows for sequence-selective recognition at defined sites along the polypeptide chain. In the context of OGT this might imply that a similar mode of substrate selection is employed for the engagement of adapter proteins or substrates.\par
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Additionally, structures of a bacterial homologue of OGT from \textit{Xanthomonas campestris}, xcOGT~\cite{Clarke:2008cl,Dorfmueller:2010kq,MartinezFleites:2008jj}, and the catalytic domain of human OGT bearing 4.5 TPRs (hOGT4.5; Figure~\ref{fig:ogt-cat-domain}) have been reported~\cite{Lazarus:2011ip}. To date, structures of hOGT4.5 have been solved with UDP, UDP-GlcNAc, the hydrolysis-resistant analog UDP-5\textit{S}GlcNAc~\cite{Gloster:2011da}, and several different peptide and glycopeptide substrates bound. Hence, binary as well as ternary substrate-like and product complexes are available~\cite{Janetzko:2016er,Jiang:2011ev,Lazarus:2011ip,Lazarus:2012gw,Lazarus:2013bq,Pathak:2015hq,Schimpl:2012ka}. A structure containing a bisubstrate analog has also been reported~\cite{Borodkin:2014it}. Taken together, these structures offer perhaps the most complete structural picture yet obtained for any Gtf. By combining the TPR structure with the catalytic domain structure, a model for human ncOGT, containing all 13.5 TPRs, can be created; in this model the TPRs form two complete superhelical turns and extend upwards from the active site.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.75\columnwidth]{intro/structure-figure-2.eps}}
	\caption[X-ray structure of the catalytic domain of human OGT]{X-ray structure of the catalytic domain of human OGT. A cartoon representation of the catalytic domain of human OGT (PDB: 3PE4). Coloring matches the domain schematic shown in Figure~\ref{fig:ogt-domain-tpr-structure}a. The N-terminal catalytic domain is in blue; the C-terminal catalytic domain is in red; the intervening domain (Int-D) is in green; the TPR domain (TPRs 9--13.5) is in gray. In addition, Helices H2 and H3 are labeled, as are UDP (magenta spheres) and the peptide substrate (CKII) (yellow ball and sticks) derived from the well-studied OGT substrate, casein kinase II~\cite{Lazarus:2011ip}.}
	\label{fig:ogt-cat-domain}
\end{figure}
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The structures of OGT have revealed many important features. As expected, the catalytic domain of OGT possesses a GT-B fold, reminiscent of that found in MurG (CAZy family GT-28), GtfB (CAZy family GT-1), and many other Gtfs \cite{Bourne:2001bf,Coutinho:2003da,Ha:2000db,Lairson:2008cn,Lombard:2013kz,Wrabl:2001ce}. GT-B Gtfs possess a catalytic domain comprising two lobes that each adopt a Rossmann-like fold~\cite{Rao:1973tq}. OGT differs from other GT-B Gtfs in several key respects, however. The N-terminal catalytic domain bears two additional helices that are crucial components of the active site~\cite{Lazarus:2011ip}. Moreover, it possesses an about 120 amino acid insertion (termed the intervening domain; Int-D) between its two catalytic lobes~\cite{Lazarus:2011ip}. This insertion forms a domain that packs exclusively against the C-terminal catalytic domain. Although its structure is known (Figure~\ref{fig:ogt-cat-domain}), its function remains mysterious. The intervening domain does however contain a large number of surface-exposed basic residues, which might suggest a role in mediating interactions with anionic macromolecules or membrane interfaces~\cite{Janetzko:2014fy,Lazarus:2011ip,Levine:2016du}. It has been suggested that OGT is recruited to membranes through binding to phosphatidylinositol (3,4,5)-triphosphate (PIP3);~\cite{Kebede:2012bc,Yang:2008js} however, mutational studies of the residues implicated in binding PIP3 are, at present, ambiguous~\cite{Kebede:2012bc,Lazarus:2011ip,Yang:2008js}, warranting further study.
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The peptide substrate binds over the UDP-GlcNAc, and the peptide binding site spans a groove formed between the two catalytic lobes and the TPR domain (Figure~\ref{fig:ogt-cat-domain}). The extended conformation of the peptide substrate likely explains the preference for proline and $\beta$-branched amino acids near the site of glycosylation~\cite{Lazarus:2011ip,Pathak:2015hq}. There is an interaction between the $\alpha$-phosphate of the nucleotide-sugar and the amide N–-H of the acceptor serine that is proposed to play a role in peptide binding to OGT~\cite{Lazarus:2011ip} and may further serve to orient the substrate for reaction (Figures~\ref{fig:intro-NT-sugar-surface} and~\ref{fig:intro-active-site})~\cite{Lazarus:2012gw}. While several OGT residues contact the backbone of the peptide substrate, few make specific contacts to the peptide side chains, consistent with the lack of a clear consensus sequence~\cite{Lazarus:2011ip,Liu:2014im,Pathak:2015hq}.
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Peptide binding is accompanied by widening of the cleft between the catalytic domain and the TPRs, and molecular dynamics (MD) simulations have suggested that the TPRs are able to pivot about a hinge region between TPRs 12 and 13, giving rise to open and closed states for the active site~\cite{Lazarus:2011ip}. Such molecular motion would be crucial for accommodating larger substrates, such as loops in folded proteins. Moreover, the flexibility is consistent with that seen in the TPR crystal structure, and with the previous suggestion that the TPRs mediate substrate binding~\cite{Jinek:2004bg}.
\par
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\section{Mechanism of Ser/Thr glycosylation by OGT} \label{sec:intro-glyco-mechanism} %DONE TO HERE
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OGT is an inverting Gtf that forms a $\beta$-glycosidic linkage between GlcNAc and the acceptor residue of the polypeptide substrate~\cite{Levine:2016du}. Although it was initially proposed that OGT follows a random bi-bi kinetic mechanism~\cite{Kreppel:1999vn}, structural information revealed that the polypeptide substrate bound above the UDP-sugar, which suggested an ordered mechanism in which UDP-GlcNAc binds first. Product inhibition studies also supported an ordered bi-bi mechanism~\cite{Lazarus:2011ip}. Hence, UDP-GlcNAc binds first to OGT, followed by the polypeptide substrate, which makes extensive contacts with the sugar donor (Figure~\ref{fig:intro-NT-sugar-surface}). Notably, kinetic studies have shown that the K\textsubscript{m,app} values for UDP-GlcNAc vary depending on the polypeptide substrate examined, ranging from $\approx1 \mu$M for Nup62 to $\approx$ 25 $\mu$M for CARM1, which underscores the importance of substrate-substrate interactions in the mechanism~\cite{Kreppel:1999vn,Lubas:2000ue,Shen:2012kv}. After the ternary substrate complex forms, the acceptor side chain attacks the anomeric carbon, with loss of UDP, to form a $\beta$-glycosidic linkage. Finally, the glycopeptide dissociates, followed by UDP.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.55\columnwidth]{intro/intro-NT-sugar-surface.eps}}
	\caption[An extensive binding surface is formed by the nucleotide sugar]{An extensive binding surface is formed by the nucleotide sugar. UDP-5\textit{S}GlcNAc (shown as magenta spheres), a UDP-GlcNAc mimic resistant to hydrolysis~\cite{Gloster:2011da}, makes extensive interactions with a CKII peptide substrate (shown as yellow sticks and surface). A hydrogen bond between the backbone N--H of the acceptor peptide and the $\alpha$-phosphate of the nucleotide sugar donor is shown as a black dotted line.}
	\label{fig:intro-NT-sugar-surface}
\end{figure}
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In addition to binding the sugar donor and acceptor substrates in close proximity (in the case of OGT, UDP-GlcNAc and a polypeptide, respectively), it has long been presumed that Gtfs must do two things in order to catalyze glycosylation: (1) activate the leaving group of the glycosyl donor towards departure, and (2) remove the proton from the hydroxyl group on the glycosyl acceptor~\cite{Lairson:2008cn}. GT-A superfamily Gtfs use a metal ion to activate the leaving group, but OGT and other GT-B Gtfs are metal ion-independent enzymes and use a different strategy (Figure~\ref{fig:intro-GTB-mech})~\cite{Lairson:2008cn}.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.85\columnwidth]{intro/intro-GTB-mech.eps}}
	\caption[A generic GT-B Gtf mechanism]{A generic GT-B Gtf mechanism. A Br{\o}nsted base (represented by ``B''), and one or more groups to stabilize the accumulation of negative charge in the leaving group (represented by ``$\delta^+$'') are thought to be required. A possible transition state, bearing oxocarbenium character, is shown.}
	\label{fig:intro-GTB-mech}
\end{figure}
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Leaving group activation in OGT is achieved through several elements that stabilize the build-up of negative charge. First, the $\beta$-phosphate of UDP-GlcNAc is anchored by hydrogen bonds to the N-terminus of an $\alpha$-helix; these include key interactions with His920, Thr921, and Thr922. Further stabilization from this helix is derived from the interactions between the negatively charged phosphate and the electric field generated by the net helix dipole~\cite{Lazarus:2012gw}. Other GT-B superfamily members use a similar mode of activation~\cite{Lairson:2008cn} with some anchoring the $\alpha$-phosphate, rather than the $\beta$-phosphate, of the leaving group~\cite{brown2013gx,Hu:2003fq}. Second, the side chain of lysine 842 is positioned directly below the $\beta$-phosphate (Figure~\ref{fig:intro-active-site}), and presumably further activates the leaving group towards departure. Biochemical data suggests that Lys842 plays an important role in OGT’s enzymatic activity with mutation to methionine or alanine largely abolishing enzymatic activity~\cite{Lazarus:2011ip,Lazarus:2012gw,MartinezFleites:2008jj,Schimpl:2012ka}. Since Lys842 is chemically reactive, as evidenced by its nucleophilicity towards a covalent OGT inhibitor~\cite{Jiang:2011ev}, its pK\textsubscript{a} may be suppressed due to its proximity to the aforementioned helix dipole; however, its protonation state remains to be established explicitly~\cite{Jiang:2011ev}. This combination of a helix dipole, carefully positioned hydrogen bonds from the N-terminus of the helix as well as a critical lysine contact to the $\beta$-phosphate of UDP-GlcNAc serves to activate the sugar donor towards nucleophilic attack. Nevertheless, glycosylation is favored over hydrolysis, raising the question of how selectivity for reaction with an acceptor hydroxyl instead of water is achieved, especially given that several crystallographically ordered water molecules are visible within the active site of all structures below 2.0 \AA~resolution (Figure~\ref{fig:intro-active-site}). It seems likely that the aforementioned contact between the $\alpha$-phosphate of UDP-GlcNAc and the amide N–-H of the acceptor serine (Figure~\ref{fig:intro-active-site}) is an important element for this selectivity. This contact may serve to bring the acceptor and donor into close proximity and position the acceptor hydroxyl in a more favorable position to form a bond with the anomeric carbon (labeled C1 in Figure~\ref{fig:intro-rxn-trajectory}) compared with any water molecule (Figure~\ref{fig:intro-active-site})~\cite{Lazarus:2012gw}. It may also further activate the leaving group for departure.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.75\columnwidth]{intro/intro-active-site.eps}}
	\caption[Active site of OGT]{Active site of OGT. A view of the OGT active site in a ternary product complex of hOGT4.5, glycosylated CKII peptide (shown as yellow sticks) and UDP (shown as cyan sticks) (PDB: 4GYY). Key residues (side chains shown as purple sticks), two ordered water molecules, labeled W1 and W2 (shown as red spheres), and hydrogen bonds (black dotted lines) are shown.}
	\label{fig:intro-active-site}
\end{figure}
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When substrate and product ternary complexes are examined, modest changes in several positions can be seen, with the C1, C2 and O5 atoms of the GlcNAc displaying the greatest displacement (Figure~\ref{fig:intro-rxn-trajectory}). A net upward rotation of the sugar moves the anomeric carbon (C1) away from the $\beta$-phosphate of UDP and into bonding distance to the acceptor hydroxyl. The inferred reaction trajectory is consistent with an electrophilic migration-type mechanism~\cite{Lazarus:2012gw}. Another notable change is the rotation of the C2-acetamide, which contacts His498 and rotates away from C1 along the reaction coordinate (Figure~\ref{fig:intro-rxn-trajectory}). The importance of the C2-acetamide was further demonstrated by the fact that OGT can transfer UDP-GalNAc, but not UDP-glucose or UDP-2-ketoGlc, a UDP-GlcNAc analog in which the C2 ``N--H'' is replaced by a ``CH\textsubscript{2}''~\cite{Lazarus:2012gw}. Similar acetamide to $\beta$-phosphate interactions have been observed for other glycosyltransferases~\cite{LiraNavarrete:2014fz,Tvaroska:2012hs}.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.75\columnwidth]{intro/intro-rxn-trajectory.eps}}
	\caption[Reaction trajectory for Ser/Thr glycosylation by OGT]{Reaction trajectory for Ser/Thr glycosylation by OGT. An overlay of: (1) A product ternary complex (PDB: 4GYW) of OGT, a glycosylated CKII peptide (gCKII), and UDP (both ligands shown in yellow sticks), and (2) A substrate ternary complex (PDB: 4GYY) of OGT, a CKII peptide and UDP-5\textit{S}GlcNAc (both ligands shown in cyan sticks). Several positions on the NT-sugar(s) are labeled.}
	\label{fig:intro-rxn-trajectory}
\end{figure}
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In addition to providing information about leaving group stabilization and reaction trajectory, the OGT ternary structures revealed a surprise: there is no OGT side chain that can serve as a catalytic base in the immediate vicinity of the reactants~\cite{Lazarus:2012gw,Schimpl:2012ka,Withers:2012iq}. Two different side chains, His498 and His558, had previously been suggested as candidates for the catalytic base, but both were ruled out after the examination of ternary structures containing substrate analogs implicated them in other roles~\cite{Lazarus:2012gw,Schimpl:2012ka}. Two alternatives for how proton transport is achieved have been suggested (Figure~\ref{fig:intro-catalytic-base}). In one, the $\alpha$-phosphate of UDP is proposed to act as a base to deprotonate the acceptor hydroxyl (Figure~\ref{fig:intro-catalytic-base}, proposal I)~\cite{Schimpl:2012ka}.  Nuclear magnetic resonance studies showed a pK\textsubscript{a} of 6.4 for the pyrophosphoryl group of UDP-GlcNAc, making it a plausible base~\cite{Jancan:2012ka,Levine:2016du,Schimpl:2012ka}. Further support stemmed from evidence suggesting that retaining Gtfs that proceed via an S\textsubscript{N}i-like mechanism~\cite{Ardevol:2011bh,Errey:2010cl,Gomez:2012bd,Lee:2011fd} may similarly employ a phosphate as the catalytic base~\cite{Levine:2016du}. In the other, a chain of water molecules leading to an aspartate side chain, Asp554, is proposed to facilitate transport of the proton liberated during the reaction from the active site (Figure~\ref{fig:intro-catalytic-base}, proposal II)~\cite{Lazarus:2012gw}. In order to test the first hypothesis, diastereomeric phosphodithioate analogs of UDP-GlcNAc were prepared and it was found that while both had a similar binding affinity for OGT, only one of them functioned as a donor substrate~\cite{Schimpl:2012ka}. While this finding suggests that the $\alpha$-phosphate is involved in the catalytic mechanism, results involving substrate mimics must be examined cautiously. Replacing the acceptor serine with an aminoalanine may, depending on the protonation state of the amine, bring the amino group in closer proximity to the $\alpha$-phosphate than it would be in the native substrate. Furthermore, oxygen to sulfur substitution on the phosphate would likely affect the previously mentioned hydrogen bond to the backbone of the acceptor amide (Figure~\ref{fig:intro-active-site}). Given that the importance of this interaction has not been rigorously examined, its energetic contribution to binding should be quantified. The proposal that the proton is shuttled out of the active site by a Grotthuss-like mechanism~\cite{RN1229,RN1230} invokes the active site ordered water molecules in the mechanism in lieu of implicating a catalytic base within OGT. In a transition state involving an oxocarbenium ion-like species, as computational studies have suggested for OGT~\cite{Tvaroska:2012hs}, a general base may not be needed to deprotonate the acceptor hydroxyl for the reaction to occur. In this case the ordered water molecules leading to residue Asp554 may simply provide a pathway for proton extrusion from the active site. It has been reported that mutation of Asp554 to alanine\footnote{The primary sequence numbering of OGT used in reference \cite{Lazarus:2005bm} is based on mOGT. As a result, Asp554 is discussed as Asp438.} abolishes catalytic activity~\cite{Lazarus:2005bm}, but other studies have found that while mutation to asparagine impairs glycosylation, it does not abolish it~\cite{Janetzko:2016er,Schimpl:2012ka}. A computational study of these pathways suggested that the lowest energy path for catalysis employed the $\alpha$-phosphate as the base;~\cite{Kumari:2015jh}. However, further study is required to resolve the issue of how the proton on the acceptor hydroxyl is ultimately removed from the active site.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.8\columnwidth]{intro/intro-base-proposals.eps}}
	\caption[Two proposals for the operative proton transport mechanism in OGT]{Two proposals for the operative proton transport mechanism in OGT. Proposal I involves the $\alpha$-phosphate acting to deprotonate the glycosyl acceptor; Proposal II involves a Grotthuss-like mechanism wherein the proton is shuttled out of the active site through a chain of ordered water molecules.}
	\label{fig:intro-catalytic-base}
\end{figure}
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%\begin{figure}[b]
%	\caption[Mechanism of ser/thr glycosylation by OGT]{\textit{(On the following page)} Mechanism of ser/thr glycosylation by OGT. (a) UDP-5\textit{S}GlcNAc (magenta spheres), a UDP-GlcNAc mimic resistant to hydrolysis~\cite{Gloster:2011da}, makes extensive interactions with a peptide substrate (yellow sticks and surface). A hydrogen bond is shown as a black dotted line. (b) A generic GT-B Gtf mechanism. A Br{\o}nsted base (represented by “B”), and one or more groups to stabilize the accumulation of negative charge in the leaving group (represented by ``$\delta^+$'') are thought to be required. A possible transition state, bearing oxocarbenium character, is shown. (c) A view of the OGT active site in a ternary product complex of hOGT4.5, glycoCKII (yellow sticks) and UDP (cyan sticks) (PDB: 4GYY). Key residues (purple sticks), two ordered water molecules, W1 and W2 (red spheres), and hydrogen bonds (black dotted lines) are shown. (d) An overlay of a product complex, containing a glycosylated CKII peptide and UDP (both as yellow sticks), and a substrate complex, containing CKII and UDP-5\textit{S}GlcNAc (both as cyan sticks). Several positions on the sugars are labeled. (e) Two main proposals for the operative proton transport mechanism in OGT.}
%	\label{fig:glycosylation-mechanism-ogt}
%\end{figure}
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%\begin{figure}[p]
%	\centerline{\includegraphics[width=1\columnwidth]{intro/Review_fig3_edit2.eps}}
%\end{figure}
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\section{OGT is required for Host cell factor-1 proteolysis}
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In 2011 OGT was implicated as the cellular factor responsible for the maturation of an essential transcriptional co-activator, host-cell factor 1 (HCF-1)~\cite{Daou:2011bn,Capotosti:2011kd}. HCF-1 was first identified in the late 1980s as a component from HeLa cell lysate required for the stable formation of the \textit{Herpes simplex} VP16-induced complex (VIC) and the expression of immediate-early genes during herpes viral infection \cite{apRhys:1989ub,Gerster:1988vs,Katan:1990iy,Kristie:1990iu,Stern:1991ki,Vogel:2013ef,Wysocka:2003jz,Xiao:1990kp}. HCF-1 has since been implicated as a critical element in cell-cycle regulation, including both G\textsubscript{1}-to-S and M-phase progression \cite{Julien:2004cx,Luciano:2003hb,Mangone:2010cr,Michaud:2013fo,Tyagi:2007cf,Vogel:2000ds,Wysocka:2003cp,Zhou:2013kc}.
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Biochemically, the first report that described the purification of HCF-1 (then called C1), by Kristie and Sharpe~\cite{Kristie:1993tm}, suggested its existence as a set of heterogeneous, but related, polypeptide fragments; later work revealed that these polypeptide fragments were derived from a large precursor protein (\textgreater200 kDa) by some proteolytic maturation process~\cite{Wilson:1993wf}. Though the protease responsible for this cleavage remained elusive, pulse-chase studies suggested that cleavage took place largely in the nucleus, but also, albeit to a lesser extent, in the cytoplasm~\cite{Wilson:1995ec}.\par
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\begin{figure}[ht]
	\centerline{\includegraphics[width=0.8\columnwidth]{intro/hcf-schematic.eps}}
	\caption[Schematic of HCF-1 domain structure]{Schematic of HCF-1 domain structure. HCF-1 is translated as a 2035 amino acid polypeptide comprising: an N-terminal Kelch domain~\cite{Gupta:2014kv}, followed by a region rich in basic amino acids (colored in green); the middle portion of the protein bears six 26-amino acid repeating sequences (colored in yellow); a C-terminal portion with an acid domain and a fibronectin type III (Fn3) repeat domain~\cite{Wilson:2000ut,Park:2012cu} (colored in orange). The primary sequence of the second proteolytic repeat is shown, and the glutamic acid at the site of cleavage is shown in red. The cleavage site, identified by Edman degradation is denoted by the ``\textsuperscript{$\bigtriangledown$}'' symbol, between Glu10 and Thr11.}
	\label{fig:hcf-intro-schematic}
\end{figure}
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Cleavage appeared to take place at one or more of six 26-amino acid repeating elements, which constituted an atypically large protease recognition motif (See Figure~\ref{fig:hcf-intro-schematic})~\cite{Wilson:1993wf,Wilson:1995ec}. Based on N-terminal peptide sequencing, cleavage was reported to occur between Glu10 and Thr11 of a given repeat~\cite{Kristie:1995vy,Wilson:1995ec}. One striking feature of this cleavage was the fact that even though it was shown to be required for proper HCF-1 function~\cite{Julien:2003in}, the two halves of the protein appeared to remain non-covalently associated post-processing~\cite{Park:2012cu,Wilson:1995ec,Wilson:2000ut}. Expression of only the N-terminal domain of HCF-1 or of an HCF-1 variant incapable of being cleaved led to defects in cytokinesis and caused binucleation of cells~\cite{Capotosti:2011kd,Julien:2003in}. Binucleation was also observed when an HCF-1 variant lacking the proteolytic repeats, but bearing an ectopic protease site, was expressed in place of the endogenous copy~\cite{Capotosti:2011kd}. Though several mechanisms for cleavage were considered~\cite{Daou:2011bn,Capotosti:2011kd,Vogel:2000ju}, the finding that HCF-1 interacts stably with, and is modified by, OGT~\cite{Capotosti:2011kd,Khidekel:2004da,Mazars:2010kp,Wysocka:2003cp} led to the suggestion that OGT may somehow promote cleavage. Of the two groups that implicated OGT in HCF-1 cleavage, the group of Winship Herr demonstrated that recombinant OGT and UDP-GlcNAc were necessary and sufficient for this process \textit{in vitro}~\cite{Capotosti:2011kd}. Alanine scanning of the repeat suggested that several of the threonine residues in the C-terminal region of the repeat were functionally important for cleavage. This led to a division of the proteolytic repeat into a ``cleavage region'' and a ``threonine region'' (as shown in Figure~\ref{fig:hcf-intro-schematic}), with further experiments suggesting that OGT may interact with the HCF-1 repeat in a bipartite fashion~\cite{Capotosti:2011kd}. Through a series of experiments, Capotosti \textit{et al.} demonstrated that the glycosylation activity of OGT was required for HCF-1 cleavage. However, the questions of whether glycosylation by OGT could facilitate autoproteolysis, or whether Ser/Thr glycosylation of the N-terminal domain of HCF-1 was a requirement for cleavage remained unanswered~\cite{Capotosti:2011kd}.
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\section{Conclusions and focus of this dissertation}
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Though our mechanistic understanding of OGT has advanced tremendously since 1984, as has the available repertoire of chemical tools to study \textit{O}-GlcNAcylation~\cite{Worth:2017ie}, the discovery that OGT is responsible for the cleavage of HCF-1 has led to an entirely new set of questions: How might a glycosyltransferase effect amide bond cleavage? Is a second OGT active site responsible for this new transformation? What is the mechanism of amide bond cleavage? And why would Nature connect OGT activity with the cleavage of HCF-1? (see also reference~\cite{Hanover:2011jm}) In the chapters to follow, I aim to answer these questions.
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%\section{Protein aging: isoaspartate formation in \textit{in vitro} and \textit{in vivo} systems}\label{intro:isoasp}
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\section{Resource: High-resolution mass spectrometry}\label{intro:MS}
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High resolution mass spectrometry (MS) is a powerful analytic tool that exploits the intrinsic property of molecular mass to distinguish species \cite{Zhurov:2013cg}. However, a great deal of information can be garnered beyond simply the mass of a molecule. Using tandem mass spectrometry (MS\textsuperscript{n}), one can deduce structural characteristics of the molecules being examined. At its most fundamental level, MS operates by taking gaseous ions and analyzing them based on the ratio of their mass to their charge (\textit{m/z}). Three different types of mass spectrometer were used in these studies: a time-of-flight (ToF) mass spectrometer \cite{Guilhaus:1995ff}, an Orbitrap mass spectrometer \cite{Hu:2005fy,Scigelova:2011dt}, and a Fourier transform ion cyclotron resonance (FT-ICR) mass spectrometer \cite{Marshall:2002kn,Scigelova:2011dt}. The details of how gas-phase ions are produced (electrospray ionization (ESI) \cite{FENN:1989ui}, in this case) and how each of these mass spectrometers manipulate ions are beyond the scope of this thesis and will not be discussed here. Rather, since aspects of the work described in Chapters~\ref{ch:mechanism} and~\ref{ch:aspartate} rely on tandem mass spectrometry experiments, a discussion of MS\textsuperscript{n} experiments for structure elucidation follows.\par
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Today, peptide sequencing, as well as PTM determination, is routinely done by MS/MS \cite{Steen:2004eu}. The most common fragmentation methods, collision-induced dissociation (CID), and higher-energy collision-induced dissociation (HCD), effect fragmentation through vibrational excitation of the analyte ions \cite{Wells:2005ga,Zhurov:2013cg}. Excitation occurs via repeated collisions between the analyte ion and neutral gas molecules (Figure~\ref{fig:hcf-intro-mass-CID}a). In the case of HCD, this process takes on the order of microseconds, allowing accumulated kinetic energy to be statistically redistributed across all degrees of freedom for the analyte ion \cite{Wells:2005ga}. As a result, the most labile linkage within this ion are subject to rupture first. In the case of a typical polypeptide, the weakest bond is the C--N (peptide) bond \cite{Wells:2005ga,Zubarev:2003ds}, and scission results in the formation of \textit{b} and \textit{y} fragment ions (Figure~\ref{fig:hcf-intro-mass-CID}b). However, if the peptide bears a PTM wherein a more labile linkage than an amide bond is present, e.g., phosphorylation or glycosylation, that linkage becomes preferentially subject to fragmentation. In these cases, loss of the PTM prior to the generation of \textit{b} and \textit{y} ions results in a loss of positional information, with respect to that PTM, making vibrational excitation ineffective for PTM analysis. Fortunately, alternative fragmentation approaches, involving the use of low energy electrons are able to retain sequence information for labile PTMs.\par
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\begin{figure}
	\centerline{\includegraphics[width=0.9\columnwidth]{intro/mass-spectrometry-1.eps}}
	\caption[Collision-induced MS/MS]{Collision-induced MS/MS. (a) A schematic showing the activation mode of CID/HCD. N\textsubscript{2} is used as a sample inert gas. (b) Representative structures for fragment ions produced by CID/HCD from a generic peptide (tripeptide when n=1). Formation of the oxazolone \textit{b} ion structure is believed to occur concurrent with cleavage of the C--N bond (and departure of the \textit{y} ion) \cite{Chen:2005fk}.}
	\label{fig:hcf-intro-mass-CID}
\end{figure}
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Electron-capture dissociation (ECD) and electron-transfer dissociation (ETD) are fragmentation methods that exploit ion-electron interactions to generate fragment species from the parent analyte ion \cite{Zhurov:2013cg,Zubarev:2003ds,Zubarev:2004if}. The process of electron-capture-based dissociation involves the interaction of a multiply charged cation with an electron \cite{Zubarev:2003ds}. When an electron approaches a cation, there is a mutual Coulombic attraction; however, for this attraction to lead to capture the cation must dissipate the excess energy over a very short period of time ($10^{-14}$ seconds) \cite{Zubarev:2003ds} (Figure \ref{fig:hcf-intro-mass-ECD}a). This time-scale is too rapid for atomic motions to compensate and redistribute the energy across several degrees of freedom. For polyatomic cations a solution is a prompt dissociation event, which can allow for stabilization of the electron within time-scales on the order of $10^{-16}$--$10^{-15}$ s~\cite{Zubarev:2003ds}. In this scenario, electron capture and bond cleavage appear to occur as a concerted process, known as dissociative recombination \cite{Zubarev:2003ds}. Further, bond cleavage under these conditions is mostly an electronic process, proceeding faster than atomic vibrations and rearrangements. Processes wherein bond dissociation occurs faster than energy redistribution are known as non-ergodic processes \cite{Zubarev:2003ds}; however, whether ECD is a non-ergodic process has been extensively debated \cite{Turecek:2003cf,Zubarev:1998cz,Zubarev:2003ds}.\par
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There are several practical considerations for ECD/ETD worth discussing here: A consequence of electron capture is a net charge reduction of the parent ion which practically means that analyte peptides must be multiply charged in order to allow for detection. In ECD parent ions are bombarded directly with low energy electrons ($\approx$ 1--10 eV) under high vacuum ($10^{-10}$ torr) \cite{Zhurov:2013cg}. As a result, a major limitation of ECD is that it is most commonly implemented on FT-ICR MS, whereas ETD is amenable to implementation on Orbitrap instruments, making it more readily accessible \cite{Zhurov:2013cg}. To circumvent the need for a direct electron source, ETD employs a ``reagent'' gas to facilitate the electron transfer process; the radical anion is produced separately using a chemical ionization source and allowed to react with analyte ions for a defined period of time \cite{Zhurov:2013cg}. As a result, the ion-ion interactions involved in ETD occur in an ion trap, which is held at higher pressure (by several orders of magnitude). This leads to a faster collisional cooling and reduces the internal energy of the post-electron attachment complex, reducing the likelihood of more energetic fragmentation pathways occurring. In this regard ECD imparts greater energy into an ion than ETD, often leading to additional peaks in an ECD spectrum, as compared to an ETD spectrum \cite{Zhurov:2013cg}.\par
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The result of ECD (or ETD) fragmentation is not the formation of \textit{b} and \textit{y} ions, but instead $c'$ and $z^\bullet$ fragments (Figure~\ref{fig:hcf-intro-mass-ECD}b, ``$'$'' is pronounced ``prime'' and ``$^\bullet$'' is pronounced ``dot'') (Figure \ref{fig:hcf-intro-mass-ECD}b). The prime symbol denotes the fragment to which the H$^\bullet$ is transfered \cite{Zubarev:2004if}. In fact, an ECD spectrum is largely devoid of \textit{b} and \textit{y} ions. This has been attributed to several factors: First, peptide bond fragmentation proceeds by intermediates that require (relatively) long life times (up to microseconds), which are incompatible with the rate at which ECD is believed to occur (($10^{-12}$ s, or faster) \cite{Zubarev:2003ds}. As a result, only direct bond scission can occur in ECD. Second, the species formed in ECD are not even-electron species, as is the case for vibrational excitation (CID/HCD), but instead are hydrogen-abundant radical cations. Since the presence of the radical changes the proximal bond strengths, this has been cited as a potential explanation for the observed N--C$\alpha$ bond cleavage \cite{Zubarev:1999js,Zubarev:2003ds}. The preference for N--C$\alpha$ bond cleavage has been rationalized through relative electron affinities \cite{Zubarev:2003ds}. The protonated side chains of lysine, arginine and histidine can be solvated by the amide backbone in the form of a species: A$\cdots$H\textsuperscript{+}$\cdots$B \cite{Zubarev:1998cz}. An electron captured by such a species would form $^\bullet$A--H + B if the electron affinity of A\textsuperscript{+}H is greater than that of HB\textsuperscript{+} \cite{Zubarev:1998cz}. The electron affinity of a protonated nitrogen, such as an amide or amine is on the order of 4 eV, compared to 6 eV for a protonated carbonyl \cite{Zubarev:1998cz}; this also suggests that neutralization of the protonated carbonyl by electron capture would be an exothermic process. Though multiple mechanisms for bond scission have been described \cite{Zhurov:2013cg}, a representative example, termed the ``Cornell mechanism'' is shown in Figure \ref{fig:hcf-intro-mass-ECD}b.\par
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%
\begin{figure}
	\centerline{\includegraphics[width=0.9\columnwidth]{intro/mass-spectrometry-2.eps}}
	\caption[Electron-capture dissociation MS/MS]{Electron-capture dissociation MS/MS. (a) A schematic showing the activation mode of ECD/ETD. (b) Representative structures for $c'$ and $z^\bullet$ fragment ions produced by ECD/ETD from a generic peptide (tripeptide when n=1). (c) The ``Cornell mechanism'' for N--C$\alpha$ bond cleavage in ECD/ETD \cite{Zhurov:2013cg}. The pendant protonated amine at the N-terminus represents a protonated residue located generically N-terminal to the site of bond scission. The product fragments shown represent the corresponding $c'$ and $z^\bullet$ species.}
	\label{fig:hcf-intro-mass-ECD}
\end{figure}
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Unlike C--N (peptide) fragmentation, where all amino acids are able to lead to fragmentation, tertiary amides are not susceptible to ECD fragmentation. Scission of the N--C$\alpha$ bond for proline fails to produce two polypeptide fragments, so $c'$/$z^\bullet$ ion pairs are not expected at these positions \cite{Zhurov:2013cg,Zubarev:2003ds}.
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In addition to the even-electron $c'$ and radical $z^\bullet$ ion pairs, it is possible to also observe radical $c^\bullet$ and even-electron $z'$ ion pairs \cite{Savitski:2007jc}. This has been suggested to occur via hydrogen atom transfer. After N--C$\alpha$ bond cleavage of $[\mathrm{M}+n\mathrm{H}]^{n+}$, the $[c+z]^{(n-1)+\cdot}$ complex remains briefly held together by non-covalent interactions. If the lifetime of this complex is sufficiently long, the $z^\bullet$ radical can abstract a hydrogen atom from $c'$ forming $z'$ (where $z'=z^\bullet+\mathrm{H}$; a z+1 fragment ion) and $c^\bullet$ (where $c^\bullet=c'-\mathrm{H}$; a c--1 fragment ion) \cite{Savitski:2007jc,Zhurov:2013cg}.\par
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