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Development of metabolic chemical reporters for the investigation of protein glycosylation

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Content DEVELOPMENT OF METABOLIC CHEMICAL REPORTERS FOR THE
INVESTIGATION OF PROTEIN GLYCOSYLATION
by
Kelly Nicole Chuh
A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMISTRY)
December 2017
Copyright 2017 Kelly Nicole Chuh
Acknowledgements
To my family, for your endless support and patience. There are hardly words to describe
how much you mean to me. You raised me to become a daughter, sister, wife and scientist
that I am proud of. Thank you for giving me the freedom and pressure-free environment
to pursue science at an early age and for teaching me work ethic and dedication and to
always do my best. Finishing my PhD wouldn’t have been possible without you all.
To my husband and fellow PhD, Steve. We have come along way from new first year
graduate students at USC to husband and wife. You helped me survive through the worst
times and were always there to help me celebrate the best ones. You are my strongest
support system, my touch stone and my partner for life. I am lucky to call myself your
wife and I am looking forward to our next adventure in Princeton.
To my lab family. Specifically the alumni that supported me during my infancy in
graduate school. To Balyn who took me under her wing when I first joined. What started
as a mentor relationship has evolved into one of my most special and real friendships that
will exist far past the limits of science. To Tharindumala, for the comfort of your hugs
and words of wisdom and for always helping me to see the bigger picture.
To Matt, thank you for having faith in me as a graduate student and for continuing to
teach me how to be the best scientist I can be. Thank you for motivating me to keep
pushing and teaching me the skills to overcome my frustrations and insecurities. For
drinking beers with me at the end of hard day or whiskey after the the most impossible
ones. You have inspired me to be a confident and strong woman in science with a tough
skin. You are a great mentor and true friend.
To current lab member Anna, whose work was absolutely integral for the publication of
my favorite project and whose friendship was crucial for my sanity during that time. To
Narek, thank you for all of the walks to Starbucks. To all of the current graduate students
and post docs in the Pratt lab, thank you for supporting me through post doc interviews
and for bearing with me through my last months.
ii
Table of Contents
Acknowledgements
List of Figures
List of Schemes
Abstract
Chapter 1. Chemical Methods for Encoding and Decoding of
Posttranslational Modifications
Introduction
Conclusions and Future Outlook
Chapter One References
Chapter 2. Changes in Metabolic Chemical Reporter Structure
Yield a Selective Probe of O-GlcNAc Modification
Introduction
Results
Discussion and Conclusion
Materials and Methods
Chapter Two References
Chapter 3. A New Chemical Reporter of O-GlcNAc Reveals
Modification of The Apoptotic Caspases That Blocks
The Cleavage and Activation of Caspase-8
Introduction
Results
Discussion
Materials and Methods
Chapter Three References
Chapter 4. N-Propargyloxycarbamate Monosaccharides As
Metabolic Chemical Reporters of Carbohydrate Salvage
Pathways and Protein Glycosylation
Introduction
Results
Discussion and Conclusion
Materials and Methods
Chapter Four References
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Chapter 5. A Chemical Reporter for Visualizing Metabolic
Cross-Talk Between Carbohydrate Metabolism and
Protein Modification
Introduction
Results
Discussion
Conclusion
Materials and Methods
Chapter Five References
Chapter 6. The Small Molecule 2-Azido-2-Deoxy-Glucose is a
Metabolic Chemical Reporter of O-GlcNAc
Modifications in Mammalian Cells, Revealing an
Unexpected Promiscuity of O-GlcNAc Transferase
Introduction
Results
Discussion
Materials and Methods
Chapter Six References
References
Appendices
Appendix A: NMR Spectra
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List of Figures
Figure 1-1: Chemical methods to tag and enrich posttranslationally
modified proteins.
Figure 1-2: Bioorthogonal reactions occur selectively between two
abiotic functional groups.
Figure 1-3: Encoding protein phosphorylation.
Figure 1-4: Decoding protein phosphorylation.
Figure 1-5: Protein glycosylation.
Figure 1-6: Ubiquintination.
Figure 1-7: Lipidation.
Figure 1-8: Acetylation.
Figure 1-9: Methylation and ADP-ribosylation
Figure 2-1: Metabolic chemical reporters.
Figure 2-2: 6AzGlcNAc labels proteins in living cells.
Figure 2-3: The GalNAc salvage pathway.
Figure 2-4: Investigation of 6AzGlcNAc metabolism.
Figure 2-5: LC-MS analysis of 6AzGlcNAc-1-phosphate
production by AGM1.
Figure 2-6: Fluorescence incorporation of MCRs in a variety of cell
lines.
Figure 2-7: Characterization of 6AzGlcAc.
Figure 2-8: Glycoprotein specificity of 6AzGlcNAc.
Figure 2-9: Identification of O-GlcNAcylated proteins using
6AzGlcNAc.
Figure 3-1: O-GlcNAcylation and the major apoptotic caspases.
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v
Figure 3-2: Characterization of the metabolic chemical reporter
6AlkGlcNAc
Figure 3-3: 6AlkGlcNAc can label proteins in a range of
mammalian cell lines.
Figure 3-4: 6AlkGlcNAc displays improved labeling compared to
6AzGlcNAc.
Figure 3-5: Global 6AlkGlcNAc labeling is less efficient compared
to GlcNAlk.
Figure 3-6: 6AlkGlcNAc is selective for O-GlcNAcylation.
Figure 3-7: Treatment with 6AlkGlcNAc is not toxic to mammalian
cells.
Figure 3-8: Identification of O-GlcNAcylation proteins, including
caspase-3 and -8, using 6AlkGlcNAc.
Figure 3-9: Chemoenzymatic labeling of O-GlcNAc modifications.
Figure 3-10: Caspases-3 and -8 are genuine O-GlcNAcylated
proteins in mouse cells.
Figure 3-11: The apoptotic caspases are O-GlcNAcylated in human
cancer cell lines.
Figure 3-12: O-GlcNAcylation levels affect caspase-8
cleavage/activation.
Figure 3-13: Caspase-8 is O-GlcNAcylated near its
cleavage/activation sites.
Figure 3-14: Generation of caspase-8 null HeLa cells
Figure 3-15: Characterization of caspase-8 dimerization.
Figure 3-16: O-GlcNAcylation of the caspase-8 cleavage sites
affects its cleavage/activation.
Figure 4-1: N-Propargylcarbamate-containing metabolic chemical
reporters incorporated onto proteins.
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Figure 4-2: Characterization of N-propargyloxycarbamate (Poc)
bearing metabolic chemical reporters.
Figure 4-3: Flow cytometry analysis of metabolic chemical reporter
incorporation.
Figure 4-4: Incorporation of metabolic chemical reporters into the
O-GlcNAcylation pathway.
Figure 5-1: Using metabolic chemical reporters (MCRs) to detect
cellular metabolism.
Figure 5-2: Characterization of proteins that are labeled by the
MCR 1-deoxy-GlcNAlk.
Figure 5-3: Characterization of 1-deoxy-GlcNAlk labeling.
Figure 5-4: Dose-dependence and dynamics of 1-deoxy-GlcNAlk
protein labeling.
Figure 5-5: Generality of 1-deoxy-GlcNAlk labeling.
Figure 5-6: Toxicity of 1-deoxy-GlcNAlk.
Figure 5-7: Identification of posttranslationally modified proteins
using 1-deoxy-GlcNAlk.
Figure 5-8 Identification of proteins labelled by GlcNAlk.
Figure 6-1: Evaluation of peracetylated-2-azido-glucose as a
metabolic chemical reporter (MCR).
Figure 6-2: 2AzGlcNAc as an O-GlcNAc metabolic chemical
reporter.
Figure 6-3: 2AzGlcNAc is a general chemical reporter in a variety
of mammalian cell lines.
Figure 6-4: Characterization of the dynamics of 2AzGlcNAc
labeling.
Figure 6-5 The acetylation pattern and labeling by 2AzGlcNAc is
responsible for cellular toxicity
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List of Schemes
Scheme 2-1: Synthesis of Ac 36AzGlcNAc.
Scheme 2-2: Synthesis of Alkyne-azo-biotin.
Scheme 2-3: Synthesis of 6AzGlcNAc-1-phosphate.
Scheme 3-1: Synthesis of Ac 36AlkGlcNAc.
Scheme 5-1: Synthesis of 1-deoxyGlcNAc, Ac3-1-deoxyGlcNAlk,
and 1-deoxy-GlcNAlk.
Scheme 6-1: Synthesis of Ac 32AzGlcNAc.
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Abstract
Post translational modifications (PTMs) are covalent additions appended to the side
chains of amino acids that afford an extra level of biological and chemical complexity to
the genome. These modifications can range from the addition of a single methyl group to
the installation of a large oligosaccharide. Nevertheless, PTMs have been shown to have
the ability to change the function, activity or localization of their target proteins.
Furthermore, PTMs can be dynamically regulated and can be added enzymatically or
chemically thus allowing them to act as signaling mechanisms or as metabolic sensors. In
order to interrogate the consequences of PTMs, many methods have been developed to
visualize and identify substrate proteins. One of these methods involves the treatment of
live cells with a small molecule memetic of a metabolic precursor, containing an abiotic
azide or alkyne, for its enzymatic incorporation onto target proteins and subsequent
analysis using copper catalyzed azide alkyne cycloaddition (CuAAC), or click chemistry.
These small molecules are termed metabolic chemical reporters (MCRs) and have been
extensively utilized in the study of protein glycosylation. Described here is the
development and optimization of the first MCRs to be specific for a single type of protein
glycosylation called O-GlcNAc. O-GlcNAc modification is defined by the enzymatic
addition of a single monosaccharide termed N-acetyl-glucosamine (GlcNAc) to serine
and threonine residues of intracellular proteins. With this novel MCR in hand, the
modification of the apoptotic regulator proteins called the caspases was revealed and was
subsequently shown to affect the cleavage and activation of the protein during cell death.
In addition, described here are three N-propargyloxycarbamate MCRs for the
ix
investigation of carbohydrate salvage pathways, an MCR for visualizing the metabolic
cross-talk between carbohydrate metabolism and protein acetylation, and finally the small
molecule 2-azido-2-deoxy-glucose, a reporter that revealed the unexpected promiscuity
of O-GlcNAc transferase, the enzyme responsible for the addition of O-GlcNAc.
x
Chapter 1. Chemical Methods for Encoding and Decoding of
Posttranslational Modifications
Introduction
In the post-genomic era, it has become clear that the complexity of life cannot be
explained by the number of genes in the genome alone. One layer of added functional and
structural diversification beyond the genome is afforded via posttranslational
modifications (PTMs). PTMs are covalent additions introduced to amino acid side chains
or termini of proteins, either enzymatically or chemically, and represent one of the basic
mechanisms to increase the chemical and biological diversity of the genome. These
modifications range from the simple addition of a phosphate to the incorporation of large
oligosaccharide structures, and they have been shown to change the biochemical and
biophysical properties of the substrate protein. In addition to regulating activity,
localization, and interactions with other proteins, PTMs can also carry information about
the cellular environment (e.g., normal or disease state) or biochemical changes in
response to various stimuli. PTMs can be dynamic in nature, and in many cases, cells are
equipped with enzymatic machinery with opposing activities to install and remove the
modification when given a functionally relevant cue. Despite the documented importance
of PTMs in cellular biology, their identification and the study of specifically-modified
substrate proteins remain challenging. Although proteins can be harvested from cells for
study, this process often requires tedious and often difficult separation of their modified
1
and unmodified forms. Furthermore, PTMs can occur on several sites simultaneously and
substoichiometricly, making the isolation of a completely homogenous population
extremely difficult. Therefore, access to site-specifically modified proteins is of the
utmost importance for the study of PTMs. Additionally, identifying all proteins within the
proteome that are substrates for a specific PTM continues to be a challenge despite being
critical for understanding the biological pathways that control and are regulated by a
given PTM. Unfortunately, some of the traditional tools for performing these types of
analysis (e.g., antibodies) are not available for all PTMs and cannot a priori distinguish
enzyme-specific modification events. Over the years, many different approaches for
studying PTMs have emerged, including the development of selective and unique
chemical methods for the synthesis, identification, and analysis of posttranslationally
modified proteins. In general, these techniques can be broadly classified into two
categories, which both enable the installation of visualization or affinity tags:
1) Methods that use chemoselective reactions to exploit the unique chemical-reactivity of
the posttranslational modifications themselves (Figure 1-1A).
2) Strategies to install abiotic probes (chemical reporters, Figure 1-1B) that can be
selectively subjected to a range of bioorthogonal reactions (Figure 1-2).
2
Figure 1-1. Chemical Methods to tag and enrich posttranslationally modified proteins. (A)
Some posttranslational modifications (PTMs) can be specifically reacted with enrichment tags
using chemoselective reactions. (B) Bioorthogonal reactions in combination with chemical
reporters enable the installation of affinity tags. Chemical reporters can either be incorporated into
PTMs using cellular metabolism or appended to existing PTMs using enzymes or selective
chemical reactions.
In parallel, the development of more sensitive mass spectrometers and increasingly
sophisticated computer-algorithms for peptide identification have enabled the routine
proteomic identification of thousands of proteins from individual complex-samples.
Coupled with complementary ionization chemistries and quantitative methods, mass
spectrometry can offer high sensitivity, modification-site identification and the ability to
quantify changes in PTM occupancy.
This chapter details the methods that have been developed to “encode” and “decode” the
posttranslational modification of glycosylation (Figure 1-1), where “encode” relates to
3
the chemical synthesis or semisynthesis of homogeneously modified glyco-proteins or
peptides, and “decode” defines the methods that are utilized for the isolation and
identification of substrate proteins. Additionally, this chapter will highlight the currently
available chemical-strategies to enrich and identify posttranslationally-modified proteins
in mammalian cells and highlight their recent applications.
Figure 1-2. Bioorthogonal reactions occur selectively between two abiotic functional-groups.
(A) The Staudinger ligation allows for the coupling of azides with triarylphosphine reagents. (B)
The Cu(I)-catalyzed azide-alkyne cycloaddition (CuAAC) gives a stable triazole product from
azides and alkynes. This common bioorthogonal reaction can be performed in both directions;
however, azide-tags give reduced background signals. (C) Cyclooctyne reagents will react with
azides in a strain-promoted azide-alkyne cycloaddition (SPAAC). (D) Tetrazines will undergo
rapid inverse-demand Diels-Alder reactions with activated alkyenes, such as cyclopropenes.
Phosphorylation
Protein phosphorylation is the transfer of an inorganic phosphate group to a variety of
amino acid side-chains, including most commonly to the hydroxyl groups of serine,
threonine, and tyrosine residues (Figure 1-3A). The modification is installed by members
4
of the kinase family of enzymes, which transfer the high-energy gamma phosphate from
adenosine triphosphate (ATP) to the substrate residues. Phosphorylation can be
subsequently removed by phosphatase enzymes, rendering the modification dynamic. The
first protein kinase, protein kinase A, was discovered in 1981 as the enzyme that could
phosphorylate and subsequently activate the metabolic enzyme phosphorylase (Hayes
and Mayer, 1981). This discovery would be just the tip of the iceberg, as protein
phosphorylation networks have since been identified that control essentially all biological
processes, including metabolic regulation, cellular growth and movement, and immune
signaling. Therefore, it should not be surprising that altered phosphorylation plays critical
roles in a variety of human diseases, including cancer (Gross et al., 2015; Zhang et al.,
2009), neurodegeneration (Wang et al., 2004), and diabetes (Prada and Saad, 2013;
Winder and Hardie, 1999). Accordingly, there is significant interest in targeting
small-molecule inhibitors to kinases that are associated with specific pathologies. The
first significant success in this area was the development of imatinib for the inhibition of
the BCR-Abl kinase that drives chronic myelogenous leukemia (CML) (Capdeville et al.,
2002). This has been followed by the clinical development of other inhibitors of growth
factor receptor kinases, Bruton’s tyrosine kinase (BTK) (Pan et al., 2007), and others.
Despite these exciting results, gaining a complete understanding of the many roles of
phosphorylation remains extremely challenging.
5
Figure 1-3. Encoding protein phosphorylation. (A) A family of ~500 kinases will transfer a
phosphate group to certain amino acid side-chains, including serine, threonine, tyrosine, and
histidine. (B) Proteins can be synthesized using native chemical ligation (NCL). NCL involves
the specific reaction of C-terminal thioesters and N-terminal cysteine residues to form native
amide bonds. (C) Recombinant protein thioesters for use in NCL reactions can be created using
proteins termed inteins, which catalyze the formation of a branched protein thioester that can be
intercepted with exogenous thiols. (D) Unnatural amino acids can be site-specifically
incorporated into proteins by using a combination of a tRNA synthetase enzyme that will charge
an amber suppressor tRNA with an unnatural amino acid and a corresponding amber stop codon
in mRNA.
There are approximately 500 kinases in the mammalian genome (Manning et al., 2002),
100 protein phosphatases that can antagonize these modifications {Alonso:2004il}, and
hundreds of proteins with phosphate-binding domains to “read-out” a site-specific,
6
phosphorylated signal (Yaffe and Elia, 2001; Yaffe, 2002). Complicating this system
further, the precise timing and spatial localization of the modifications also play key roles
in determining the cellular response to phosphorylation signals. Importantly, chemistry
has made significant contributions to both the preparation and the analysis of
site-specifically phosphorylated proteins and the development of new methodologies that
link a specific kinase to its substrates.
Encoding phosphorylation
The majority of posttranslationally modified proteins, including phosphorylated ones,
have been generated in vitro using protein semisynthesis. Protein semisynthesis relies on
the native chemical ligation (NCL) reaction for the preparation of full-length proteins
from smaller fragments (Figure 1-3B) (Dawson et al., 1994). Since these smaller
fragments are accessible by solid-phase peptide synthesis, they can be chemically
modified with essentially any functionality, including PTMs and PTM analogs. NCL
relies on the reversible transthioesterification reaction between a peptide/protein with a
C-terminal thioester and another peptide/protein with an N-terminal cysteine residue,
which results in a thioester linkage between the two fragments. This is quickly followed
by an essentially irreversible sulfur to nitrogen acyl-transfer, resulting in a native amide
bond. This reaction proceeds in water, often with or without denaturants, and without the
need for protecting groups on the amino acid side-chains or PTMs of interest. Several
solid-phase resins have been developed for the preparation of synthetic peptide thioesters
using either Boc- or Fmoc-based peptide synthesis chemistries, and these peptides can
7
then be reacted with either synthetic or recombinant protein fragments that have
N-terminal cysteines. In the other orientation, recombinant protein thioesters can be
prepared by taking advantage of the naturally occurring process of protein splicing
(Figure 1-3C) (Vila-Perelló and Muir, 2010). Specifically, proteins or protein fragments
can be genetically fused in-frame to proteins termed inteins. These inteins will catalyze a
nitrogen to sulfur acyl-transfer to generate a linked protein thioester that can be
intercepted with exogenous thiols to give recombinant protein thioesters. These proteins
can then undergo a variant of NCL, termed expressed protein ligation (EPL) (Muir et al.,
1998). Notably, the recent development of ultra-fast inteins has greatly improved the
efficiency of the preparation of protein thioesters compared to commercially available
intein constructs (Shah et al., 2012).
Phosphorylated tyrosine, serine, and threonine residues and their corresponding
non-hydrolyzable mimics have been site-specifically incorporated into proteins using
semisynthesis. The first phosphorylated protein to be prepared using semisynthesis was
the tyrosine kinase Csk, which was prepared bearing a phosphorylated tyrosine residue in
its C-terminus during the initial report of EPL (Muir et al., 1998). Csk is responsible for
the phosphorylation of Src and Src family members at their C-terminal tail regions, which
leads to a conformational change driven by a intramolecular interaction between the
phosphorylated tail and SH2 domain in Src. Using EPL, the authors prepared a unnatural
chimera between Csk and a C-terminal phosphorylated peptide and showed that this
results in an intramolecular interaction that increases the kinase activity of Csk towards
8
its substrates. Native phosphorylated serine and threonine residues have also been
incorporated into proteins using semisynthesis. One of the best examples of this has been
the series of experiments aimed at understanding the TGFβ signaling pathway
(Massagué, 2012). Extracellular TGFβ will engage with TGFβ receptor kinase (TGFβR),
which will then go on to activate Smad transcription factors through the phosphorylation
of two serine residues in their extreme C-termini. During this process, a subunit of the
receptor itself becomes phosphorylated on several serine residues. Using NCL, a soluble
fragment of the TGFβR was prepared with four such phosphorylation sites on its
N-terminus (Huse et al., 2001). In vitro studies with this protein demonstrated that it has
improved binding to its substrate, Smad2, and reduced affinity for the protein inhibitor of
the pathway, FKBP12. The consequences of the downstream phosphorylation on Smad2
were then examined. Using EPL, Smad2 was prepared bearing different phosphorylation
patterns at serines 465 and 467, revealing that the trimerization of the transcription factor
is largely driven by phosphorylation at residue 465, with a smaller contribution from
modification of serine 467 (Ottesen et al., 2004). In a subsequent elegant display of the
flexibility of EPL, phosphorylated Smad2 was also prepared with photoactivatable groups
and fluorescent dyes, enabling the analysis of the kinetics of relocalization of trimerized
Smad2 to the nucleus (Hahn and Muir, 2004; Pellois et al., 2004). In addition to the
incorporation of natural phosphorylated residues, protein semisynthesis has also been
used for the site-specific incorporation of non-hydrolyzable, difluoro- and
non-substituted-methylene phosphonate analogs. In the case of tyrosine phosphorylation,
these analogs have been most useful for the study of protein tyrosine phosphatases
Type to enter text
9
(PTPases). These enzymes can be phosphorylated at their C-termini, but the effect of
these modifications is difficult to study as the PTPase will remove their own
phosphorylation marks. Incorporation of non-hydrolyzable analogs of phosphorylated
tyrosine using EPL overcame this limitation and showed that phosphate modifications
had site-specific effects on the PTPase enzymatic activity (Lu et al., 2001; 2003;
Schwarzer et al., 2006; Zhang et al., 2003). Serine and threonine phosphorylation have
also been studied using the same approach. For example, EPL was used to prepare a
semisynthetic version of casein kinase II alpha (CK2α) with a difluoro-phosphonate at
threonine residue 344. A subsequent combination of microinjection and in vitro assays
demonstrated that this modification increases the cellular stability of CK2α and alters its
substrate selectivity (Tarrant et al., 2012). More recently, Lashuel and coworkers used
EPL to prepare both unmodified and phosphorylated versions the Huntingtin exon 1
protein that is prone to aggregation that causes Huntington’s disease {Ansaloni:2014ka}.
Using these proteins, the author showed that phosphorylation at threonine residue 3
significantly slows the aggregation of this protein.
Although semisynthesis has been quite powerful for the preparation of phosphorylated
proteins, it does require some synthetic expertise and in some cases the refolding of
proteins after purification. Another strategy that has the potential to overcome these
issues, which has been used with success for the site-specific incorporation of
phosphorylated residues or their structural analogs is based on the use of the expanded
genetic code and unnatural amino acids (Figure 1-3D). This technique relies on the
10
recognition of a stop codon (typically the amber stop codon UAG) by an engineered
tRNA
CUA that has been chemically or enzymatically charged with an unnatural amino
acid, thereby generating a orthogonal codon that can be read out by the ribosome during
protein translation (Chin, 2014; Lang and Chin, 2014; Liu and Schultz, 2010). Dozens of
unnatural amino acids have now been incorporated into proteins in E. coli, yeast, and
mammalian cells using this system. One of the most successful orthogonal
aminoacyl-tRNA synthetase/tRNA
CUA pairs has been based on the tyrosine pathway
(TyrRS/tRNA
Tyr
) from Methanococcus jannaschii. Although this system has not yet been
engineered to incorporate phosphorylated tyrosine, it has been used to site-specifically
incorporate the tyrosine analog, p-methoxylmethyl-phenylalanine, into recombinant
protein in E. coli (Xie et al., 2007). Specifically, the target protein this study focused on
was the transcription factor signal transducer and activator of transcription-1 (STAT1),
and the procedure generated a constitutively active protein. The approach developed to
incorporate phosphorylated serine into proteins is somewhat different and takes
advantage of an interesting two-step pathway for the introduction of cysteine residues
into proteins used by some methanogenic archaea. For these organisms the first step in
making cysteine aminoacylated tRNA needed for protein synthesis, is to attach
phosphorylated serine onto a tRNA (tRNA
Cys
GCA) using the phosphoseryl-tRNA
synthetase (SepRS). Then a second enzyme converts the phosphorylated serine to
cysteine to give tRNA
Cys
GCA that is ready for protein synthesis. This raises an interesting
question of how methanogenic archea select for the final product, tRNA
Cys
GCA, and avoid
incorporating the intermediate tRNA carrying a phosphorylated serine. One explanation
11
put forward was that the corresponding tRNA
Sep
GCA would be a poor substrate for the
elongation factor EF-Tu and thus discriminated against by the protein synthesis
machinery. This suggested that EF-Tu could be engineered to favor incorporation of
phosphorylated serine into recombinant proteins. This was done and worked as expected
(Lee et al., 2013; Park et al., 2011) although the yields were lower than for other
unnatural amino acid systems. More recently, this same orthogonal system was improved
to allow for the incorporation of phosphorylated serine and its difluoro-phosphonate
analog (Rogerson et al., 2015). Specifically, engineering of the aminoacyl-tRNA
synthetase/tRNA
CUA pair (SepRS/tRNA
pSer
CUA) yielded mutants that will incorporate
phosphorylated serine into recombinant proteins in E. coli 18 times more efficiently
without the need for engineering of EF-Tu. Notably, in E. coli phosphorylated serine is
generated in the serine biosynthesis pathway and this endogenous reaction can serve as
the source of modified serine by the system. Additionally, mutation of this biosynthetic
pathway to reduce the cellular concentration of phosphorylated serine enabled the
incorporation of an exogenously added phosphonate analog of serine. Undoubtedly, the
further optimization of these technologies, including other orthogonal synthetase/tRNA
pairs, will allow for the incorporation of both phosphorylated tyrosine and serine in a
variety of cell types.
Decoding phosphorylation
The identification of phosphorylated proteins under different cellular states, in different
tissues, etc. is critical for understanding the specific roles of this PTM. Site- and
12
pan-specific antibodies can give a broad picture of the level of individual or global
phosphorylation levels, respectively, and enrichment methods (e.g., metal affinity
chromatography) coupled with proteomics has enabled the global characterization of the
phosphoproteome. However, mapping kinases to their specific substrates has been more
difficult and has inspired the creation of different chemical methods. The first such
method uses engineered protein kinases, termed analog-sensitive kinases, that are
optimized to exclusively accept an analog of ATP that cannot be utilized by any wild-type
kinases (Figure 1-4A) (Liu et al., 1998; Shah et al., 1997; Ubersax et al., 2003; Zhang et
al., 2005). Specifically, this method takes advantage of the fact that most kinases have a
large (threonine, methionine, etc.) conserved residue in the ATP-binding pocket. This
gate-keeper residue prevents the binding of an N
6
-benzyl-modified ATP in the kinase
active site. However, when the gate-keeper residue is mutated to a smaller amino acid
(e.g., glycine), the resulting analog-sensitive kinases will use bulky ATP derivatives, such
as N
6
-(benzyl) ATP, to phosphorylate their substrates. This method has been generally
termed the “bump-hole” strategy. When radioactive N
6
-benzyl ATP is used with a specific
analog-sensitive kinase, the substrates of that kinase can be visualized in a complex
cellular lysate. Notably, these same mutant kinases can be selectively inhibited by bulky
ATP-competitive kinase inhibitors (Bishop et al., 1998; 2000). By genetically
incorporating the mutant kinase into a cell or organism, one can achieve highly selective
inhibition and investigate the phenotypic and biochemical effects. While this method
enables the visualization of specific kinase substrates and has been shown to be broadly
applicable, it does not immediately allow for the unbiased identification of these
13
substrates from a complex mixture. To accomplish this, the fact that kinases will transfer
the ATPγS analog to thiophosphorylate proteins was exploited (Allen et al., 2005; 2007).
In this elegant strategy, the resulting protein thiophosphate can then be alkylated with a
p-nitro-benzyl group (Figure 1-4B). An antibody that specifically recognizes this
alkylated structure can then be used to either visualize or enrich kinase substrates for
proteomic identification. Combining analog-sensitive kinases with this approach allows
for the enrichment and identification of the substrates of in principal any kinase of
interest. Subsequent iterations of this strategy have used direct alkylation of the
thiophosphate by solid-phase resins followed by their selective elution (Blethrow et al.,
2008), and the selective capping of cysteine residues to eliminate the potential
enrichment of false-positive proteins (Garber and Carlson, 2013).
14
Figure 1-4. Decoding protein phosphorylation. (A) Development of analog-sensitive kinases
using a bump-hole strategy. Wild-type kinases are incapable of using the “bumped” ATP analog
N
6
-benzyl ATP. However, mutation of the kinase in its active site creates a “hole” that will allow
N
6
-benzyl ATP to function as a substrate. (B) Identification of kinase substrates using
analog-sensitive kinases. A gatekeeper mutant kinase of interest is first incubated with cell lysate
and N
6
-benzyl-ATPɣS, resulting in selective thiophosphorylation of that kinase’s substrates. The
resulting thiophosphate is then alkylated to generate a p-nitro-benzyl group that is recognized by
a specific antibody for visualization or enrichment. (C) Linking a known phosphorylated substrate
with the kinase responsible using cross-linking. An ATP based cross-linker is first incubated with
a complex mixture of kinases in a cell lysate, transferring a Michael acceptor to the conserved,
catalytic lysine residue. Then a substrate peptide bearing a cysteine residue at the known site of
phosphorylation is added, yielding a covalent cross-link between the substrate and kinase of
interest.
The thiophosphate method above starts with a kinase of interest and proceeds to the
identification of the corresponding substrate proteins. An equally important challenge is
15
to map a known phosphorylated protein back to the kinase responsible for the
modification. Again chemistry contributed to solving this problem, in particular with the
development of several small molecule cross-linkers. One class of these compounds
relies on a three-component reaction between a substrate protein (or peptide) of interest,
the small-molecule, and the conserved active-site lysine residue of a kinase (Riel-Mehan
and Shokat, 2014; Statsuk et al., 2008). All of these cross-linkers rely on the genetic
incorporation of a cysteine at the normal site of phosphorylation in the substrate. In the
most robust system to date, the cross-linker is an analog of ATP that binds to the kinase
active site and results in the modification of the catalytic lysine to generate a
methacrylamide, which will then undergo a Michael-addition reaction with the
cysteine-containing substrate bait (Figure 1-4C). This method has been successfully
applied to a model system in a complex lysate, but unfortunately, not to date for the
unbiased identification of a kinase-substrate pair. An alternative strategy was also
explored that relied on the incorporation of two photo-cross-linking groups into an analog
of ATP (Parang et al., 2002). Upon photo-irradiation, one of the cross-linkers near the
adenosine ring will covalently label the kinase, while the other located near the gamma
phosphate traps the substrate protein. This system was successfully applied to an in vitro
model system but has not been employed in a large scale discovery experiment.
Glycosylation
Glycosylation is a common PTM where carbohydrate chains of various lengths and
composition are added to a large fraction of proteins, highlighting the utility of
16
glycosylation in biological events. Specifically, proteins that reside in the secretory
pathway, at the cell surface, or are excreted into extracellular space, can be modified by
typically large and elaborate oligosaccharides in the endoplasmic reticulum (ER) and
Golgi, resulting in N-linked and mucin O-linked glycosylation. The most common type
of O-linked glycosylation, mucin-type glycosylation, is characterized by the core addition
of N-acetyl-galactosamine (GalNAc) through an α-O-linkage to the β-hydroxyl group of
serine or threonine residues. This monosaccharide can then be elaborated through
addition of sialic acid, fucose and/or additional units of Galβ1,4GlcNAc to form large,
branched glycan structures. Mucin-type glycosylation (Hang and Bertozzi, 2005) is found
on many cell-surface proteins and has been shown to play an essential role in protein
localization and cell-cell communication in the immune response (Wolfert and Boons,
2013). A more prevalent type of protein glycosylation is N-linked glycosylation. In
contrast to the synthesis of mucin-type glycoproteins, N-linked glycoproteins are
synthesized first by the assembly of a dolichol-linked oligosaccharide precursor in the
cytosol and ER. The large structure is subsequently transferred by an oligosaccharide
transferase to the amide side chain of an asparagine residue of a nascent polypeptide.
Intracellularly, N-linked glycans regulate protein trafficking and act as quality control for
protein folding (Helenius and Aebi, 2001). Outside the cell, N-linked glycans can
function as ligand receptors and have been shown to mediate cell-interactions with
proteins, other cells and pathogens. Serine and threonine residues of intracellular proteins
can also be O-glycosylated by the single monosaccharide, N-acetyl-glucosamine
(GlcNAc). Termed, O-GlcNAc, this PTM is known to affect protein localization and
17
signal transduction. Unlike cell-surface glycosylation, O-GlcNAcylation is a dynamic
modification that plays critical roles in cellular responses to changes in metabolism and
stress, particularly in diseases such as cancer, diabetes and neurodegeneration (Hart et al.,
2011). Additionally, altered states of O-GlcNAc glycosylation have also been associated
with oncogenic transformation (Ma and V osseller, 2013). All types of glycoproteins are
fundamental in biology and therefore have conjured a great deal of interest within the
scientific community. Their study, however, has proved challenging due to the fact that
naturally occurring glycoproteins are not synthesized homogeneously and current
chromatographic methods are not sophisticated enough to separate the variety of
glycoforms on a reasonable scale. Further more, complex oligosaccharide structures are
not synthesized in a template-dependent manner; therefore, no straightforward genetic
methods for controlling expression of specific carbohydrates exist, leaving researchers
with a restricted set of tools for their study. Therefore, there is a demand for a source of
structurally homogeneous glycoproteins for functional studies (Davis, 2002; Grogan et
al., 2002), which involves methodologies from carbohydrate and peptide chemistries
alike, as well as the creation of chemical tools for the visualization and identification of
glycoproteins.
Encoding glycosylation
The most common method to synthesize glycopeptides/glycoproteins relies heavily on
solid phase peptide synthesis (SPPS) to incorporate glycosylated amino acids onto
growing amino acid chains resulting in a native glycan-amino acid bond. The majority of
18
naturally occurring, O-linked glycans can be synthesized by the glycosylation of
protected serine or threonine residues using common glycosyl donors, resulting in α-O-
linked “cassettes” for solid phase peptide synthesis (Figure 1-5A). For example, the
Boons lab has demonstrated the synthesis of the T
n and T f antigen building blocks using
the thiophilic Ph
2SO/Tf 2O promoter system for subsequent glycosylation of an
Fmoc-protected threonine for the synthesis of glycopeptides (Cato et al., 2005). For the
synthesis of larger glycan structures, the core α-O-Ser/Thr linkage is formed first, and
orthogonally removable protecting groups can then be used to direct the additional
elaboration of branching sugars. Impressively, this linear, cassette-based method has been
employed by the Kunz lab in the synthesis of part of the CD62P ligand PSGL-1 that is
involved in the inflammatory response and leukocyte recognition. The resulting
18-residue fragment containing an O-linked hexasaccharide was synthesized in sufficient
amounts for use in biomedical studies (Baumann et al., 2008). In the synthesis of
N-linked glycopeptides, glycosylated asparagine cassettes are most commonly prepared
before or after SPPS through the reaction of a protected or deprotected anomeric
glycosyl-amine with an aspartate residue (Figure 1-5B). For example, the Keissling lab
has developed a method in which the glycosyl-amide bond is formed through reaction
with glycosyl azides and asparagine-derived phosphinothioesters, avoiding anomerization
and formation of isomeric mixtures (He et al., 2004). More recently, Doores and
co-workers have demonstrated a elegant procedure for glycosyl-asparagine ligation that
can be utilized with fully deprotected substrates, is stereocontrolled and is compatible
with linear and convergent approaches that are free from the need of complex auxiliaries
19
(Doores et al., 2006). The development of site-selective chemical glycosylation of
proteins has allowed for the synthesis of larger and more complex glycopeptides;
however, site-specific installation of native glycans in full-length proteins remains
challenging. To overcome the size limitation of SPPS, larger glycoproteins can be
prepared in a stepwise fashion using NCL. For example, to mechanistically explore the
role N-linked glycosylation during protein folding in the ER, Kajihara and Ito
synthesized misfolded, interleukin-8 (IL-8, CXCL8) bearing an N-linked glycan. Using
this material, they found that the enzyme UDP-glucose:glycoprotein glucosyltransferase
(UGGT), prefers misfolded over correctly folded glycoproteins, indicating that UGGT
plays a distinct role in quality control of protein folding (Izumi et al., 2012). Extending
the use of NCL, the Danishefsky group published the complete chemical synthesis of the
signaling glycoprotein, erythropoietin, with all carbohydrate domains at all native
glycosylation sites (Wang et al., 2013a). Our lab has utilized EPL to study the function of
O-GlcNAc modification on the protein α-synuclein, the major aggregating protein
associated with Parkinson’s disease and dementia with Lewy Bodies (DLBs). Chemical
synthesis of homogenous, full-length, O-GlcNAcylated α-synuclein was achieved using
EPL in three steps and subsequent aggregation assays showed that O-GlcNAc
modification at threonine residue 72 inhibits α-synuclein aggregation and is non-toxic to
neurons in cell culture (Marotta et al., 2015).
Extension of these synthetic techniques to larger glycans has been stymied due to the low
efficiency of couplings and the complicated nature of both the carbohydrate and amino
20
acid protecting group chemistries involved in SPPS. Additionally, some glycosidic bonds
cannot survive the strong acidity of final deprotection conditions of peptides (e.g. TFA).
To address these shortcomings, chemoenzymatic methods to catalyze the addition of
single monosaccharides onto to synthetic glycopeptides or growing glycan structures
have been developed as an alternative. Sometimes referred to as glycoprotein remodeling,
recombinant enzymes with unique glycosyltransferase/glycosidase activity can be utilized
in a stepwise fashion to produce more complex glycopeptide libraries that are out of
reach of canonical chemical synthesis. For example, Bello and co-workers recently
demonstrated the stepwise, chemoenzymatic synthesis of O-linked glycosylated MUC1
peptides utilizing Drosophila glycosyltransferases (Bello et al., 2014). In the area of
N-linked glycoproteins, the Boons and Paulson labs collaborated to create a library of
complex multi-antennary glycans by utilizing a chemically synthesized core
oligosaccharide to which asymmetric, branched antenna were chemoenzymatically
installed (Wang et al., 2013c). The resulting library was printed on a microarray and was
subsequently screened for binding to lectins and influenza-virus hemagglutinins (HAs).
The results illustrate the complex, environment-dependent recognition of glycan epitopes
and highlight the importance of understanding the receptor specificity to further elucidate
their biological consequences in disease. Incorporating complex N-glycans into peptides
can also be difficult due to the involved chemistry required for their synthesis and
conjugation to aspartic acid. In order to allow for the use of isolated N-linked glycans and
improve the coupling of isolated and synthesized structures onto peptides, endo-β-N-
acetylglucosaminidases (ENGases) have been utilized to perform transglycosylation
21
reactions (Figure 1-5C) (Wang, 2011). These enzymes normally cleave N-linked glycans,
leaving only the first N-acetylglucosamine (GlcNAc) residue at the N-linked site(s).
However, through mutagenesis and screening of reaction conditions, it was found that
certain ENGases will essentially perform this reaction in reverse to install complex
N-linked glycans onto single GlcNAc residues on peptides. In an early example of this
method, the enzyme EndoM was used to prepare glycosylated versions of the the
N-linked glycopeptide hormone calcitonin, a 32-amino acid calcium-regulating hormone
used in the treatment for hypercalcemia, Paget’s disease and osteoporosis (Haneda et al.,
1998). Other Endo enzymes have also been explored, such as EndoF2 and F3, that were
found to glycosylate α-1,6-fucosylated GlcNAc derivatives, producing native, core
fucosylated, complex-type glycopeptides (Huang et al., 2011). More recently, the Davis
lab has explored a bacterial endoglycosidase, EndoS, that is complimentary to other
endoglycosidases, EndoA and EndoH (Goodfellow et al., 2012). Specifically, the authors
showed that a synthesized tetrasaccharide oxazoline could be transferred onto human IgG
using EndoS and that EndoS also shows tolerance to the presence of core fucosylation,
broadening its synthetic utility.
Although useful, chemoenyzmatic methods do not always allow for control over the site
of glycosylation. Therefore, to further increase the selectivity of protein glycosylation,
various chemoselective methods have been developed. Specifically, pre-formed glycans
are attached to peptides or proteins through mild and selective chemical reactions that
tolerate numerous functional groups, therefore minimizing the need for protecting groups.
22
The Bertozzi group pioneered this area in the synthesis of O-linked glycopeptides that
contain unnatural bonds at the C-6 and C-3 branch points, as oximes
{Rodriguez:1997up} and thioethers respectively (Marcaurelle and Bertozzi, 2001), as
well as thioether linkages to install antennae onto a N-linked core (Pratt and Bertozzi,
2003). More recently, other methods that take advantage of the native chemistry of the
cysteine thiol have also been utilized. Notably, Dondoni and co-workers demonstrated a
ligation strategy that utilizes the thiol-ene coupling (TEC) reaction in which an alkenyl
C-glycoside is coupled via photo-irradiation with a protein or peptide containing a free
sulfhydryl group resulting in a thioether bond (Dondoni et al., 2009). Thioglycosides are
an attractive synthetic mimic due to their likeness in length to the native glycosidic bond
as well as their increased stability (Chalker et al., 2011). The Davis group published a
“tag-and-modify” method that involved the use of TEC-chemistry in the synthesis of
S-glycosyl amino acids through the addition of glycosyl-thiols to homoallylglycine (Hag)
following its incorporation into a peptide/protein as a nonnatural amino acid (Floyd et al.,
2009). The tag-and-modify method has also utilized bioorthogonal azide-alkyne
cycloaddition reactions. Specifically, through the introduction of the unnatural amino acid
azidohomoalanine (Aha), GlcNAc modified protein Np276 from Nostoc punctiforme was
synthesized (Fernández-González et al., 2010). In a similar fashion, an alkyne-containing
nonnatural amino acid, homopropargylglycine (Hpg) can be incorporated as a methionine
surrogate. Interestingly, the Davis lab utilized both Aha and Hpg to enable attachment of
multiple glycans to bacterially expressed protein scaffolds (van Kasteren et al., 2007).
23
Figure 1-5. Protein glycosylation. (A) O-linked glycopeptides are typically synthesized through
the solution-phase preparation of an Fmoc-protected amino-acid cassette that can be used directly
in solid phase peptide synthesis. (B) N-linked glycopeptides have been prepared using the
cassette approach but can alternatively synthesized after peptide synthesis through the coupling of
an aspartic acid residue to a glycosyl-amine or amine equivalent. (C) Enzymatic installation of
large N-linked glycans onto peptides and proteins with transglycosylation reactions. Under
certain reaction conditions, some endoglycosidases will use isolated or synthesized glycans as
substrates and transfer them onto single N-acetyl-glucosamine residues on peptides or proteins.
(D) Metabolic chemical reporters of glycosylation. Living cells are treated with analogs of
monosaccharides containing bioorthogonal functionality (e.g., an alkyne). These reporters are
metabolized by the cell and installed onto proteins. Bioorthogonal reactions can then be
performed for the installation of visualization or affinity tags. (E) Chemoenzymatic detection of
O-GlcNAc modifications. Endogenous O-GlcNAc modifications in a cell lysate can be
enzymatically modified with a GalNAz residue, followed by the installation of tags using
bioorthogonal chemistry.
24
Decoding protein glycosylation
Different cellular states and tissues can display unique glycoproteins and glycan
structures, and direct identification of modified proteins is essential to uncovering the role
of this posttranslational modification. Pioneered by the Bertozzi lab, several labs have
investigated the use of metabolic chemical reporters (MCRs) for the visualization and
identification of glycosylated proteins (Figure 1-5D). Typically, this technique involves
treatment of cells with chemically synthesized analogs of naturally occurring
monosaccharides that contain bioorthogonal reactivity (commonly an azide or alkyne).
These MCRs are then accepted by living systems and metabolically converted into
high-energy UDP-sugar donors that are subsequently utilized in their incorporation into
glycans by endogenous glycosyltransferases. The first completely orthogonal MCR,
Ac
4ManNAz, was developed by the Bertozzi lab (Saxon, 2000). Once this compound
diffuses into cells, the O-acetates are removed by endogenous lipases and the resulting
ManNAz is biosynthetically converted to the corresponding azide-containing sialic acid
analog and enzymatically installed onto the termini of cell surface glycans. Subsequent
bioorthogonal labeling of the azide, using reactions like the Staudinger ligation or the
copper-catalyzed azide-alkyne cycloaddition can then be used for the selective
installation of visualization or affinity tags. Since this transformative initial study, many
different MCRs of glycosylation have been developed, including other azide-bearing
monosaccharides like Ac
4GalNAz (Hang et al., 2003), Ac 4GlcNAz (V ocadlo et al., 2003),
and Ac
4FucAz for the labeling of mucin O-linked glycans, O-GlcNAc modifications, and
fucose-containing glycans, respectively. Our lab subsequently found that the alkyne
25
MCR, Ac 4GlcNAlk, displayed improved signal to noise and was more selective for
O-GlcNAc modifications compared to Ac
4GlcNAz (Zaro et al., 2011b), and other alkyne
MCRs for the visualization of sialic acid and fucose have also been developed (Hsu et al.,
2007). Recently, research has also focused on the effects of metabolic cross-talk between
glycosylation pathways on the cellular fate of different metabolic chemical reporters. For
example, it has been shown that UDP-GalNAz and UDP-GlcNAz can be enzymatically
interconverted in living cells (Boyce et al., 2011; Zaro et al., 2011b), allowing the
proteomic identification of O-GlcNAcylated proteins from cells treated with Ac4GalNAz
(Palaniappan et al., 2013). While Ac4GalNAz-treatment appears to label O-GlcNAcylated
proteins more robustly under some conditions (Boyce et al., 2011), its lack of specificity
requires cell fractionation to separate O-GlcNAcylated and mucin O-linked
glycoproteins. Overcoming this limitation, treatment with peracetylated
6-azido-6-dexoy-N-acetyl-glucosamine (Ac36AzGlcNAc, Figure 3D) was recently shown
to selectively label O-GlcNAcylated proteins (Chuh et al., 2014). Notably, using
biochemistry and proteomics the same study demonstrated that 6AzGlcNAc is the most
selective for O-GlcNAcylation, followed by GlcNAz and finally the relatively
nonspecific GalNAz. These results suggest that selective metabolic chemical reporters of
other types of glycoproteins could be developed in the future. Currently, there are also
monosaccharide MCRs that contain other bioorthogonal functionalities, including alkenes
(Niederwieser et al., 2013; Späte et al., 2014) and cyclopropenes (Cole et al., 2013;
Patterson et al., 2014a; 2012), which can take advantage of the rapid tetrazine ligation for
26
the installation of tags. Notably, monosaccharide MCRs have been used in a variety of
contexts from cell culture to living animals for the visualization (Chuh and Pratt, 2015a)
and proteomic identification (Chuh and Pratt, 2015b) of glycoproteins. For example,
Ac
4ManNAz and Ac 4GalNAz have been used to visualize cell surface glycans in
zebrafish embryos {Laughlin:2008ky, Baskin:2010ir} and C. elegans (Laughlin and
Bertozzi, 2009). Additionally, many monosaccharide MCRs have been used to perform
proteomic analysis of different glycoproteins, including quantitative comparisons of
cancer versus normal cell populations using Ac
4GalNAz (Slade et al., 2012). Recently,
the Bertozzi lab has developed a technique termed isotope-targeted glycoproteomics
(IsoTaG), a mass-independent chemical glycoproteomics method for the identification of
intact, metabolically labelled (using Ac
4ManNAz or Ac 4GalNAz) glycopeptides from the
total proteome (Woo et al., 2015). In contrast to traditional tandem MS proteomics that is
performed on the most abundant species in the total-scan mass spectra, IsoTaG enables
the specific detection of glycoproteins with isotopic signatures, improving the selection of
low-abundance glycopeptides. MCRs have been transformative in their ability to report
on different types of glycoproteins; however they necessarily compete with natural
metabolites, meaning that they are inherently unreliable indicators of the overall levels of
a modification. To address this limitation in the area of O-GlcNAc modification, the
Hsieh-Wilson lab has developed chemoenzymatic detection methods that enable the
capture of a snap-shot of endogenous O-GlcNAc modified proteins. In the most common
iteration of this technology, incubation of cell lysates with an recombinantly expressed,
mutant β-1,4-galactosyltransferase and chemically prepared UDP-GalNAz results in the
27
transfer of GalNAz to O-GlcNAc residues (Figure 1-5E) (Clark et al., 2008). The
resulting azide-containing disaccharide can be bioorthogonally reacted with different
visualization and affinity tags. For example, small polyethylene glycol (PEG) chains can
be installed that will shift the mass of O-GlcNAc modified proteins, enabling the
stoichiometry of some O-GlcNAc modifications to be quantitated using Western blotting
(Ortiz-Meoz et al., 2014; Rexach et al., 2010). Additionally, this strategy has been
combined with both β-elimination (Khidekel et al., 2007) and electron transfer
dissociation mass spectrometry [e.g., (Alfaro et al., 2012)] for the proteomic
identification of O-GlcNAcylated proteins and a subset of modification sites.
Identification of Glycosylated Proteins
Many types of cancer display altered levels of both sialic acid and mucin O-linked
glycans. Treatment of highly metastatic prostate cancer with Ac
4ManNAz (Figure 3A)
allowed for the identification of cell-surface proteins, the majority of which were
implicated in cell motility, migration and invasion, supporting a potential role for sialic
acid in metastasis (Yang et al., 2011). In a separate study, a prostate cancer cell-line was
treated with Ac
4GalNAz to enable the enrichment and identification of 29 cell-surface
glycoproteins, including many proteins involved in cell adhesion (Hubbard et al., 2011).
More recently, Ac
4GalNAz (Figure 3B) was used in combination with iTRAQ (isobaric
tag for relative and absolute quantitation) to enrich and compare the secreted mucin
glycoproteins from two different CHO cell-lines (Slade et al., 2012), demonstrating that
28
these chemical reporters could be used in the future to compare cellular glycoproteins
from two states (e.g., cancer vs. normal tissue).
O-GlcNAc modification of serine and threonine residues is an abundant
modification of proteins in the cytosol, nucleus and mitochondria. Unlike cell-surface
glycosylation, O-GlcNAcylation is a dynamic modification that plays critical roles in
cellular responses to changes in metabolism and stress, particularly in diseases such as
cancer, diabetes and neurodegeneration (Hart et al., 2011). The first metabolic chemical
reporter of O-GlcNAcylation was peracetylated N-azidoacetyl-glucosamine (Ac
4GlcNAz,
Figure 3D) (V ocadlo et al., 2003), which was recently used to identify over one-thousand
potentially O-GlcNAcylated proteins from a single cell-line and confirm the modification
sites on 80 of these substrates (Hahne et al., 2013). To provide improved signal-to-noise
in the CuAAC bioorthogonal reaction, the alkyne-analog N-pentynyl-glucosamine
(GlcNAlk, Figure 3D) has also been developed and revealed the O-GlcNAc modification
of the ubiquitin ligase NEDD4 (Zaro et al., 2011b). This chemical reporter was also used
to identify O-GlcNAcylated proteins associated with diabetic retinopathy
{Gurel:2014bn}. As an alternative to metabolic methods, a chemoenzymatic method has
been developed (Figure 3E). Specifically, incubation of cell-lysates with an engineered
β-1,4-galactosyltransferase and azide- or ketone-containing UDP-donor sugar results in
modification of O-GlcNAc residues (Clark et al., 2008). The resulting disaccharide can
then be subjected to bioorthogonal labeling for visualization and proteomics. For
example, this chemoenzymatic chemical reporter was recently used in conjunction with a
29
photo-cleavable biotin tag to enrich and identify 274 O-GlcNAcylated proteins and 458
specific O-GlcNAc modification sites on 195 proteins (Alfaro et al., 2012).
Recently, research has also focused on the effects of metabolic cross-talk between
glycosylation pathways on the cellular fate of different metabolic chemical reporters. For
example, it has been shown that UDP-GalNAz and UDP-GlcNAz can be enzymatically
interconverted in living cells (Boyce et al., 2011; Zaro et al., 2011b), allowing the
proteomic identification of O-GlcNAcylated proteins from cells treated with Ac4GalNAz
(Palaniappan et al., 2013). While Ac4GalNAz-treatment appears to label O-GlcNAcylated
proteins more robustly under some conditions (Boyce et al., 2011), its lack of specificity
requires cell fractionation to separate O-GlcNAcylated and mucin O-linked
glycoproteins. Overcoming this limitation, treatment with peracetylated
6-azido-6-dexoy-N-acetyl-glucosamine (Ac36AzGlcNAc, Figure 3D) was recently shown
to selectively label O-GlcNAcylated proteins (Chuh et al., 2014). Notably, using
biochemistry and proteomics the same study demonstrated that 6AzGlcNAc is the most
selective for O-GlcNAcylation, followed by GlcNAz and finally the relatively
nonspecific GalNAz. These results suggest that selective metabolic chemical reporters of
other types of glycoproteins could be developed in the future.
Ubiquitination
The covalent addition of a small (76-residue) protein called ubiquitin, to lysine side
chains of protein through isopeptide bonds, is a posttranslational modification that is
30
implicated in numerous cellular processes including proteasomal degradation, signal
transduction, receptor endocytosis, and DNA damage response (Figure 1-6A) (Chen and
Sun, 2009; Komander and Rape, 2012). Ubiquitin is added an enzymatic cascade that
involves three classes of proteins: E1 ubiquitin-activating enzymes, E2
ubiquitin-conjugating enzymes, and E3 ubiquitin ligases(Hershko and Ciechanover,
1998). First, the C-terminal glycine residue of ubiquitin is activated by ATP to yield a
thioester intermediate with the catalytic cysteine of the E1 enzyme. Then, ubiquitin is
transferred onto an E2 conjugating enzyme to generate a second, active site-thioester
intermediate. The final step of the ubiquitin enzymatic cascade is catalyzed by an E3
ligase to form an isopeptide bond between the C-terminal glycine reside of ubiquitin and
the ε-amino group of the target lysine reside of the substrate protein. Interestingly,
proteins can be modified with either a single ubiquitin molecule (monoubiquitination) or
ubiquitin chains (polyubiquitination), both of which give rise to a plethora of ubiquitin
signals. Specifically, monoubiquitination can be found at more than one site, giving rise
to multi-monoubiquitination. On the other hand, polyubiquitination can be formed
through one of the seven ubiquitin lysine resides (Lys6, Lys11, Lys27, Lys29, Lys33,
Lys48 and Lys63) or through the amino terminal methionine residue to generate linear
chains (Kulathu and Komander, 2012). Given all the possible outcomes of ubiquitination,
it is not surprising that this PTM accounts for a wide range of functions within the cell.
Akin to phosphorylation, ubiquitination is a dynamic modification and can be removed
from protein substrates by the action of the enzyme family of deubiquitinases (DUBs)
Akin to phosphorylation, ubiquitylation is a dynamic modification and can be removed
31
from protein substrates by the action of the enzyme family of deubiquitinases (DUBs)
(Komander et al., 2009). Given its pervasive role in cellular function, the understanding
the role of ubiquitination is of the utmost importance, and chemistry has contributed to
this goal through both the synthesis of site-specifically ubiquitinated proteins and the
development of probes for the identification of ubiquitin modifying enzymes.
Encoding ubiquitination
Although the chemical synthesis of ubiquitin was achieved over 20 years ago through
SPPS, the synthesis of ubiquitinated substrate-proteins remained challenging until the
advent of NCL. Muir and co-workers were the first to synthesize a monoubiquitinated
protein, histone H2B, containing a native isopeptide-linkage using EPL to study the
mechanistic role of ubiquitination in enhancing subsequent methylation of lysine residue
79 in histone H3 (H3K79) (Figure 1-6B) (McGinty et al., 2008). The synthetic portion of
the protein was a peptide corresponding to residues 117-125 of H2B. This peptide
contained an N-terminal cysteine residue protected with a photo-labile o-nitrobenzyl
group and a photo-labile, thiol-bearing ligation auxiliary attached to the ε-amino group of
Lys120 through a glycine linker (eventually to become Gly76 of ubiquitin). This ligation
auxiliary then mediated ubiquitination through an NCL reaction with a recombinant
ubiquitin(1-75) C-terminal thioester, and the ligation auxiliary and the cysteine protecting
group were subsequently removed by irradiation with ultraviolet light. A second EPL
reaction was preformed between this ubiquitinated peptide and a recombinant
H2B(1-116) C-terminal thioester to give full-length, ubiquitinated H2B. Since H2B has
32
no native cysteine residues, chemical desulfurization was used to convert the cysteine
required for the ligation to alanine to yield the monoubiquitinated H2B protein with no
mutations. This synthetic protein was used to show that monoubiquitination of H2B
directly stimulates methylation at H3K79 by the methyltransferase hDot1L, and it
inspired a series of important innovations in the synthesis of site-specifically
ubiquitinated proteins. The first significant advanced was the synthesis of δ-
mercaptolysine unnatural amino acid building blocks for solid phase peptide synthesis
first developed by the Brik lab (Ajish Kumar et al., 2009; Kumar et al., 2010) followed
shortly by Ovaa and coworkers (Figure 1-6C) (Oualid et al., 2010). Like the previous
photo-liable ligation auxiliary, this amino acid can be directly incorporated into peptides
and then undergo NCL reactions with ubiquitin C-terminal thioesters, followed by
desulfurization for the site-specific installation of ubiquitin. When combined with heroic
protein chemistry efforts, these amino acids have enabled the synthesis of quite large
proteins. For example, the Parkinson’s disease associated protein α-synuclein was
prepared bearing either mono-, di-, or tetraubiquitination at the physiologically-relevant
lysine residue 12 and the different effects on the protein aggregation and stability was
measured {HajYahya:2013df}. However, the size of many proteins can make their
chemical synthesis difficult. To address this limitation the Chin and Kommander labs
collaborated to use unnatural amino acid mutagenesis to introduce their GOPAL
(genetically encoded orthogonal protection and activated ligation) strategy (Virdee et al.,
2010). Briefly, a lysine residue of interest can be genetically replaced with Nε-(tert-
butyloxycaronyl)-L-lysine (NHBoc-Lys) using unnatural amino acid mutagenesis and the
33
Methanosarcina barkeri MS pyrrolysine tRNA synthetase (MbPyrlRS) and its
corresponding amber suppressor tRNA (MbtRNA
CUA). After recombinant expression, the
remaining lysine residues can be orthogonally protected by treatment with
N-(benzyloxycarbonyloxy)succinimide (Cbz-OSu). After deprotection of the
NHBoc-Lys, a suitably protected ubiquitin molecule can be site-specifically installed,
followed by global deprotection. Unfortunately, the GOPAL system relies on extensive
protection group chemistry that can contribute to poor yields. To overcome this issue, the
Chin lab designed a synthetic scheme in which a δ-thiol-L-lysine is incorporated at the
desired site of ubiquitylation using an improved pyrrolysyl-tRNA synthetase/tRNA
CUA
pair (Figure 1-6D) (Virdee et al., 2011). Using this method, Chin and co-workers
prepared K6-linked diubiquitin and site-specifically ubiquitinated SUMO (small
ubiquitin-like modifier protein) at K11 through the formation of native isopeptide bonds.
A great deal of research has been dedicated to the synthesis of native isopeptide-bond
conjugated ubiquitin through NCL and EPL, and has allowed insight into the biological
roles that this PTM can play. However, many of these approaches are synthetically
challenging and require multiple ligation and purification steps thus limiting the
widespread use of NCL and EPL for the semi-synthesis of ubiquitinated proteins. To
address this shortcoming, several groups have created strategies to introduce ubiquitin
onto substrate proteins through isopeptide bond mimics (Figure 1-6E). One of the first
such methods was a disulfide-directed approach that was developed simultaneously by
the Muir and Zhuang labs (Chatterjee et al., 2010; Chen et al., 2010). This approach takes
34
advantage of a ubiquitin-intein fusion that is trapped with cysteamine to incorporate a
thiol moiety at the C-terminus of ubiquitin (Ub-SH), which can be subsequently activated
as a mixed disulfide by treatment with specific reagents like 2,2’-dithiobis(5-
nitropyridine) (DTNP). The target protein carrying a cysteine residue at the desired
modification site is then reacted with the activated ubiquitin resulting in the formation of
a disulfide between the substrate and ubiquitin moieties. Although, this technique has
been successfully applied to a handful of ubiquitination sites in vitro [e.g., (Abeywardana
et al., 2013; Meier et al., 2012)] the disulfide linkage is not chemically stable. Therefore,
other chemical approaches have been used for the installation of stable analogs of the
ubiquitin linkage. For example, the Strieter lab took advantage of thiol-ene coupling
(TEC) to install thioether linkages for conjugation of ubiquitin to substrate proteins
(Valkevich et al., 2012). Furthermore, they were able to show that this Nε-Gly-L-
homothialysine isopeptide linkage was hydrolyzed by DUBs in a manner similar to that
of the wild-type isopeptide bond, demonstrating that it is a good structural mimic. In yet
another thioether approach, the Brik lab utilized either an α-bromo acetamide or
maleimide synthetically tethered to the C-terminus of ubiquitin that can be subsequently
reacted with a cysteine of a substrate protein (Hemantha et al., 2014).
Copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC) has also been used for the
installation of ubiquitin to a substrate protein via a triazole linkage. For example, the
installation of a cysteine into the target protein followed by treatment with iodoacetamide
ethyl azide allows for the installation of a site-specific azide, which can be reacted with a
ubiquitin bearing a C-terminal alkyne (installed using intein chemistry) (Weikart and
35
Mootz, 2010). More recently, similar strategies have been developed that take advantage
of the incorporation of azide- or alkyne-containing amino acids (Eger et al., 2011; Rösner
et al., 2015; Sommer et al., 2011). Additional methods based on the use of nonnatural
linkages continue to be developed and applied, including propanone (Yin et al., 2000) and
oxime (Shanmugham et al., 2010) linkages.
Figure 1-6. Ubiquitination. (A) Ubiquitination is the addition of the small protein ubiquitin to
protein side chains, most often lysine, resulting in an isopeptide bond. This first modification
event can then be polymerized in various ways to form polyubiquitin chains. (B) Synthesis of
ubiquitinated histone H2B using a photo-cleavable auxiliary. Using an NCL reaction ubiquitin is
first installed onto a synthetic peptide through a lysine residue bearing the auxiliary. Photolysis
then both removes the auxiliary and reveals the N-terminal cysteine residue that can be used in
subsequent NCL reactions. (C) Ubiquitination of proteins using a δ-mercapto-lysine residue. The
δ-mercapto-lysine residue is first incorporated into a peptide using solid phase peptide synthesis,
where it can then undergo an NLC reaction with a ubiquitin thioester. The δ-thiol group is then
removed by chemical desulfurization. (D) A δ-mercapto-lysine residue can be site specifically
installed into recombinant proteins using unnatural amino acid mutagenesis. (E) Examples of
isopeptide linkages that have been used for the installation of ubiquitin.
Decoding protein ubiquitylation
Visualization and identification of ubiquitin enzymatic machinery has been accomplished
using activity based protein profiling (ABPP), which utilizes enzyme-specific probes that
react, in most cases covalently, within the active site of various enzymes (Nomura et al.,
36
2010). Although the targets of activity-based probes vary, typically they contain two
elements: a reactive group containing an electrophile to react with a nucleophilic residue
within the enzyme, and a tag. For example, a panel of radioactive- or HA-tagged,
DUB-specific probes containing a C-terminal thiol-reactive group or “warhead” was
synthesized using EPL. They were subsequently utilized as suicide substrates to first
visualize and then identify active DUBs from mammalian cells (Borodovsky et al., 2002).
The range of electrophilic warheads was then expanded beyond the vinylmethylester and
vinylethoxysulfone groups used previously to enable the identification of both DUBs and
E3 ligases (Love et al., 2009). These probes used known cysteine-selective electrophiles;
however, an active-site directed probe containing an alkyne as the C-terminal electrophile
was recently demonstrated to also react with active-site cysteines (Ekkebus et al., 2013).
The researchers were surprised to discover that their C-terminally propargylated Ub
(Ub-Prg), originally synthesized for site-specific ubiquitination of peptides, inhibited the
human DUB ubiquitin carboxyl-terminal hydrolase isoenzyme L3 (UCHL3). Confirmed
by X-ray crystallography, it was found that the resulting quaternary vinyl thioether
conferred selectivity towards de-ubiquitinating enzymes. Continued success in this field
with these types of atypical warheads will undoubtedly enable the preparation of ever
more selective probes and potentially extend this technique to the metalloprotease
members of the DUB family.
37
Lipidation
The attachment of long-chain fatty acids to proteins, termed lipidation, is a PTM that
regulates membrane affinity, localization, and trafficking (Figure 1-7A) (Hang and
Linder, 2011). There are many types of lipid modifications and their covalent attachment
to proteins has revealed a complicated network of membranes and lipidated proteins that
are at the center of basic cellular function and human disease. This complexity, combined
with an almost complete lack of appropriate biological reagents (e.g., antibodies), have
increased the pressure to develop specific and sensitive methods to probe their function
and identify modified substrates. Myristoylation and palmitoylation are the two most
common classes of fatty-acylation events that typically occur co- and posttranslationally,
respectively. Myristoylation is characterized by the irreversible, covalent attachment of a
14-carbon fatty acid, myristic acid, to the N-terminus of a substrate protein via an amide
linkage that only occurs in eukaryotes (Hannoush, 2015). N-myristoylation commonly
occurs cotranslationally, although posttranslational myristoylation was observed during
programmed cell-death and occurs due to proteolytic cleavage revealing an N-terminal
glycine within a cryptic myristoylation consensus sequence which can then under go
myristoylation (Martin et al., 2011b). In contrast, S-palmitoylation is the dynamic,
reversible addition of a 16-carbon fatty acid, palmitic acid, to cysteine amino-acid side
chains via a thioester linkage. Targets of this PTM include ion channels, regulatory
enzymes, scaffolding proteins and membrane receptors (Chamberlain and Shipston,
2015). For example, S-palmitoylation of the pro-apoptotic protein BAX regulates its
subsequent targeting to the mitochondrial outer membrane to initiate programmed cell
38
death, and the S-palmitoylation of death receptor Fas regulates its protein expression by
circumventing its degradation through the lysosome (Fröhlich et al., 2014). S-Prenylation
is another class of lipidation that affects about 2% of the proteome in mammals (Resh,
2006a). It is characterized by the irreversible addition of an isoprenoid, either at
15-carbon farnesyl or a 20-carbon geranylgeranyl to one or two C-terminal cysteines of a
protein through a thioether linkage. Substrate proteins of S-prenylation require the
modification for membrane association and subsequent regulation of function (Berndt et
al., 2011; Zhang and Casey, 1996). Much like the other PTMs discussed above, the
development of homogeneous, synthetic lipopeptides and lipidated proteins have
contributed to the biochemical understanding of these modifications. Additionally, the
creation of a range of chemical probes has transformed the ability to track and identify
lipidated proteins from living systems.
Encoding lipidation
In cells, short amino acid sequences encode the recognition elements for lipidation and
are sufficient enough to promote the modification. In fact, a 10-amino acid sequence
transplanted from H-Ras, a known target of S-palmitoylation, to another soluble protein
can promote its modification (Hang and Linder, 2011). However, the purification of these
proteins in sufficient amounts and to homogeneity is still a very difficult task. Synthetic
lipopeptides and proteins have given access to fully functional lipidated proteins and have
allowed for the study of complete functional proteins. Often the rate-limiting step in these
studies is the preparation of lipidated peptides that can be incorporated into larger
39
proteins by techniques like NCL, and accordingly, several different approaches have been
developed (Brunsveld et al., 2006). Early work in this area was carried out by Waldmann
and co-workers who developed elegant protecting group strategies that enabled the
solution-phase synthesis of both base-sensitive palmitate and acid-sensitive farensyl
groups (Schmittberger and Waldmann, 1999; Stöber et al., 1997). Solid-phase peptide
synthesis has clear advantages over solution preparation, but it could not be immediately
applied to lipidated peptides due to the harsh deprotection and cleavage conditions. In an
early effort to circumvent this issue, phenylselenocysteine was introduced into peptides
using solid-phase peptide synthesis and subsequently eliminated under oxidative
conditions to give site-specific dehydroalanine residues (Zhu and van der Donk, 2001).
These residues can then undergo Michael additions with various thiol-containing
nucleophiles, including protected farensyl-thiol. Although this three-step method is
attractive due to its ease and versatility, the lack of diastereoselectivity in the reaction is a
limitation. Alternatively, on-resin lipidation was used where selectively protected
cysteine residues were revealed and alkylated or esterified with farnesyl or palmitoyl
electrophiles, respectively (Ludolph et al., 2002). This approach has also been reversed:
β-bromo-alanine residues were incorporated into a peptide and then displaced by an
appropriate thiol-containing lipid as the nucleophile (Pachamuthu et al., 2005).
Unfortunately, these methods require a significant excess of the lipids. Notably, the
problems with solid-phase peptide synthesis with pre-lipidated amino acids was first
overcome through the use of a hydrazide linker to the solid support, enabling a more
straight-forward cassette approach (Kragol et al., 2004; Lumbierres et al., 2005). More
40
recently, the concept of post-peptide-synthesis modification has been revised using
thiol-ene chemistry (Triola et al., 2008) by selectively reacting cysteine thiols in
unprotected peptides with lipid alkenes for the generation of palmitoylation analogs
(Calce et al., 2014; Wright et al., 2013) and native farnsylated structures (Calce et al.,
2014). Much like the synthesis of other modifications, the generation of analogs of the
lipid linkage has also been explored. For example, the Davis lab created disulfide-linked
lipid modifications by taking advantage of Lawesson’s reagent (LR) for the site-selective
lipidation of cysteine residues in full-length proteins (Gamblin et al., 2008). Specifically,
the cysteine sulfhydryl on the protein is first activated as a phenyl selenenyl sulfide by
treatment with phenylselenenyl bromide, followed by addition of lipid thiols, resulting in
reasonable modification yields of a model protein (geranyl >90% while farensyl >50%).
These peptide synthesis strategies have enabled the generation of lipidated analogs of the
protein NRas that were then microinjected into living cells to directly monitor palmitate
turnover kinetics and interestingly, enabling the localization of depalmitoylation events
throughout the cell, the palmitoylation machinery to the Golgi (Rocks et al., 2010).
41
Figure 1-7. Lipidation. (A) Proteins can be modified by several types of lipids, including
myristoylation at the N-termini and palmitoylation and prenylation at cysteine residues. (B)
Examples of lipid metabolic chemical reporters for the visualization and identification of lipidated
proteins and lipid-dependent protein-protein interactions. (C) Acyl-biotin exchange for the
analysis of palmitoylation. Free cysteine residues are first capped by incubation of cell lysates
with N-ethylmaleimide. Any palmitate thioesters are then cleaved using hydroxylamine and the
resulting free thiols can be reacted with a variety of electrophilic tags.
Decoding lipidation
Classically, protein lipidation has been investigated by treatment of cells with radioactive
lipids (
3
H or
14
C) that are metabolically incorporated by cells. However, visualization of
the modified proteins requires days-long exposure times on radioactive film, and this
technique offers no opportunity for the enrichment or identification of lipidated proteins.
To overcome these challenges, fatty-acid MCRs containing azides or alkynes have been
developed that take advantage of bioorthogonal chemistry in the same way as the
carbohydrate reporters discussed above (Figure 1-7B) (Hang et al., 2011). Specifically,
probes are designed to mimic the hydrocarbon chain length, incorporating the “click-
able” substituent at the omega end, therefore minimizing interference with acyl-CoA
42
recognition and enzymatic catalytic efficiency (Hannoush, 2015). Recently, for example,
Alk-12 (13-tetradecynoic acid) was used in conjunction with a biotin-enrichment tag for
the identification of myristoylated proteins in the malaria pathogen Plasmodium
falciparum, which allowed for the identification of several proteins implicated in the
parasite’s life cycle and disease transmission (Wright et al., 2014). Similarly, Alk-16
(17-octadecyonic acid) has been utilized in the proteomic identification and subsequent
characterization of palmitoylated proteins. For example, global proteomic profiling in a
mouse dendritic cell-line identified 150 potentially palmitoylated proteins, including the
innate immune effector IFITM3, which the authors subsequently demonstrated requires
fatty-acylation for its anti-viral activity (Yount et al., 2010). Another proteomics study
utilized Alk-16 as well as Alk-12 and Alk-14 (i.e., Alk-12 for myristoylation and Alk-16
for palmitoylation), to highlight the selectivity of the chain length for different
modifications (Wilson et al., 2011). The chemical reporter Alk-16 has also been used in
conjunction with stable isotope labeling in cell culture (SILAC) to perform quantitative
proteomics and subsequently demonstrated that some palmitoylation events are stable
over time while others are more dynamic (Martin et al., 2011a). More recently, a diazirine
photo-cross-linking functionality was incorportated into Alk-16, which enabled the
identification of protein-binding partners of lipidated IFITM3 (Figure 1-7B) (Peng and
Hang, 2015) An alternative strategy takes advantage of the thioester linkage of
S-palmitoylation to use chemoselective reactions to perform a “biotin switch”, a method
termed acyl-biotin exchange (ABE) (Figure 1-7C) (Drisdel and Green, 2004).
Specifically, free cysteines in a cell lysate are alkylated with N-ethylmaleimide, followed
43
by treatment with hydroxylamine, which acts to cleave any palmitate thioesters revealing
sulfhydryl groups that are then selectively modified with biotinylation reagents.
Importantly, this method was used for the identification of palmitoylated proteins in yeast
(Roth et al., 2006), mammalian cells (Ivaldi et al., 2012), malaria parasites (Jones et al.,
2012), and mouse tissue (Wan et al., 2013). S-prenylation has also been investigated
using alkyne derivatives of isoprenoids (Charron et al., 2011; DeGraw et al., 2010). For
example, alkyne-farnesol was utilized in a large-scale enrichment of isoprenoid-modified
proteins in a macrophage cell-line that led to the identification of both known and
unpredicted S-prenylated proteins, including the zinc-finger antiviral protein (ZAP)
(Charron et al., 2013).
Other probes developed to investigate palmitoylation machinery take advantage of some
irreversible pan palmitoylation inhibitors. For example, alkyne analogs of the inhibitor
2-bromopalmitate (2-BP) were utilized as activity-based probes for protein palmitoyl
acyltransferases (PATs) as well as other palmitoylating and 2-BP-binding enzymes
(Zheng et al., 2013). Specifically, the corresponding terminal alkyne analogs of
1,2-bromohexadec-15-yonic acid (16C-BYA) and 2-bromohexadec-17-ynoic acid
(18C-BYA) were synthesized and subsequently used to identify endogenous proteins in
HEK293A and the pancreatic cancer cell line PANC1. While 18C-BYA was able to
identify three endogenous PATs from HEK293A cells, other acyltransferases and
acyl-CoA enzymes were also enriched, as well as many known palmitoylated substrates,
raising the possibility that 2-BP could be incorporated into the cellular lipid pool and
44
used as an acyl donor during palmitoylation. Improving on this study, the same authors
developed a clickable analog of the natural product cerulenin, an inhibitor of fatty acid
biosynthesis and protein palmitoylation that acts through irreversible alkylation of the
cysteine residues in the enzymes (Zheng et al., 2014). The cerulenin-derived probe was
demonstrated to be more specific than the first generation 2-BP probe, perhaps because it
does not require metabolic transformation within the cell. The Tate lab utilized a potent
and specific human N-myristoyltransferase (NMT) inhibitor in combination with SILAC
and an alkyne-containing myristate analog to identify potentially myristoylated proteins
(Thinon et al., 2014). In brief, cells were treated with tetradec-13-ynoic acid (YnMyr)
(Heal et al., 2008) and were either grown in standard media containing NMT inhibitor or
in SILAC media with no inhibition. Following enrichment using an azido-biotin affinity
tag, substrates (and non-substrates) were assigned according to the response in
enrichment to the inhibition of NMT. Using this method, over 100 N-myristoylated
proteins were identified including novel targets such as nucleolar protein 3 (NOL3/ARC),
a protein implicated in inhibition of apoptosis, tumorigenesis, metastasis and
chemoresistance (Thinon et al., 2014).
Identification of Lipidated Proteins
Classically, fatty-acylation has been studied using metabolic radiolabeling (
4
H or
14
C);
however, radioactivity usually requires days to weeks to visualize modified proteins and
offers no opportunities for their selective enrichment and identification (Resh, 2006b). To
overcome these limitations, azide and alkyne fatty-acids have been developed (Hang et
45
al., 2011). Recently, Alk-12 (13-tetradecynoic acid, Figure 4A) was used in combination
with a biotin-enrichment tag to identify mrysitoylated proteins in the malaria pathogen P .
falciparum, including several proteins that are important for the parasite’s life cycle and
disease transmission (Wright et al., 2014). In a similar fashion, 17-octadecynoic acid
(Alk-16, Figure 4B) has been particularly useful for the proteomic identification and
subsequent characterization of palmitoylated proteins. For example, Alk-16 enabled
global proteomic profiling in a mouse dendritic cell-line and identified 150 potentially
palmitoylated proteins. This included several new palmitoylation substrates, such as the
innate immune effector IFITM3, which the authors demonstrated requires fatty-acylation
for its anti-viral activity (Yount et al., 2010). Subsequent proteomic-profiling of Jurkat T
cells with Alk-16 and two other alkyne fatty-acids (Alk-12 and Alk-14) highlighted the
selectivity of different length alkyne fatty-acids for different modifications (i.e., Alk-12
for myristoylation and Alk-16 for palmitoylation) (Wilson et al., 2011). More recently,
Alk-16 was combined with stable isotope labeling in cell culture (SILAC) to perform
quantitative proteomics and demonstrated that some palmitoylation events are very stable
over time, while others are rapidly removed from substrate proteins (Martin et al., 2011a).
As an alternative to bioorthogonal chemistry, the thioester linkage of S-palmitoylation
enables the use of chemoselective reactions to perform a “biotin switch” (Drisdel and
Green, 2004). This method, termed acyl-biotin exchange (ABE, Figure 4C), involves the
alkylation of free cysteine residues with N-ethylmaleimide, followed by treatment with
hydroxylamine, which cleaves any palmitate thioesters revealing sulfhydryl groups that
can be selectively modified with biotinylation reagents. Most notably, this method was
46
used for the identification of palmitoylated proteins in yeast (Roth et al., 2006) and then
in neurons (Kang et al., 2008), including a palmitoylated Cdc42-isoform that plays a
important role in the induction of dendritic spines (Kang et al., 2008). Recently, this
approach was also used to identify palmitoylation candidates in B lymphocytes (Ivaldi et
al., 2012) in malaria parasites (Jones et al., 2012) and in tissue from a mouse model of
Huntington’s disease (Wan et al., 2013).
S-Prenylation is an irreversible addition of an isoprenoid, either 15-carbon
farnesyl or 20-carbon geranylgeranyl to one or two C-terminal cysteines of a protein
through a thioether linkage that effects 2% of the proteome in mammals (Resh, 2006a).
Typically, these modifications are found near the C-terminus of proteins where they are
critical for localization to the cell membrane (Resh, 2006a). Like fatty-acids, alkyne
derivatives of isoprenoids have also been created (Charron et al., 2011; DeGraw et al.,
2010). Recently, an alkyne-farnesol (Figure 4D) permitted a large-scale enrichment of
isoprenoid-modified proteins in a macrophage cell-line that identified both known and
unpredicted S-prenylated proteins (Charron et al., 2013). Notably, these proteomic data
served as a starting point to demonstrate that S-farensylation is important for the
localization and antiviral activity of zinc-finger antiviral protein (ZAP).
Acetylation
Protein acetylation describes the reversible posttranslational transfer of an acetyl group
from acetyl-coenzymeA (Acetyl-CoA) to a target protein, most commonly on the
side-chain amine of a lysine residue (Figure 1-8A). This process is catalyzed by lysine
47
acetyltransferases (KATs) and is removed by lysine deacetylases (KDACs) (Yu-Ying et
al., 2011). KDACs can be broken into two families: classic metallohydrolases that act
directly to hydrolyze acetimide and the sirtuins that utilize an nitotinamide adenine
dinucleotide (NAD) cofactor (Dancy et al., 2012). Protein acetylation was first identified
on lysine-rich N-terminal tails of histones isolated from calf thymus in 1963
{Phillips:1963uh}. Approximately one year later, Vincent Allfrey and colleagues showed
that radiolabelled acetate was rapidly sequestered from media and incorporated onto
histones of isolated nuclei while not being affected by treatment with puromycin, a
translation inhibitor, suggesting that the acetylation events take place posttranslationally.
Additionally, Allfrey was able to show that histone acetylation decreased inhibition of
RNA synthesis, giving rise to the widely accepted theory that posttranslational histone
acetylation serves as a dynamic and reversible mechanism for the regulation of
transcription (Allfrey et al., 1964). Since Allfrey had made his initial discovery, his
theory has been thoroughly validated and it is currently accepted that lysine acetylation,
in particular that of histones, plays a large role in regulating epigenetic changes through
both gene transcription (Shahbazian and Grunstein, 2007) and non-chromatin associated
proteins (Yang et al., 2010). Given that changes in transcriptional regulation are a feature
of human diseases, most notably cancer, there has been significant interest in the
modulation of protein acetylation levels as a therapeutic strategy. For example V orinostat,
a histone deacetylase (HDAC) inhibitor, is the first FDA-approved drug for acetylation
regulation for the treatment of T cell lymphoma, and V orinostat and two other HDAC
inhibitors, romidepsin and panabinostat, are currently going through clinical trials for the
48
treatment of other cancers, as well as HIV infection (Shirakawa et al., 2013). Despite
these successes and the creation of several anti-acetylation antibodies, deciphering the
effects of the thousands of known human acetylation sites remains an obstacle, and a
variety of chemical tools have been developed to help move the field forward.
Encoding acetylation
Examination of site-specific acetylation events is needed to fully understand the
biological implications associated with the modification. In contrast to the modifications
discussed above, the incorporation of acetylated lysine residues into peptides using solid
phase peptide synthesis is quite straight forward. Lysine residues directly bearing an ε-N-
acetate group can be used without the need for further protection and then these peptides
can readily participate in NCL reactions for the preparation of synthetic proteins
{He:2003eq}. For example, NCL was used to generate histone H4 site specifically
acetylated at K16 {ShogrenKnaak:2006gt}. Subsequent biochemical analysis showed that
acetylation played an integral role in chromatin compaction through inhibition of
cross-fiber interactions. They also found that this acetylation event inhibits
ATP-dependent chromatin assembly and remodeling enzyme (ACF) from mobilizing the
mononucleosome, suggesting that acetylation of K16 is sufficient to regulate both higher
order chromatin structure as well as protein-chromatin interactions in a manner that
affects its function {ShogrenKnaak:2006gt}. In addition to NCL-based approaches,
acetyl-lysine has also been incorporated into recombinant proteins using unnatural amino
acid mutagenesis (Figure 1-8B) (Neumann et al., 2009; 2008). The M. bakeri pyrrolysyl
49
tRNA synthetase/tRNA CUA was once again used to incorporate acetylated lysine at amber
stop codons in E. coli expressed proteins for biochemical experiments. For example,
acetyl-lysine was site-specifically introduced at residue 56 in histone H3, and
single-molecule FRET experiments were used to show that it does not have a direct effect
on the compaction of chromatin (Neumann et al., 2009). Finally, an analog of
acetyl-lysine has also been encoded through modification of cysteine residues (Figure
1-8C) (Huang et al., 2010). Specifically, Cole and co-workers demonstrate that cysteine
residues in peptides and proteins can be selectively alkylated in high yield with
methylthiocarbonyl-aziridine to generate a thiocarbamate analog of acetyl-lysine.
Importantly, this analog is recognized by both antibodies and an acetyl-lysine-binding
bromodomains, and the authors confirmed its ability to mimic the natural modification in
the activation of two full-length proteins. Notably, this analog is stable to enzymatic
deacetylation, potentially enabling the specific effects of an acetylation mark to be tested
in cell lysates or through microinjection.
50
Figure 1-8. Acetylation. (A) The most common form of protein acetylation is the dynamic
modification of lysine side chains. (B) Acetylated lysine residues can be incorporated into
recombinant proteins using unnatural amino acid mutagenesis. (C) Analogs of lysine acetylation
can be installed onto cysteine residues using alkylation chemistry, resulting in stable
thiocarbamate structures. (D) Chemical reporters of acetylation, malonylation, and
aspirin-dependent acetylation.
Decoding acetylation
Although anti-acetyl-lysine antibodies exist, chemical reporters of acetylation have
certain advantages, including reporting on non-lysine acetylation events and robust
recovery in proteomics experiments. Towards this goal, sodium 4-pentynoate has been
developed as an MCR lysine acetylation both in living cells and in vitro (Figure 1-8D).
Treatment of living cells with 4-pentynoate or cell lysates with chemically synthesized
4-pentynoyl-CoA enabled the visualization and identification of both known and new
acetylation targets (Yang et al., 2010). Subsequently, 4-pentynoyl-CoA was incubated
with the acetyltransferase p300 to identify enzyme specific substrates. The newly
discovered acylation substrates included a cysteine residue in histone H3 variants (Wilson
51
et al., 2011), demonstrating that these reporters can be used to visualize acylation events
that would not be picked-up by traditional anti-acetyl lysine antibodies. Additional
acylation events with diverse structures on lysine residues have also begun to emerge. To
further probe these understudied modifications, an alkyne-bearing MCR of lysine
malonylation was synthesized as a protected version of 2-propargyl malonate, termed
Mal-AMyne (Figure 1-8D) (Bao et al., 2013). Treatment of HeLa cells with Mal-AMyne
resulted in the identification of 14 previously known malonylated proteins as well as 361
new potential substrates (Bao et al., 2013). Another form of acylation that occurs is a
non-enzymatic transfer of acetate from small-molecules, such as aspirin, to protein
substrates. Our lab introduced a chemical reporter of aspirin acetylation, AspAlk (Figure
1-8D) that enabled the visualization of these chemical events and identification of 120
potential substrate proteins including some of the core histones (Bateman et al., 2013b).
More recently, this same probe was combined with quantitative proteomics to identify
523 proteins and many specific sites of aspirin-mediated acetylation (Wang et al., 2015).
Identification of Acetylated Proteins
Proteome-wide studies using anti-acetyl lysine antibodies have unveiled ~4700 human
acetylation sites, making the investigation of the dynamics of these modifications and the
substrate specificity of acetyltransferases and deacetylases an important goal. An alkyne
analog of acetate, 4-pentynoic acid, has been shown to be chemical reporter of lysine
acetylation in living cells and in vitro. Treatment of a variety of cell lines with sodium
4-pentynoate resulted in metabolic labeling of a wide range of proteins, enabling the
52
proteomic identification of both known lysine-acetylated proteins, like histones, and new
potential substrates (Yang et al., 2010). Additionally, synthetically prepared
4-pentynoyl-CoA (Figure 5A) is a substrate for purified p300 acetyltransferase, enabling
the identification of transferase-specific substrates from cell lysates (Yu-Ying et al.,
2011). Acetylation can also occur on the N-terminus of proteins and serine and threonine
side-chains through an ester linkage. This raises the possibility that these chemical
reporters could be used to investigate these acetylation events that would not be
recognized by anti-acetyl lysine antibodies.
Other types of lysine acetylation events, including propinylation, butyrylation,
malonylation and succinylation, can also occur {Lin:2012ic}. While it is not completely
clear if these modifications are a result of enzymatic addition or simply chemical
acetylation from their corresponding high-energy CoA thioesters, several of them are
reversible through action of the sirtuin deacylases. Towards applying chemical reporters
to the investigation of these modifications, a alkyne-analog of malonylation, 2-propargyl
malonate (Mal-yne, Figure 5B) has recently been developed (Bao et al., 2013). Notably,
due to the low cell-permeability of Mal-yne, the small molecule was masked as a
pro-reporter through the introduction of acetoxymethyl protecting groups that are
removed by endogenous lipases in living cells. Treatment of HeLa cells with this
pro-reporter (Mal-AMyne) enabled the identification of 375 potential substrates,
including 14 previously confirmed malonylated proteins. Some small-molecules contain
electrophilic acetates that can be chemically transferred to protein substrates. For
example, aspirin (acetylsalicylic acid), a nonsteroidal anti-inflammatory drug (NSAID)
53
commonly used for the treatment pain, is known to acetylate proteins by transfer of its
acetate group to amino acid side-chains {Alfonso:2009ij}. Recently, aspirin-dependent
acetylation was investigated using an alkyne-aspirin chemical reporter (AspAlk, Figure
5C) that replaced the electrophilic acetate-group with 4-pentynoic acid (Bateman et al.,
2013b). Treatment of HCT-15 cells with AspAlk allowed for the identification of 120
potentially aspirin acetylated proteins including core histone proteins H2B and H3,
implicating aspirin as a potential modulator of gene transcription.
Methylation
Protein methylation describes the transfer of a methyl group onto the side chains of
lysines, arginines, and less commonly histidines (Figure 1-9A) (Lee et al., 2005). Lysine
residues can be mono-, di- or tri-methylated, while arginines can mono-methylated or
di-methylated. Histidines have only been reported to be mono-methylated, however, this
modification is uncommon and its role remains unclear (Greer and Shi, 2012). Histone
methylation plays well-documented roles in transcriptional regulation, and non-histone
protein methylation, although historically less characterized, has emerged as a prevalent
PTM that plays an important role in cellular signaling. The installation of
posttranslational methyl modifications onto proteins is catalyzed by a class of enzymes
called methyltransferases and is removed by demethylases. Methyltransferases generally
use S-adenosylmethionine (SAM) as a methyl donor (Biggar and Li, 2015).
Demethylases use a FAD cofactor and molecular O
2 to produce formaldehyde, hydrogen
peroxide, and the demethylated peptide (Dancy et al., 2012). Protein methylation on
54
lysine was first recognized to occur posttranslationally in 1965 (Kim and Paik, 1965),
shortly after methylated lysine was found in bacterial flagellar protein {Ambler:1959vz}
and on the histones isolated from calf thymus, wheat germ, and multiple rat organs
{Murray:1964vx}. Since then, protein methylation has been implicated in a variety of
cellular functions including cell-cycle regulation, DNA damage and stress response, and
the development and differentiation through modulation of chromatin-bound histones as
well as chromatin-associating non-histone proteins (Huang and Berger, 2008;
Kouzarides, 2007). Not surprisingly, the misregulation of protein methylation has been
linked to a variety of diseases including cancer (Chi et al., 2010), intellectual disability
{Iwase:2010bv}, and aging {Scaffidi:2006ck}. Antibodies that recognize the different
methylation states of both lysine and arginine are available, as well as small molecule
inhibitors of methyltransferases and demethylases. These tools have contributed greatly
to the investigation of protein methylation, but these methods have limitations for the
analysis of specific methylation events. Fortunately, chemical approaches for the
preparation of site-specifically modified proteins, as well as probes that can deconvolute
the substrate specificity of methyltransferases have both contributed to our understanding
of this key modification.
Encoding methylation
Like acetylation, methylated lysine residues can be easily incorporated into peptides
using solid phase peptide synthesis; di-methyl and tri-methyl lysines can be directly
incorporated, and the mono-methylated side chain can be protected as an appropriate
55
carbamate. NCL has been frequently used for incorporating methylated lysine with native
linkages into proteins {He:2003eq}. In an excellent example of the power of NCL, a
mononucleosomes bearing either a di- or tri-methylated lysine (K7) histone H3 and a
series of acetylated lysine residues in histone H4 were prepared (Ruthenburg et al., 2011).
Using these homogenous proteins, the authors were able to demonstrate the cross-talk
that occurs between the two PTMs by showing a significant increase in binding by the
BPTF PHD-Bromodomain in response to a specific pattern of both types of modification.
In another example, NCL was used to generate histone H3 with tri-methylated lysines at
residues 4, 9, and 27 (Bartke et al., 2010). This semi-synthetic methylated H3 was
combined with biotinylated DNA and immobilized on streptavidin beads to enrich for
binding partners of H3 that are specific for the tri-methylation modifications. As a
complementary strategy, unnatural amino acid mutagenesis using the M. bakeri
pyrrolysyl tRNA synthetase/tRNA
CUA has also been used to incorporate mono-methylated
lysine residues (Figure 1-9B). A pyrrolysyl-tRNA synthetase pair has been used
extensively for incorporation of mono- and dimethylated lysine into recombinant proteins
as it prefers methylated lysine over unmodified lysine. In one example, mono- and
dimethylated Lys were incorporated into recombinant proteins using a PylRs-tRNA pair.
Here, mono-methylated lysine was installed as a tert-butoxycarbonyl protected analog,
which was then subsequently deprotected using TFA {Nguyen:2009wp}. Similar
two-step genetic approaches have been used for the installation of site-specifically
modified lysine using alternative protection groups including allylcarbamoyl and
photocaged methylated lysine that allow for alternative deprotection methods (Groff et
56
al., 2010; Wang et al., 2010). The Chin lab has also used a GOPAL-like strategy to reveal
a specific lysine residue that can be di-methylated using reductive methylation (Nguyen
et al., 2010). Unnatural amino acid mutagenesis methods are yet to be used for the direct
incorporation of di- and tri-methylated lysine residues. However, a chemical method for
the facile incorporation of mono-, di, and tri-methylated lysine analogs was developed
(Figure 1-9C) (Simon et al., 2007). Briefly, cysteine resides at the desired site of
modification can be selectively alkylated it to create a methyl lysine analog (MLA),
N-methylated aminoethylcysteine. Notably, these mono-, di-, and tri-methylated lysine
analogs were functionally similar to their native counterparts when incorporated into
recombinant proteins.
Decoding methylation
Linking a specific methyltransferase to a particular substrate is key to understanding the
complex cellular biology of this modification. Working to identify and analyze the protein
methylome, two alkyne-analogs of SAM were developed as the first chemical reporters of
protein methylation. Several endogenous methyltransferases accepted these stable alkyne
analogs of SAM as cofactors and were able to transfer an alkylated methyl group onto a
lysine residue of both a peptide and a recombinant protein. The alkylated methyl group
could then subsequently be subjected to labeling with CuAAC using an azide tag (Peters
et al., 2010). Interestingly, the alkyne SAM analogs were utilized selectively by different
methyltransferases, suggesting that a chemical reporter/methyltransferase pair could be
created in the same way that selective reporters of phosphorylation were developed as
57
described above. Towards this goal, an additional series of azide- and alkyne-SAM
analogs of various sizes were prepared (Luo, 2012). Several of the developed SAM
analogs were not turned over by wild-type methyltransferases but were selective for
rationally engineered methyltransferase mutants using the bump-hole strategy, enabling
the identification of the specific methyltransferase-dependent substrates (Figure 1-9D)
(Islam et al., 2013). Although this strategy was useful for in vitro testing and the
screening of cellular extract, the poor cell-permeability of SAM analogs limited their use
in experiments involving living cells. To overcome this, the biosynthetic pathway for
SAM was engineered in mammalian cells (Wang et al., 2013b). Briefly, cells can be
treated with cell-permeable alkyne-methionine analogs that will be enzymatically
transformed to the corresponding SAM analogs. Due to the physiological instability of
these methionine-based SAM analogs, a more stable selenium-based SAM reporter was
also created that can probe both arginine and methyltransferases in vitro (Willnow et al.,
2012). Continuing the work with selenium-based reporters, it was shown that a
propargylic Se-containing SAM analog could be used by endogenous methyltransferases
and was stable in whole-cell lysates (Bothwell et al., 2012).
58
Figure 1-9. Methylation and ADP-ribosylation. (A) A variety of methylation marks can be
dynamically installed onto both lysine and arginine side chains. (B) Mono-methylation can be
incorporated into recombinant proteins using unnatural amino acid mutagenesis followed by
acid-based deprotection. (C) Mono-, di-, and tri-methylated lysine analogs can be generated by
alkylation of cysteine residues with the appropriate ethyl-amino electrophile. (D) Using a
bump-hole approach, the alkyne-bearing SAM analog will be transferred by engineered
methyltransferases, enabling the identification of transferase-specific substrates. (E) ADP-ribose
can be enzymatically added to a variety of protein side chains and then subsequently polymerized
to form long poly-ADP-ribose chains. (F) ADP-ribose analogs can be installed onto peptides after
solid phase peptide synthesis by taking advantage of oxime chemistry. (G) Examples of
ADP-ribosylation reporters for use in cell lysates.
59
Identification of Methylated Proteins
The methyl donor for the methyl transferase reaction is the cofactor
S-adenosyl-L-methionine (SAM, also referred to as AdoMet). The first chemical reporters
of protein methylation were two alkyne-analogs of SAM (e.g., AdoEnYn in Figure 6A)
(Peters et al., 2010). Notably, these two chemical reporters displayed selective turnover
by different methyltransferases, raising the possibility that orthogonal
chemical-reporter/transferase pairs could be developed to interrogate substrate specificity.
In this regard, a series of differentially-sized azide- and alkyne-analogs of SAM have
been characterized (Luo, 2012). Notably, several of these compounds were not substrates
for wild-type methyltransferases but were utilized by rationally-engineered
methyltransferase mutants, following a “bump-hole” strategy. This enabled the selective
identification of >500 methylation substrates of the closely related lysine
methyltransferases EuHMT1/2 (Islam et al., 2013). Unfortunately, the poor permeability
of the SAM analogs restricted their use in live cells. To overcome this limitation, the
SAM biosynthetic pathway has been recently engineered (Wang et al., 2013b).
Specifically, an alkyne-analog of SAM can be generated in living cells that express a
mutant SAM synthetase when they are treated with the corresponding cell-permeable
alkyne-analog of methionine. Although the aforementioned methionine-based SAM
reporters are functional, they are subject to degradation at physiological pH and result in
nonspecific labeling. To overcome this, a more stable selenium-based SAM reporter was
developed that can be utilized by both lysine and arginine methyl transferases in vitro
(Willnow et al., 2012). It was subsequently demonstrated that a propargylic,
60
selenium-containing SAM analog (ProSeAM) was compatible with native PMTs in
whole-cell lysate, enabling the identification of 297 potentially methylated proteins, many
of which were novel (Bothwell et al., 2012). Combining all of these tools should facilitate
the identification of specific methyltransferase substrates in living cells in the future.
ADP-Ribosylation
Protein adenosine diphosphate (ADP) ribosylation describes the posttranslational transfer
of an ADP-ribose moiety from β-nicotinamide adenine dinucleotide (NAD
+
)

to a variety
of amino acid side chains on protein acceptors, including aspartate, glutamate, lysine,
arginine and cysteine (Figure 1-9E) (Daniels et al., 2015; Leung, 2014). This
mono-ADP-ribosylation is then often polymerized to generate a long chain of repeating
units, termed poly-ADP-ribosylation. In humans, ADP-ribosylation is installed by a
family of 17 diphtheria toxin-like ADP-ribosyltransferases (ARTDs), commonly known
as poly-ADP-ribose polymerases (PARPs). Majority of these enzymes catalyze
mono-ADP-ribosylation, and four PARPs (PARP1, 2, 5a/b) are known to catalyze
poly-ADP-ribosylation through the transfer of multiple ADPr units onto target proteins
(Carter-O’Connell et al., 2014; Morgan and Cohen, 2015). ADP-ribosylation can be
removed by endogenous enzymes that cleave poly-ADP-ribose polymers such as
poly(ADP-)ribose glycohydrolase (Moyle and Muir, 2010). With NAD
+
serving as a
substrate for ADP-ribosylation, NAD
+
consumption and energy metabolism is directly
linked to the production of ADP-ribose derivatives (Schreiber et al., 2006). Like many
PTMs, ADP-ribosylation plays a unique role in many important cellular processes
61
including but not limited to stress signaling, DNA damage repair, telomere homeostasis,
transcriptional regulation, and centrosomal targeting (Carter-O’Connell et al., 2014;
Schreiber et al., 2006). ADP-ribosylation has also been shown to have important
therapeutic consequences in cancers, neurodegenerative diseases, ischemia, and
inflammatory disorders {Curtin:2013us}. The specific function of majority of ribosylation
events, in particular mono-ADP-ribosylation, is not well understood. Obstacles
contributing to difficulties in studying ADP-ribosylation include stability of the
ester-linked ADP-ribose at basic pH, the ability of the modification to be rapidly removed
by endogenous enzymes, lack of commercial antibodies, and overlapping target
specificities among the 17 ARTDs (Carter-O’Connell et al., 2014; Moyle and Muir,
2010). Again, chemical methods have begun to address these limitations through the
site-specific installation of ADP-ribose analogs and the development of chemical probes
for the global and isoform-specific identification of ARTD substrates.
Encoding ADP-ribosylation
As with the more extensively studied modifications above, the preparation of
site-specifically ADP-ribosylated peptides and proteins is key to investigating its
biochemical effects. However, the chemically sensitive nature of the pyrophosphate bond
makes peptide synthesis challenging. In order to circumvent these issues, Filippov and
co-workers installed a selectively protected ribosylated asparagine or glutamine residue
into peptide using solid phase peptide synthesis (van der Heden van Noort et al., 2010).
An alternative approach has also been developed for the generation of ADP-ribosylation
62
analogs (Figure 1-9F) (Moyle and Muir, 2010). More specifically, this method uses
aminooxy-functionalized amino acids for the specific conjugation of ADP-ribose onto
peptides and semi-synthetic proteins, with oxime ligation occurs between the
aminooxy-functionalized amino acid of choice and the anomeric carbon of ribose at pH
4.5. Notably, this mimic of mono-ADP-ribosylated in a H2B-derived peptide was a
substrate for PARP1, producing site-specific-poly-ADP-ribosylated peptide conjugates.
Finally, the same authors used NCL to prepare ADP-ribosylated H2B proteins with
benzophenone cross-linkers and were able to enrich for previously unknown
ADP-binding proteins histone mH2A1.1 and PARP9 (Moyle and Muir, 2010).
Decoding ADP-ribosylation
In an effort to enrich and identify the ADP-ribosylated proteome, the alkyne-containing
chemical reporters 6-alkyne-NAD and 8-alkyne-NAD have been synthesized (Du et al.,
2009; Jiang et al., 2010). Incubation of these analogues with cell lysate and recombinant
PARP1 led to the discovery of 70 potentially new PARP1 protein substrates (Figure
1-9G) (Jiang et al., 2010). Orthogonal NAD variants have also been used in combination
with several engineered PARPs to identify direct substrates of specific members of the
PARP superfamily (Carter-O’Connell et al., 2014). A “bump-hole” strategy was used to
create a mutant ARTD (K903A) that could accept an NAD analog containing an
ethyl-substituent at the C5’ position of the nicotinamide moiety of NAD. Incubation of
these orthogonal pairs in nuclear extracts resulted in the proteomic identification of a pool
of substrates specific to either PARP1 or PARP2. Affinity purification and tandem mass
63
spectrometry allowed for identification of unique targets for both enzymes, which could
be further applied to all 17 members of the PARP superfamily to delineate the role of
specific enzyme-substrate interactions. More recently, an aminooxy alkyne (AO-alkyne)
probe was synthesized to detect mono-ADP-ribosylation in cells using CuAAC and an
azide-containing tag. The probe was used to show that PARP10 and PARP11 are
auto-ADP-ribosylated as well as used to monitor stimulus-induced ADP-ribosylation in
cells (Morgan and Cohen, 2015).
Identification of ADP-Ribosylated Proteins
Although the role of ADP-ribosylation has been characterized in DNA repair, other
biological functions are relatively unknown and isolation of the ADP-ribosylated
proteome would prove useful for their elucidation. Towards this goal, 6- and
8-alkyne-NAD analogs have been developed (Figure 6B) (Du et al., 2009; Jiang et al.,
2010). Incubation of cell lysates with 6-alkyne-NAD and recombinant ARTD1 (PARP1)
enabled the identification of 70 potentially novel ADP-ribosylated proteins including
many mitochondrial proteins (Jiang et al., 2010). Very recently, orthogonal
6-alkyne-NAD/ARTD pairs have been engineered to identify the direct substrates of
specific ARTDs (Carter-O’Connell et al., 2014). Using a “bump-hole” strategy, a
ethyl-substituent was added to the C-5 position on the nicotinamide moiety of NAD,
which could only be accepted by a corresponding mutant ARTD (K903A). Treatment of
nuclear extracts with these orthogonal pairs made possible the proteomic identification of
64
hundreds of specific substrates of either ARTD1 or ARTD2, including 42 proteins that
were unique ARTD1 substrates.
Conclusions and future outlook
Chemical techniques have been particularly valuable for the synthesis and identification
of posttranslationally modified proteins. Advances in protein-peptide ligations for the
semisynthesis of modified target proteins has enabled the development of unique and
site-specific chemical reactions for the installment of modifications through either native
or nonnative linkages, further expanding our toolbox. For example, in a tour-de-force of
NCL mediated synthesis, the Muir lab has recently prepared histones with different
combinations of ubiquitination, acetylation, and methylation marks (Nguyen et al., 2014).
These proteins can be combinatorially combined to form mononucleosomes with
different modification patterns resulting in a PTM library, which paired with DNA
sequencing techniques can rapidly identify specific protein binding partners for different
patterns of modifications. In parallel, improvements in the quality of chemical reporters
and mass spectrometry methods have allowed for an unprecedented volume of identified
proteins and coupled with chemical tools developed for the enrichment of modified
proteins, has resulted in new protein targets for research and drug discovery. Specifically,
the application of chemical reporters for the incorporation of bioorthogonal chemical
moieties has enabled the study of a variety of posttranslational modifications, that due to
their chemical structure and complex regulation, have been difficult to study with
traditional biological methods. For example, Hang and co-workers used the Alk-16 MCR
65
to show that changes in the palmitoylation of specific proteins controls the entry into
meiosis in fission yeast, suggesting that single enzymes that install PTMs can control
complex biological processes {Zhang:2013ev}.
Unfortunately, despite these successes, challenges still remain. For example, in some
cases, the use of SPPS for the synthesis of modified peptides relies on the chemical
synthesis of complex, pre-modified amino acids that require unique protecting group
strategies, limiting its use to labs with a requisite chemical expertise. Furthermore, NCL
suffers from concentration dependent reaction rates, which limits its utility with folded
protein substrates that cannot be concentrated to reasonable levels. Therefore, there is a
need for the continued collaboration between chemists and biologists to devise
recombinant strategies for the preparation of evermore challenging protein targets. In the
case of chemical reporters, treatment with metabolic analogs may perturb the cellular
environment, causing unnatural changes in cellular metabolism and altering metabolic
pathways. This highlights the need for a more comprehensive investigation, including
understanding the cellular fate of these analogs to uncover which PTMs are being
enriched with a specific reagent. Beyond the identification of modified substrate proteins,
retrieving site-specific information still remains a challenge, as many PTMs are not stable
during MS/MS analysis. These shortcomings should be met with a research effort that
focuses on enhanced mass spectrometry methods, including new ionization techniques
and computer programs that allow for the direct identification of modification sites.
Chemical methods have been instrumental in the investigation of PTMs towards
66
elucidating their biological role, and we believe that certain modifications, such as
arginine methylation, are particularly ripe for the development of additional chemical
tools. The exciting advancements in this field have enabled the use of multiple chemical
methods to both synthesize homogenous proteins for study while unambiguously
identifying their modification status, and undoubtedly, chemical biologists will continue
to have a huge impact in the field of protein modifications.
67
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Chapter 2. Changes in Metabolic Chemical-Reporter Structure Yield a
Selective Probe of O-GlcNAc Modification
*
Introduction
There are three common types of protein glycosylation that modify large numbers of
protein substrates in mammalian cells. Proteins localized to the secretory pathway and the
cell surface or secreted into the extracellular space can be modified by oligosaccharide
structures, such as N-linked glycosylation (linked through asparagine) or mucin O-linked
glycosylation (linked through serine and threonine). Additionally, cytoplasmic, nuclear,
and mitochondrial proteins can be substrates for the addition of the single
monosaccharide N-acetyl-glucosamine, termed O-GlcNAc modification
(O-GlcNAcylation, linked through serine and threonine) (Holt and Hart, 1986; Love and
Hanover, 2005; Torres and Hart, 1984; Zachara and Hart, 2002). Unlike other forms of
glycosylation, O-GlcNAcylation is dynamic. It is added to protein substrates by one of
three isoforms of O-GlcNAc transferase (OGT) and removed by two isoforms of
O-GlcNAcase (OGA) (V ocadlo, 2012). The expression of these enzymes is also required
for embryonic development in mice and Drosophila (Shafi et al., 2000; Sinclair et al.,
2009; Yang et al., 2012). O-GlcNAc modification displays significant crosstalk with other
posttranslational modifications (PTMs), most significantly phosphorylation and
ubiquitination, setting up O-GlcNAcylation as a key regulator of cellular pathways (Hart
*
Balyn W. Zaro (University of Southern California) contributed to the work presented in this chapter.
103
et al., 2011). A wide variety of proteins have been shown to be O-GlcNAc modified,
including regulators of transcription and translation, cytoskeletal proteins, signaling
proteins, and metabolic enzymes. The specific consequences of most of these
modifications are unknown; however, limited biochemical analyses demonstrate that
O-GlcNAc modification can change protein localization, stability, molecular interactions,
and activity. Critically, O-GlcNAcylation is also misregulated in Alzheimer’s disease and
cancer. For example, in neurodegenerative disorders such as Alzheimer’s disease,
O-GlcNAcylation levels are diminished directly leading to protein aggregation and cell
death (Yuzwa et al., 2012), and we have demonstrated that it likely plays a similar role in
Parkinson’s disease (Marotta et al., 2012). Finally, higher levels of O-GlcNAc
modification are a common feature of many cancers and are necessary for tumorigenesis
and proliferation (Caldwell et al., 2010; Kang et al., 2009; Shi et al., 2010).
To identify and characterize O-GlcNAc modifications, complementary chemical methods
have been developed (Banerjee et al., 2013; Rexach et al., 2008). In general, these
technologies take advantage of bioorthogonal chemistries, such as the copper(I)-catalyzed
azide-alkyne cycloaddition (CuAAC or “click chemistry”, Figure 2-1A) (Best, 2009;
Sletten and Bertozzi, 2011; Tornøe et al., 2002). This reaction relies upon small, abiotic
chemical-reporters (azides and alkynes) that can be selectively reacted with alkyne- and
azido-probes, respectively, for the installation of visualization- and affinity-tags. One of
these methods, initiated by the Bertozzi laboratory, takes advantage of monosaccharide
analogs that directly incorporate azides or alkynes into their structures {Dube:2003vo}.
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These analogs, termed metabolic chemical-reporters (MCRs) (Grammel and Hang, 2013),
are taken up by cells through carbohydrate salvage pathways and subsequently feed into
the biosynthesis of nucleotide sugar-donors for use by glycosyltransferases. For example,
the first O-GlcNAc-targeted MCR, N-azidoacetyl-glucosamine (GlcNAz, Figure 6-1B),
has been used for the visualization and proteomic identification of labeled proteins
(Nandi et al., 2006; V ocadlo et al., 2003; Zaro et al., 2011b). Unlike other methods (e.g.,
Western blotting), MCRs do not necessarily read-out on endogenous levels of
O-GlcNAcylation, as they must compete with GlcNAc in the cell. However, because they
must be metabolically transformed before their incorporation onto proteins, they not only
report on O-GlcNAc modification but also on the integration of upstream metabolic
pathways. Additionally, they can be used much like radioactivity to isolate new
modification events and subsequent rates of removal in pulse and pulse-chase labeling
experiments. Despite the clear utility of this technology, the previous iterations have
limitations. Until recently, GlcNAz and other MCRs were presumed to label only one
type of glycosylation (i.e., GlcNAz treatment results in O-GlcNAcylation labeling), but
several enzymatic pathways exist that can interconvert different monosaccharides, raising
the possibility that MCRs are converted in the same manner (Yarema and Bertozzi,
2001). Upon careful characterization, it was demonstrated that GlcNAz can be readily
transformed to N-azidoacetyl-galactosamine (GalNAz, Figure 2-1B) and vice versa,
resulting in the labeling of both O-GlcNAcylated and mucin O-linked glycosylated
proteins (Banerjee et al., 2010; Boyce et al., 2011; Yarema and Bertozzi, 2001; Zaro et
al., 2011b). Furthermore, we showed that GlcNAz treatment leads to labeling of N-linked
105
glycosylation (Zaro et al., 2011b). This can be overcome using cellular fractionation
(Boyce et al., 2011); however, since we have complete chemical control over the MCR,
we predicted that structural alterations can limit this “off-target” labeling and produce an
O-GlcNAcylation-specific reporter. Indeed, we previously demonstrated that an
alternative MCR, N-pentynyl-glucosamine (GlcNAlk), which contains a larger functional
group at the N-acetyl position, is not converted to the galactosamine derivative and
therefore could not label mucin O-linked glycoproteins (Zaro et al., 2011b).
Unfortunately, GlcNAlk was still incorporated into N-linked glycans, preventing its use
as a completely-selective O-GlcNAcylation reporter.
Figure 2-1. Metabolic chemical reporters (MCRs). (A) Copper(I)-catalyzed azide-alkyne
cycloaddition (CuAAC). (B) Peracetylated MCRs used in this study.
We report here the development and application of the metabolic chemical reporter
(MCR) 6-azido-6-deoxy-N-acetyl-glucosamine (6AzGlcNAc, Figure 2-1B and Scheme
2-1) as a MCR in living cells. Cellular analysis of this MCR using CuAAC and
fluorescent probes demonstrated that, unlike previous reporters, it is highly-selective for
O-GlcNAcylated proteins, allowing for the robust visualization of O-GlcNAc
modifications using in-gel fluorescence scanning. Furthermore, comparative proteomics
106
using 6AzGlcNAc, GlcNAz, and GalNAz confirmed the specificity of 6AzGlcNAc
towards O-GlcNAc modifications. 6AzGlcNAc-labeling resulted in the enrichment of
zero proteins, out of 367, which are annotated to have exclusively extracellular or
lumenal localization. In contrast, GlcNAz and GalNAz identified 9 and 72 such proteins,
respectively. Finally, we also demonstrate that 6AzGlcNAc can bypass an assumed
biosynthetic roadblock by being phosphorylated by the enzyme
phosphoacetylglucosamine mutase.
Scheme 2-1. Synthesis of Ac 46AzGlcNAc (6.1). (a) 4-toluenesulfonyl chloride, pyridine, -20 ℃,
18 h; (b) NaN3, DMF, 50 ℃, 3 d; Ac 2O, pyridine, rt, 16 h, 60% over three steps.
Results
6AzGlcNAc is a robust metabolic chemical-reporter in living cells.
Our previous data using MCRs demonstrated that even small alterations in chemical
structure can have dramatic effects on the distribution of chemical reporters into different
types of glycosylation (Bateman et al., 2013a; Zaro et al., 2011b). Therefore, to find a
specific MCR of O-GlcNAcylation, we synthesized a small panel of O-acetylated
N-acetyl-glucosamine analogs bearing azides at different positions; the acetate
protecting-groups allow diffusion across the cell membrane and are subsequently
107
removed by endogenous lipases/hydrolases. NIH3T3 cells were treated with these
compounds at 200 μM concentrations for 16 hours, followed by lysis, CuAAC with an
alkyne-containing rhodamine dye (alk-rho), and analysis by in-gel fluorescent scanning.
One of these compounds, Ac
36AzGlcNAc (Figure 2-1B), gave a protein-labeling pattern
that was subjectively similar in both intensity and pattern to Ac
4GlcNAz (Figure 2-2A).
Figure 2-2. Ac 36AzGlcNAc labels proteins in living cells. (A) NIH3T3 cells were treated with
Ac4GlcNAz (200 μM), Ac 36AzGlcNAc (200 μM), or DMSO vehicle for 16 h, followed by
CuAAC and analysis by in-gel fluorescence scanning. (B) NIH3T3 cells were treated with
varying concentrations of Ac4GlcNAz or Ac 36AzGlcNAc for 16 h, followed by CuAAC and
analysis by in-gel fluorescence scanning. (C) NIH3T3 cells were treated with Ac36AzGlcNAc
(200 μM), or DMSO vehicle for the times indicated and were tested for toxicity using an MTS
assay. (D) Proteins modified by 6AzGlcNAc were enriched from NIH3T3 cells treated with
Ac36AzGlcNAc (200 μM) or DMSO vehicle using CuAAC with alkyne-azo-biotin and analyzed
by Western blotting.
To further characterize this MCR, NIH3T3 cells were treated with various concentrations
of Ac
36AzGlcNAc or Ac 4GlcNAz for 16 hours before reaction with alk-rho. In-gel
fluorescence scanning revealed labeling of a wide-range of proteins in concentrations as
low as 50 μM and maximal labeling achieved at approximately 200 μM (Figure 2-2B),
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consistent with other MCRs of glycosylation(Bateman et al., 2013a; Zaro et al., 2011b;
2011a). To examine the limits of treatment time with our MCR, the viability of NIH3T3
cells was tested after treatment with 200 μM Ac
36AzGlcNAc for 16 hours or 72 hours
using a cell proliferation assay. Only minimal loss of cell growth/survival was seen even
after 72 hours of treatment (Figure 2-2C). To determine if 6AzGlcNAc could report on
O-GlcNAc modifications, we treated NIH3T3 cells with Ac
36AzGlcNAc (200 μM) for 16
hours. The cells were then lysed and reacted with an alkyne-containing cleavable affinity
tag (alk-azo-biotin, Scheme 2-2) using CuAAC. Labeled proteins were enriched using
streptavidin beads before elution with sodium dithionite. Enriched proteins were then
subjected to Western blotting using antibodies against the known O-GlcNAc modified
proteins NEDD4 (Alfaro et al., 2012; Zaro et al., 2011b), pyruvate kinase (Zaro et al.,
2011b), and nucleoporin 62 (nup62) (Lubas et al., 1995). All three proteins were
selectively enriched using 6AzGlcNAc (Figure 2-2D), showing that the MCR does label
known O-GlcNAcylated proteins.
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Scheme 2-2. Synthesis of alkyne-azo-biotin. (a) propargyl chloride, 0.1 M KOH, EtOH, reflux,
20 h, 28%; (b) i.) NaNO2, methyl-4-amino-benzoate, 6 M HCl, K 2CO 3, H 2O:THF (2:1), 0 °C, 30
min; ii.) rt, 18 h, 90%; (c) NaOH, rt, 24 h, 70%; (d) N-hydroxysuccinimide, N,N’-
dicyclohexylcarbodiimide, THF, rt, 18 h, 56%; (e) EZ-link Amine PEG3-biotin, DMF, rt, 18 h,
31%.
110
Figure 2-3. The GlcNAc salvage pathway. (A) Peracetylated GlcNAc accesses the HBP through
the GlcNAc salvage pathway. Ac4GlcNAz is accepted by these enzymes and is ultimately
transformed into the UDP donor sugar the which is utilized by OGT to modify protein substrates.
(B) 6AzGlcNAc cannot be phosphorylated at the 6-position by GNK. We demonstrate that
6AzGlcNAc can be directly transformed to 6AzGlcNAc-1-phosphate by AGM1 and subsequently
transformed to UDP-6AzGlcNAc by AGX1.
6AzGlcNAc is metabolically incorporated by bypassing GlcNAc-6-kinase.
MCRs are enzymatically transformed into their nucleotide sugar-donors by
monosaccharide salvage pathways. Previous O-GlcNAc MCRs are thought to largely
utilize the GlcNAc salvage pathway (Figure 2-3A) (V ocadlo et al., 2003). The first step of
this pathway is the phosphorylation of MCRs at the 6-position of the carbohydrate ring
by N-acetylglucosamine kinase (GNK). This is followed by enzymatic mutation of the
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phosphate to the 1-position and conversion to the uridine-diphosphate (UDP) sugar donor
by N-acetylglucosamine-phosphate mutase (AGM1) and
uridine-diphosphate-N-acetylglucosamine pyrophosphorylase (AGX1/2), respectively.
Although UDP-6AzGlcNAc is known to be accepted by OGT (Mayer et al., 2011),
6AzGlcNAc cannot be phosphorylated at the 6-position, as we have replaced the
6-hydroxyl functionality with an azide. Therefore, we first took a candidate-based
approach to identify a kinase that could directly phosphorylated 6AzGlcNAc at the
1-position and chose N-acetyl-galactosamine kinase (GalK2), which performs this
reaction on N-acetylgalactosamine (GalNAc) and poorly on GlcNAc
{Pastuszak:1996vw}. To test this possibility, NIH3T3 cells were stably transformed with
five different short-hairpin RNA vectors targeting GalK2 using retroviral infection and
then treated with Ac
36AzGlcNAc (200 μM) for 16 hours. Subsequent CuAAC with
alk-rho and in-gel fluorescent scanning showed no loss of fluorescent signal, despite a
clear reduction of GalK2 mRNA as measured by semi-quantitative RT-PCR (Figure
2-4A), suggesting that GalK2 is not the enzyme responsible for 6AzGlcNAc metabolism.
To confirm this result, we subjected 6AzGlcNAc (2.8, Scheme 2-3A) to in vitro
phosphorylation by recombinant GalK2 (Pouilly et al., 2012). Specifically, GalK2 was
incubated with 40 mM concentrations of GalNAc, GlcNAc, or 6AzGlcNAc and
[
32
P]ɣATP (5 mM). At these elevated substrate-concentrations, GalK2 readily
phosphorylated both GalNAc and GlcNAc but gave no detectable modification of
6AzGlcNAc (Figure 2-4B).
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Figure 2-4. Investigation of 6AzGlcNAc metabolism. (A) Cell-lines with stable knockdown
(shRNA) of galactosamine kinase (GalK2) were treated with 6AzGlcNAc (200 μM) for 16 hours
before visualization by in-gel fluorescence. (B) The indicated monosaccharides (40 mM
concentration) were tested as sub- strates for purified GalK2 in vitro. (C) Proposed mechanism
by-which AGM1 directly phosphorylates 6AzGlcNAc in the presence of GlcNAc-6-phosphate.
(D) Kinetic constants for the enzymatic production of UDP sugar donors from
GlcNAc-1-phosphate and 6AzGlcNAc-1-phosphate by the enzyme UDP-N- acetylhexosamine
pyrophosphorylase (AGX1).
Next, we tested whether phosphoacetylglucosamine mutase (AGM1) could directly
generate 6AzGlcNAc-1-phosphate. AGM1 typically converts GlcNAc-6-phosphate to
GlcNAc-1-phosphate during the biosynthesis of UDP-GlcNAc. As part of its enzymatic
cycle, AGM1 removes the 6-phosphate from substrate sugars, resulting in a
phosphoenzyme intermediate (Nishitani et al., 2006). Therefore, once loaded,
phosphorylated AGM1 might be capable of phosphorylating 6AzGlcNAc. To test this
possibility, human AGM1 was heterologously expressed in E. coli and purified. The
enzyme was then incubated with 6AzGlcNAc (2.25 mM) with or without different
“cofactors” that could generate phosphorylated AGM1, specifically glucose-6-phosphate
or glucose-1,6-bisphosphate or GlcNAc-6-phosphate (all at 1 mM). To isolate any
113
6AzGlcNAc-1-phosphate that had been produced, the enzymatic reactions were first
subjected to copper-free click chemistry with a fluorescein-conjugated cyclooctyne tag.
Fluorescein-labeled compounds (i.e., 6AzGlcNAc and 6AzGlcNAc-1-phosphate) were
then separated from the phosphorylated-cofactors by paper chromatography. Finally, any
fluorescent-spots were eluted and analyzed by mass spectrometry (LC-MS, Figure 2-5).
Incubation of AGM1 with 6AzGlcNAc alone, or with glucose-6-phosphate or
glucose-1,6-bisphosphate, resulted in no detectable formation of
6AzGlcNAc-1-phosphate. However, in the presence of GlcNAc-6-phosphate as a
cofactor, the formation of 6AzGlcNAc-1-phosphate was unambiguously detected. This
demonstrates that direct phosphorylation of 6AzGlcNAc by AGM1 represents one
pathway that circumvents the GNK biosynthetic-roadblock. However, because the
conversion is very low (< 1% conversion to product based on ion-intensities in ESI-MS),
AGM1 may not be the only enzyme that can produce 6AzGlcNAc-1-phosphate in living
cells. We next analyzed the final enzyme in the biosynthetic pathway,
UDP-N-acetylhexosamine pyrophosphorylase (AGX1), by first synthesizing
6AzGlcNAc-1-phosphate (2.16, Scheme 2-3B). Recombinant AGX1 was then incubated
with different concentrations of GlcNAc-1-phosphate or 6AzGlcNAc-1-phosphate and
[
3
H]UTP. Subsequent Michaelis-Menten kinetic analysis demonstrated that
6AzGlcNAc-1-phosphate is a substrate of AGX1, although at a significantly lower
efficiency than GlcNAc-1-phosphate (Figure 2-4D). Taken together, these data suggest
that in living cells 6AzGlcNAc can be directly phosphorylated by AGM1 and enter the
remainder of the GlcNAc salvage pathway to generate UDP-6AzGlcNAc (Figure 2-2).
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Scheme 2-3. Synthesis of 6AzGlcNAc-1-phosphate. (A) Reagents: (a) p-Toluenesulfonyl
chloride, pyridine, -20 °C, 16 h; (b) NaN3, DMF, 50 °C, 3 d, 14% over two steps. (B) Reagents:
(a) benzyl alcohol, concentrated HCl, 75 ℃, 4 h, 35%; (b) p-toluenesulfonyl chloride, pyridine,
-20 ℃, 1 h, 52%; (c) acetic anhydride, pyridine, 3 h, quantitative yield; (d) Pd(OH)2/C (10% Pd),
H2, MeOH, 48 h; (e) i) 5-(ethylthio)-1H-tetrazole, diallyl-N,N’-diisopropylphosphoramidite,
CH2Cl 2, 2 h; ii) m-chloroperoxybenzoic acid, CH 2Cl 2; -78 ℃, 10 min, 74% over 2 steps; (f)
sodium methoxide, MeOH, 1.5 h, 57%; (g) sodium azide, DMF, 48 h, 71%; (h) p-toluenesulfinic
acid sodium salt, tetrakis(triphenylphosphine)-Palladium(0), 4 d, 99%.
115

Extrait d’ion correspondant au composé phosphorylé (isotope le plus intense de l’ion dichargé).
Extraction of the ions corresponding to the phosphorylated compound (most intense isotope of the double-charged ion)
X014864CYC.d
X014865CYC.d
X014956CYC.d
X014957CYC.d
X014958CYC.d
X014959CYC.d
0
20
40
60
Intens.
0
1
2
5
x10
0
50
100
150
200
0
500
1000
0
100
200
0
100
200
0.0 2.5 5.0 7.5 10.0 12.5 15.0 17.5 20.0 Time [min]
Fraction 0.2
Fraction 1.2
Fraction 3.2
Fraction 2.2
Fraction 4.2
Fraction 5.2
6 azidoGlcNAc-Bicyclooctyn-POE3-Fluorescein (control)
6 azidoGlcNAc-1P-Bicyclooctyn-POE3-Fluorescein (control)
PGM3 + 6 azidoGlcNAc
PGM3 + 6 azidoGlcNAc + GlcNAc6P
PGM3 + 6 azidoGlcNAc + GlcP6P
PGM3 + 6 azidoGlcNAc + Glc1,6diP
Figure 2-5. LC-MS analysis of 6AzGlcNAc-1-phosphate production by AGM1. AGM1
enzymatic reactions were subjected to copper-free click chemistry with
bicyclooctyn-POE3-fluorescein and subjected to separation by paper chromatography.
Fluorescent spots were eluted and analyzed using LC-ESI-MS. Ions corresponding to the
fluorescein-conjugated 6AzGlcNAc-1-phosphate (most intense isotope of the double-charged ion)
were extracted (blue trace). Only in the presence of AGM1 (PGM3) and GlcNAc-6-phosphate is
the formation of 6AzGlcNAc-1-phosphate observed. Units on the y-axis are not uniform.
6AzGlcNAc is a general and dynamic metabolic chemical-reporter in living cells.
Next, to explore the generality of 6AzGlcNAc as a MCR, we labeled a panel of different
cell lines. Specifically, Cos-7, H1299, HEK293, HeLa, MCF7, mouse embryonic
fibroblasts (MEFs), and NIH3T3 cells were treated with Ac
36AzGlcNAc (200 μM) for 16
hours. In-gel fluorescence scanning after CuAAC with alk-rho showed labeling in all the
cell lines examined and a diversity of the pattern and intensity of modified proteins
(Figure 2-6). To qualitatively compare 6AzGlcNAc to previous MCRs of
O-GlcNAcylation, the same panel of cell lines was treated with 200 μM Ac
4GlcNAz or
Ac
4GalNAz for 16 hours (Figure 2-6). In-gel fluorescence scanning showed incorporation
116
of previously characterized MCRs in each cell line with varying intensities and patterns,
which were more pronouncedly different for GalNAz, when compared to 6AzGlcNAc
and GlcNAz.
117
Figure 2-6. Fluorescence incorporation of MCRs in a variety of cell lines. The indicated
cell-lines were treated with 200 μM Ac36AzGlcNAc, Ac 4GlcNAz or Ac 4GalNAz for 16 hours
before modified proteins were subjected to CuAAC with alk-rho and analysis by in-gel
fluorescence scanning.
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As stated above, MCRs only report on modifications that occur during the labeling time,
raising the possibility that they can be used to isolate O-GlcNAcylation events in a short
time frame via a pulse-labeling experiment. To determine the kinetics of protein labeling
by 6AzGlcNAc, NIH3T3 cells were treated with Ac
36AzGlcNAc (200 μM) for different
lengths of time. The cells were then lysed, reacted with alk-rho using CuAAC, and
analyzed by in-gel fluorescence scanning (Figure 2-7A). Modified proteins can be clearly
visualized over background in 2 to 4 hours, similar to the kinetics of protein labeling by
Ac
4GlcNAz at 200 μM (Figure 2-7A). MCRs also have the ability to read-out on the
turnover of protein modifications using a pulse-chase format. Accordingly, we treated
NIH3T3 cells with either Ac
36AzGlcNAc or Ac 4GlcNAz at concentrations of 200 μM.
After 16 hours, the cells were washed and fresh media containing Ac
4GlcNAc (200 μM)
was added. Cells were collected after different lengths of time, lysed, and subjected to
CuAAC with alk-rho. In-gel fluorescence scanning showed a steady loss of protein
labeling over the course of 48 hours (Figure 2-7B), in line with radioactive (tritiated
glucosamine) pulse-chase experiments of O-GlcNAc on α-crystallin and cytokeratin (t
1/2
of ~10 and 55 hours, respectively) (Chou et al., 1992; Roquemore et al., 1996).
O-GlcNAcase (OGA) is responsible for the dynamic removal of O-GlcNAc from
substrate proteins. To demonstrate that 6AzGlcNAc is incorporated into
O-GlcNAcylation and a substrate for OGA, cells were first treated with Ac
36AzGlcNAc
(200 μM) or DMSO for 5 hours. Media was then exchanged for fresh media containing
200 μM Ac
4GlcNAc with or without Thiamet-G (10 μM), a potent and highly-selective
OGA inhibitor (Yuzwa et al., 2008). After 12 hours, cells were harvested and subjected to
119
CuAAC with alk-rho. In-gel fluorescence scanning shows that cells that were treated with
Thiamet-G maintain higher levels of 6AzGlcNAc labeling compared to those without
(Figure 2-7C), demonstrating that 6AzGlcNAc is incorporated into O-GlcNAc
modifications that can be subsequently removed by OGA.
Figure 2-7. Characterization of Ac 36AzGlcNAc. (A) NIH3T3 cells were treated with 200 μM
Ac36AzGlcNAc or Ac 4GlcNAz for the indicated times, followed by CuAAC and analysis by
in-gel fluorescence scanning. (B) NIH3T3 cells were treated with 200 μM Ac36AzGlcNAc or
Ac4GlcNAz for 16 h at which time media was exchanged for fresh media containing 200 μM
Ac4GlcNAc. Cells were harvested after the indicated lengths of time, subjected to CuAAC and
analyzed by in-gel fluorescence scanning. (C) HeLa cells were treated with 200 μM
Ac36AzGlcNAc or Ac 4GlcNAz for 16 h at which time media was exchanged for fresh media
containing 200 μM Ac4GlcNAc and 10 μM of the OGA inhibitor Thiamet-G or DMSO. Cells
were harvested at the times indicated and subjected to CuAAC before being analyzed by in-gel
fluorescence scanning.
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6AzGlcNAc is a specific metabolic chemical-reporter of O-GlcNAc modification.
As noted above, previous MCRs of O-GlcNAcylation are not selective for O-GlcNAc
modifications because they are also incorporated into either N-linked or mucin O-linked
glycans or both (Banerjee et al., 2010; Boyce et al., 2011; Zaro et al., 2011b). To
determine if 6AzGlcNAc specifically modifies O-GlcNAcylated proteins, we first took
advantage of the chimeric, secreted protein GlyCAM-IgG that contains both an N-linked
and multiple mucin O-linked glycosylation sites (Zaro et al., 2011b). NIH3T3 cells that
stably express GlyCAM-IgG via retroviral transformation were treated with
Ac
36AzGlcNAc, Ac4GlcNAz, or Ac 4GlcNAc at 200 μM concentrations for 48 hours. At
this time, GlyCAM-IgG was immunoprecipitated from the media using
protein-A-conjugated beads. In-gel fluorescence scanning, following CuAAC with
alk-rho, showed that while GlcNAz robustly labels GlyCAM-IgG, as expected based on
our previous results(Zaro et al., 2011b), 6AzGlcNAc does not (Figure 2-8A). This
demonstrates that while GlcNAz does label the major types of cell-surface glycosylation,
6AzGlcNAc does not. Next, to confirm that 6AzGlcNAc labels O-GlcNAcylated
proteins, we treated NIH3T3 cells that were stably transfected with the FLAG-tagged
transcription factor FoxO1 with Ac
36AzGlcNAc, Ac 4GlcNAz, or Ac 4GlcNAc at 200 μM
concentrations for 24 hours. In contrast to GlyCAM-IgG, in-gel fluorescence showed that
both MCRs robustly labeled FoxO1A (Figure 2-8B), demonstrating that 6AzGlcNAc is a
highly-selective MCR of O-GlcNAc modifications.
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To rule out the possibility that 6AzGlcNAc was excluded from GlyCAM-IgG but labeled
other cell-surface glycoproteins, NIH3T3 cells were treated with Ac
36AzGlcNAc,
Ac
4GlcNAz, or Ac 4GalNAz at 200 μM for 16 hours before being harvested and submitted
to copper-free click chemistry using commercially available DBCO-biotin. After
subsequent incubation with FITC-conjugated avidin, cell-surface glycoprotein labeling
by each chemical reporter was analyzed using flow cytometry. No labeling over
background was observed with Ac
36AzGlcNAc while labeling was observed with
Ac
4GlcNAz and Ac 4GalNAz (Figure 2-8C). Notably, this corroborates live-cell flow
cytometry data from the Bertozzi lab where they observed some cell-surface labeling
with GlcNAz and no labeling with 6AzGlcNAc (Saxon et al., 2002).
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Figure 2-8. Glycoprotein specificity of 6AzGlcNAc. NIH3T3 cells stably expressing either
GlyCAM-IgG (A) or Flag-tagged FoxO1 (B) were treated with the indicated MCRs or
Ac4GlcNAc, followed by immunoprecipitation, CuAAC, and analysis by in-gel fluorescence
scanning. (C) NIH3T3 cells were treated with Ac36AzGlcNAc, Ac 4GlcNAz, Ac 4GalNAz, or
Ac4GlcNAc (all at 200 μM) for 16 hours at which time cells were harvested and subjected to
copper-free click chemistry with DBCO-biotin. After incubation with FITC-avidin, live-cell
surface labeling was analyzed by flow cytometry.
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Direct comparison of 6AzGlcNAc, GlcNAz, and GalNAz as metabolic chemical-reporters
of glycosylation.
To identify the proteins labeled by 6AzGlcNAc and compare them to those enriched by
the previous MCRs GlcNAz and GalNAz, NIH3T3 cells were treated in triplicate with
either Ac
36AzGlcNAc, Ac4GlcNAz, Ac 4GalNAz, or Ac4GlcNAc as a control (all at 200
μM) for 16 hours. At this time cells were lysed using denaturing conditions (4% SDS)
and subjected to CuAAC conditions with an alkyne-bearing biotin tag. The proteomes
were then reduced, alkylated, and subjected to biotin-enrichment using
streptavidin-conjugated beads. After extensive washing to remove unlabeled proteins,
on-bead trypsinolysis afforded peptides that were analyzed using LC-MS/MS, identified
using Proteome Discoverer and Mascot, and quantified by spectral counting. Labeled
proteins were identified as those that met the following threshold criteria: First, proteins
must have been identified by at least 1 unique peptide in each of the three data sets and a
total of 3 spectral counts in the sum of three replicate data sets. Second, the sum of
spectral counts of the MCR-treated samples must be 3-times greater than those in the
GlcNAc labeled samples. Finally, the number of spectral counts in the MCR treated
sample compared to the control must be statistically significant (p < 0.05, t-test). Using
these criteria, 366 proteins were identified as being labeled by 6AzGlcNAc, including
many known O-GlcNAcylated proteins, such as the three annotated in black in Figure
2-9A (MAP4 , NEDD4, and HCF1). GlcNAz and GalNAz labeling identified 359
proteins and 348 proteins, respectively. In contrast to 6AzGlcNAc, these lists included
both known O-GlcNAcylated proteins and proteins that are exclusively localized to the
124
extracellular space or the lumen of the secretory pathway and lysosome, such as the three
annotated in red in Figure 2-9A (fibronectin, calumenin, and α-glucosidase). Comparison
of the three proteomics lists showed that 6AzGlcNAc has greater overlap with GlcNAz
than GalNAz (Figure 2-9B), consistent with with previous studies that show more
efficient global-incorporation of GalNAz versus GlcNAz into cell-surface glycoproteins
(Hang et al., 2003; Saxon et al., 2002). Importantly, many of the proteins that were
identified by 6AzGlcNAc have been previously identified in other O-GlcNAc proteomic
studies (Slade et al., 2012; Steentoft et al., 2013).
Figure 2-9. Identification of O-GlcNAcylated proteins using 6AzGlcNAc. (A) NIH3T3 cells
were treated with Ac36AzGlcNAc, Ac 4GlcNAz, Ac 4GalNAz, or Ac 4GlcNAc (all at 200 μM) for
16 hours. At this time, the corresponding cell-lysates were subjected to CuAAC with
alkyne-biotin, enrichment with streptavidin-coated beads, and on-bead trypsinolysis. Proteins
identified by LC-MS/MS are graphically presented as total number of positive minus total
number of control spectral counts. Three known O-GlcNAcylated proteins are annotated in black
and three known extracellular/lumenal proteins are annotated in red. (B) Overlap between
proteins identified using 6AzGlcNAc, GlcNAz and, GalNAz. (C) Graphical representation of
enriched proteins based on whether their localization is exclusively intracellular (i.e.,
cytoplasmic, nuclear, or mitochondrial), exclusively extracellular or lumenal (i.e., ER, Golgi,
lysosome), or have domains in both (e.g., transmembrane protein).
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We next annotated the proteins in our lists based on their characterized localizations
(Figure 2-9C). Proteins with uncharacterized localizations were omitted. Consistent with
specific labeling of O-GlcNAcylated proteins, 6AzGlcNAc treatment enriched 350
exclusively intracellular proteins (i.e., nuclear, cytosolic, and mitochondrial) and 8
proteins that can be localized to both the cytosol and extracellular space or lumenal
compartments (e.g., transmembrane proteins). Notably, only 1 exclusively extracellular or
lumenal protein (galectin-1) was found in this list. In contrast, 10 and 72 exclusively
extracellular or lumenal proteins were found using GlcNAz and GalNAz treatment,
respectively (Figure 6-9C), reenforcing the data demonstrating the non-specific labeling
of multiple types of glycosylation by GlcNAz and GalNAz.
Discussion and Conclusion
The use of MCRs for the visualization and identification of protein glycosylation has
expanded the ability to investigate these key post-translational modifications. However,
recent evidence from our lab and others has demonstrated that many MCRs of protein
glycosylation lack specificity, as they are incorporated into multiple types of glycans
(Banerjee et al., 2010; Boyce et al., 2011; Zaro et al., 2011b). We previously showed that
small changes to the chemical structure of MCRs can have a large impact on their
distribution into different glycans (Bateman et al., 2013a; Zaro et al., 2011b). Following
this chemical-optimization theme further, we identified an MCR (6AzGlcNAc) that
robustly labeled a variety of proteins in living mammalian cells. Using a fluorescent
alkyne-tag, we compared 6AzGlcNAc to the previous MCR, GlcNAz, and demonstrated
126
that 6AzGlcNAc is efficiently incorporated onto proteins allowing visualization in as
little as 2 to 4 hours after treatment. Furthermore, 6AzGlcNAc removal from protein is
dependent on the activity of OGA, demonstrating that it is dynamically incorporated into
O-GlcNAcylated proteins. Using two reporter proteins, we next demonstrated that while
GlcNAz labels both secreted glycoproteins and O-GlcNAcylated proteins, 6AzGlcNAc is
specific for O-GlcNAc modifications. This is consistent with our flow cytometry data and
previous reports (Saxon et al., 2002) that both showed essentially no cell-surface labeling
by 6AzGlcNAc and that chemically-synthesized UDP-6AzGlcNAc is a substrate for
recombinant O-GlcNAc transferase (Mayer et al., 2011).
Unlike GlcNAz, 6AzGlcNAc cannot be metabolized to the corresponding UDP-sugar
donor by the canonical GlcNAc salvage-pathway (Figure 2-3), as the first step involves
phosphorylation at the 6-hydroxyl of the monosaccharide. Therefore, an alternative
enzyme must directly phosphorylate 6AzGlcNAc at the 1-hydroxyl to bypass this
roadblock. Taking a candidate-based approach, we tested GalK2 and AGM1 in vitro to
determine if they could generate 6AzGlcNAc-1-phosphate. We did not observe any
product formation using GalK2, and knockdown of GalK2 in living cells using shRNA
did not result in reduced protein-labeling by 6AzGlcNAc (Figure 2-4A). Notably,
however, we found that AGM1 is capable of directly generating 6AzGlcNAc-1-phosphate
when its normal substrate, GlcNAc-6-phosphate is added to the reaction mixture (Figure
2-5). Based on the enzymatic mechanism, we conclude that AGM1 removes the
phosphate from GlcNAc-6-phosphate to generate the known phosphoenzyme
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intermediate (Nishitani et al., 2006), followed by binding of 6AzGlcNAc and
phosphorylation of the 1-hydroxyl. This is consistent with the reversible nature of
AGM1’s activity, where the monosaccharide substrates can bind the active site with
either the 1- or 6-hydroxyl groups oriented towards the catalytic serine. Additionally, we
also showed that once 6AzGlcNAc-1-phosphate is formed it can be enzymatically
transformed to UDP-6AzGlcNAc by AGX1 (Figure 2-4C). We do not know if
UDP-6AzGlcNAc can be epimerized to UDP-6AzGalNAc in cells; however, even if this
metabolite is formed, previous studies by Bertozzi and co-workers demonstrated that
UDP-6AzGalNAc is not a substrate for the polypeptide-N-acetyl-galactosamine
transferases (Hang et al., 2003). Together, these results suggest an unappreciated
metabolic flexibility in mammalian cells. AGM1 and potentially other, yet unidentified,
small-molecule phosphotransferases may contribute to the salvaging of natural
monosaccharides from the environment. Furthermore, they have potentially important
implications for the metabolism of bacterial or abiotic carbohydrates that would
otherwise be assumed to not enter mammalian biosynthetic pathways. Finally, our results
challenge a dogma in MCR design, which relies on well-established metabolic pathways
and directly resulted in the previous dismissal of 6AzGlcNAc as a viable MCR in living
cells (Mayer et al., 2011; Saxon et al., 2002).
To further confirm the specificity of 6AzGlcNAc and demonstrate any advantages over
other MCRs previously used to study O-GlcNAcylated proteins, we performed a
proteomics experiment using 6AzGlcNAc, GlcNAz, and GalNAz in combination with
128
alkyne-biotin and on-bead trypsinolysis. We found that enrichment with 6AzGlcNAc
resulted in the identification of essentially only intracellular proteins that cannot contain
glycans (e.g., N-linked or mucin O-linked) that are added in the secretory pathway. This
confirms the high degree of specificity of 6AzGlcNAc for O-GlcNAcylated proteins.
Consistent with our fluorescence data, GlcNAz was less selective, resulting in the
enrichment of 28 proteins that potentially bear secretory-pathway glycans and 10
exclusively extracellular or lumenal proteins. Finally, GalNAz was the least selective,
since it enriched only 226 exclusively intracellular proteins and 72 proteins that are only
extracellular or lumenal. We believe that this lack of selectivity is one reason why a
recent study using GalNAz required subcellular fractionation and two-dimensional
electrophoresis to identify the potential O-GlcNAcylation of the voltage-dependent
anion-selective channel protein 2 (VDAC2) (Palaniappan et al., 2013), while the same
protein was readily identified by 6AzGlcNAc labeling without any biochemical
manipulations. This specificity is a significant improvement over other MCRs that require
biochemical manipulations (e.g., cell fractionation) to exclude cell-surface
glycoproteins(Boyce et al., 2011).
Previous direct-comparisons of the selectivity of different glycoprotein MCRs are
somewhat limited (Banerjee et al., 2010; Bateman et al., 2013a; Boyce et al., 2011; Saxon
et al., 2002; Zaro et al., 2011b). The Bertozzi lab reported that GalNAz has superior
O-GlcNAc labeling-efficiency compared to GlcNAz due to more efficient metabolic
conversion of GalNAz to UDP-GalNAz and subsequent epimerization to UDP-GlcNAz
129
(Boyce et al., 2011). Our in-gel fluorescence data do not support these data, as GlcNAz
and GalNAz resulted in qualitatively similar levels of protein labeling in a variety of cell
lines (Figure 6-6). Interestingly, only a minority of cell-lines show similar global-patterns
of labeling between GlcNAz and GalNAz (e.g., MCF7), while most are significantly
different. This also is true of 6AzGlcNAc, which often shows different labeling patterns
and intensities from both GlcNAz and GalNAz (Figure 2-6). This supports our results
that each of the MCRs is incorporated into different types of glycoproteins and utilizes
independent metabolic enzymes for the generation of the corresponding UDP
donor-sugars. Notably, while 6AzGlcNAc is the most selective reporter of
O-GlcNAcylation, it requires longer labeling-times to achieve the same signal-to-noise as
GlcNAz (Figure 2-7A), highlighting a potential tradeoff between labeling efficiency and
specificity. However, based on our results, we predict that any bottlenecks in the
metabolism of 6AzGlcNAc will not dramatically hamper the visualization and
identification of O-GlcNAcylated proteins. Furthermore, the extent of O-GlcNAcylation
and identity of modified proteins has been shown to be dependent on the cellular
concentration of UDP-GlcNAc (Comer and Hart, 2001; Kreppel et al., 1997). Therefore,
it could be advantageous to have limited metabolic conversion of an MCR to minimize
the chances of altering the endogenous repertoire of O-GlcNAcylated proteins, as long as
the labeling is above the detection limit. We are currently exploring if different
concentrations of MCR treatment change the overall levels of O-GlcNAcylation. Despite
the increased labeling-efficiency of GlcNAz and GalNAz compared to 6AzGlcNAc,
approximately the same total number of spectral counts were found in our comparative
130
proteomics experiment. We believe that this could be due to an excess of input that
exceeded the capacity of the streptavidin beads, resulting in equal total levels of protein
enrichment prior to trypsinolysis and identification.
Coupled with the ever-growing toolkit of commercially available and custom
azide-reactive tags, including terminal alkynes, cyclooctynes, and phosphines, we predict
that 6AzGlcNAc will become the most powerful and readily used MCR for the study of
O-GlcNAcylation. In particular, metabolic labeling strategies have the unique ability to
isolate time-resolved protein modifications that only occur during cell labeling.
Furthermore, MCRs can be used in pulse-chase experiments to measure the dynamic
removal of O-GlcNAc modifications in living cells. Finally, the successful application of
synthetic chemistry to identify a selective MCR of O-GlcNAcylation suggests that the
same chemical-strategy could be used to create reporters that are specific for other types
of protein glycosylation. Coupled with new bioorthogonal reactions (e.g., tetrazene
clycoadditions) that enable more diverse functional groups to be incorporated into MCRs
(Cole et al., 2013; Niederwieser et al., 2013; Patterson et al., 2012), we predict that a
diverse library of MCRs can be created to enable the specific visualization and
identification of the several types of glycosylation in mammals and other organisms.
Materials and Methods
All reagents used for chemical synthesis were purchased from Sigma-Aldrich, Alfa Aesar
or EMD Millipore unless otherwise specified and used without further purification.
131
DBCO-biotin was purchased from Click Chemistry Tools. FITC-avidin was purchased
from Sigma. All anhydrous reactions were performed under argon or nitrogen
atmosphere. Analytical thin-layer chromatography (TLC) was conducted on EMD Silica
Gel 60 Å F
254 plates with detection by ceric ammonium molybdate (CAM), anisaldehyde
or UV . For flash chromatography, 60 Å silica gel (EMD) was utilized.
1
H spectra were
obtained at 400, 500, or 600 MHz on a Varian spectrometers Mercury 400, VNMRS-500,
or -600. Chemical shifts are recorded in ppm (δ) relative to solvent. Coupling constants
(J) are reported in Hz.
13
C spectra were obtained at 100, 125 or 150 MHz on the same
instruments.
Chemical Synthesis.
Known chemical reporters Ac
4GlcNAz(Saxon et al., 2002) and Ac 36AzGlcNAc (Saxon et
al., 2002), were synthesized according to literature procedures. The fluorescent detection
tag alk-rho(Charron et al., 2009) and the OGA inhibitor Thiamet-G(Yuzwa et al., 2008)
were also synthesized in lab according to literature procedures. Ac
36AzGlcNAc and
alkyne-azo-biotin were synthesized according to literature procedures as described below.
Compound 2.1 1,3,4-Tri-O-acetyl-2-acetamido-6-azido-2,6-dideoxy-D-
glucopyranose (6AzGlcNAc). Commercially available
2-deoxy-2-N-acetyl-glucopyranose (1.00 g, 4.52 mmol) was
dissolved in anhydrous pyridine under nitrogen and cooled to -20
°C. p-Toluenesulfonyl chloride (1.04 g, 5.43 mmol) was dissolved
O
AcO
NH
O
N
3
AcO
OAc
132
in anhydrous pyridine (3 mL) and the solution was added dropwise over 20 minutes to
the above reaction mixture. Upon completion of the addition, the reaction was warmed to
rt and stirred for 18 hours. The mixture was concentrated by vacuum and used without
further purification. The crude was then resuspended in N, N-dimethylformamide (20
mL) under a nitrogen atmosphere. Sodium azide (1.47 g, 22.6 mmol) was added and the
reaction was stirred for 3 d at 50 °C. The reaction mixture was then concentrated by
vacuum and resuspended in pyridine (30 mL). Acetic anhydride (10.0 mL, 66.0 mmol)
was added and the mixture was stirred for 16 h at rt. Upon completion, solvent was
removed under reduced pressure and the resulting mixture was redissolved in CH
2Cl 2
(200 mL) and washed with 1 M HCl (2 × 100 mL), saturated aqueous sodium bicarbonate
(2 × 100 mL) and water (2 × 100 mL). Organic layer was dried over sodium sulfate. The
resulting crude mixture was purified by column chromatography (65% ethyl acetate in
hexanes) to afford 1.01 g of the product in 60% yield over three steps.
1
H NMR (400
MHz, CDCl
3): δ (ppm) 6.18 (d, J = 3.7 Hz, 1H), 5.61 (d, J = 8.9 Hz, 1H), 5.22 (ddd, J =
10.8, 9.5, 0.4 Hz, 1H), 5.13 (dd, J = 9.9, 9.4 Hz, 1H), 4.46 (ddd, J = 10.9, 9.0, 3.7 Hz,
1H), 4.14 – 4.02 (m, 1H), 3.94 (dddd, J = 10.0, 5.6, 3.1, 0.6 Hz, 1H), 3.38 – 3.26 (m, 2H),
2.19 (s, 3H), 2.06 – 2.01 (m, 6H), 1.92 (d, J = 2.6 Hz, 3H).
Compound 2.2 4-(prop-2-yn-1-yl)phenol (Kimura et al., 2004). Hydroquinone (11.0 g,
0.910 mmol) and propargyl chloride (7.50 g, 0.101 mmol) were
dissolved in ethanol (20 mL) under an argon atmosphere in a
three-neck flask equipped with an addition funnel. The reaction
OH
133
was heated to reflux to dissolve all solids. KOH (0.10 M in water) was added dropwise
through the addition funnel. The mixture was then stirred for 20 h upon which time the
reaction was cooled and solvent was removed by vacuum. The resulting crude mixture
was dissolved in CH
2Cl 2 and extracted with dilute, aqueous KOH. The aqueous layer was
then brought to a neutral pH by the addition of 1 M HCl and subsequently extracted with
CH
2Cl 2. The organic layer was washed with water and dried over sodium sulfate, filtered
and concentrated. The crude mixture was purified by column chromatography (10% ethyl
acetate:hexanes) to afford the pure product (4.07 g, 28%).
1
H NMR (400 MHz, CDCl 3): δ
(ppm) 6.87 (dd, J = 9.1, 1.1 Hz, 2H), 6.79 (d, J = 9.0 Hz, 2H), 4.62 (dd, J = 2.4, 1.0 Hz,
2H), 2.51 (t, J = 2.4 Hz, 1H).
Compound 2.3 (E)-methyl 4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)benzoate. To
a suspension of methyl-4-amino-benzoate in 6 M HCl was
added sodium nitrite at 0 °C. The reaction was let stir for
30 mins. The crude reaction mixture was used in the
following reaction. Compound 2.2 (1.30 g, 18.9 mmol)
was dissolved in water:THF (2:1) and cooled to 0 °C. Potassium carbonate (52.0 g, 376
mmol) was added and reaction let stir for 30 mins upon which time
4-(methoxycarbonyl)benzenediazonium chloride was added dropwise. The reaction was
allowed to warm to rt and was stirred for 18 h. The reaction was poured over water and
extracted with ethyl acetate (3 x 200 mL). The organic layer was dried over sodium
sulfate, filtered, and concentrated. The resulting crude mixture was purified by column
OH
N
N
O
O
134
chromatography by first starting at 10% ethyl acetate:hexanes and increasing to 20%
ethyl acetate:hexanes to elute the product. Concentration under decreased pressure
affords the product as a yellow oil (2.00 g, 90%).
1
H NMR (400 MHz, CDCl 3): δ (ppm)
8.21 (d, J = 8.5 Hz, 2H), 7.93 (d, J = 8.6 Hz, 2H), 7.56 (d, J = 3.1 Hz, 1H), 7.12 (dd, J =
9.1, 3.1 Hz, 1H), 7.01 (d, J = 9.1 Hz, 1H), 4.76 (d, J = 2.3 Hz, 2H), 3.97 (s, 3H), 2.57 (t, J
= 2.4 Hz, 1H).
13
C NMR (125 MHz, CDCl 3): δ (ppm) 166.25, 153.08, 150.83, 148.27,
137.01, 131.92, 130.80, 130.17, 128.36, 123.68, 122.02, 119.17, 111.60, 78.41, 75.86,
56.77, 52.38. MALDI-MS calculated for C
17H 15N 2O 3 [M+H]
+
: 295.1083, found
293.9045.
Compound 2.4 (E)-4-((2-hydroxy-5-(prop-2-yn-1-
yl)phenyl)diazenyl)benzoic acid. Compound 2.3 (0.124 g,
0.421 mmol) was dissolved in tetrahydrofuran (2 mL).
NaOH (0.758 mg, 1.90 mmol) dissolved in water was
added and reaction let stir 18 h. Upon completion, a color change from purple to orange
is seen. The reaction was neutralized by the dropwise addition of acetic acid and
subsequently concentrated under reduced pressure to remove solvent. The resulting crude
mixture was dissolved in CH
2Cl 2 and washed with water (2 x 50 mL). The organic layer
was dried over sodium sulfate, filtered, and concentrated. The crude mixture was column
purified (8:1.5:0.5 ethyl acetate:methanol:water) to afford the pure product as a bright
orange solid (0.825 g, 70%).
1
H NMR (600 MHz, D 6-DMSO): δ (ppm) 8.05 (d, J = 8.5
Hz, 2H), 7.85 (d, J = 8.5 Hz, 2H), 7.46 (d, J = 3.0 Hz, 1H), 7.01 (dd, J = 9.2, 3.0 Hz, 1H),
OH
N
N
OH
O
135
6.88 (d, J = 8.9 Hz, 1H), 4.68 (d, J = 2.3 Hz, 2H), 2.87 (t, J = 2.4 Hz, 1H).
13
C NMR (125
MHz, D
6-DMSO): δ (ppm) 216.29, 178.12, 162.96, 160.46, 160.22, 148.17, 146.80,
140.14, 132.53, 132.07, 129.05, 115.05, 89.05, 88.03, 65.94, 40.41. MALDI-MS
calculated for C
16H 12N 2O 3Na [M+Na]
+
: 303.0740, found 302.9541.
Compound 2.5 (E)-2,5-dioxopyrrolidin-1-yl-4-((2-hydroxy-5-(prop-2-yn-1-yl)
phenyl)diazenyl)benzoate. To a solution of 2.4 (0.180 g,
0.642 mmol) in THF under argon was added
N-hydroxysuccinimide (0.177 g, 1.54 mmol) and N,N’-
Dicyclohexylcarbodiimide (0.317 g, 1.54 mmol). The
reaction was let stir for 18 h at rt at which time the reaction was concentrated by vacuum.
The mixture was dissolved in ethyl acetate and filtered to remove solids. The
flow-through was concentrated and the crude product was purified by column
chromatography (1:10 ethyl acetate:CH
2Cl 2) to afford the product as a dark red solid that
was used in the subsequent reaction without further purification.
Compound 2.6 (E)-4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)-N-(13-
oxo-17-(2-oxohexahydro-1H-thieno[3,4-
d]imidazol-4-yl)-3,6,9-
trioxa-12-azaheptadecyl)benzamide
(Alk-azo-biotin). Compound 2.5 (0.040 g, 0.101 mmol) was dissolved in anhydrous N,N’-
dimethylformamide (1 mL) under argon. EZ-Link Amine PEG
3-Biotin (0.460 mg, 0.111
OH
N
N
O
O
N
O
O
OH
N
N
O
H
N
O
3
H
N
O
S
NH
HN
O
136
mmol) (Thermo Scientific) was added and reaction let stir for 18 h upon which time
solvent was removed by vacuum. The resulting crude mixture was purified by RP-HPLC
over a C18 semi-preparative column (The Nest Group) using a 5.5-44% B linear gradient
over 10 min before switching to a 44-100% B linear gradient over 40 mins,
t
R = 18 min
(buffer A: 0.1% TFA in water, buffer B: 0.1% TFA, 90% ACN in water) and lyophilized
to afford the pure product as an orange solid (0.022 g, 31%). ESI-MS calculated for
C
34H 40N 6O 8S (oxidized at the biotin cysteine) [M+Na]
+
: 719.28, found 719.20.
Compound 2.7 6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine. Commercially
available 2-deoxy-2-N-acetyl-glucopyranose (2.50 g, 11.3 mmol)
was co-evaporated from toluene and dissolved in anhydrous
pyridine (20 mL). The reaction mixture was cooled to -20 °C.
p-Toluenesulfonyl chloride (2.59 g, 13.6 mmol) was then dissolved is anhydrous pyridine
(5 mL) and added drop-wise to the stirring mixture. Upon completion of addition, the
reaction was allowed to warm to room temperature and stirred for 16 h under an argon
atmosphere. To purify, the reaction was concentrated under reduced pressure and the
crude mixture purified by column chromatography (7:1:0.5 ethyl acetate:methanol:water)
to afford the product as a yellow oil (1.78 g).
1
H NMR (500 MHz, CD 3OD) α-anomer: δ
7.74 (d, J = 8.3 Hz, 2H), 7.26 (d, J = 8.2 Hz, 2H), 5.13 (d, J = 3.6 Hz, 1H), 3.99 (m, 1H),
3.90 (dd, J = 2.2, 10.6 Hz, 1H), 3.73 (m, 1H), 3.55 (dd, J = 3.2, 13.3 Hz, 1H), 3.44 (dd, J
= 5.4, 12.6 Hz, 1H), 3.38 (m, 1H), 2.39 (s, 3H), 2.01 (s, 3H). The product was used in the
subsequent reaction with no further characterization.
O
AcHN
OTs
HO
HO
OH
137
Compound 2.8 6-azido-6-deoxy-N-acetyl-glucosamine (6AzGlcNAc, 9) (Saxon et al.,
2002). Compound 2.7 (1.78 g, 4.73 mmol) was coevaporated
from toluene and dissolved in anhydrous N,N’-
dimethylformamide (20 mL). Sodium azide (1.54 g, 23.7 mmol)
was then added and the reaction warmed to 50 °C. The reaction was stirred for 3 d after
which time the reaction was cooled and concentrated under reduced pressure. The crude
mixture was purified by silica gel chromatography (9:1:0.5 ethyl acetate:methanol:water)
to afford the product as a white solid (402 mg, 14% yield over 2 steps). The sugar was
further purified by RP-HPLC over a C18 semi-preparative column (The Nest Group)
using a 5-15% B linear gradient over 10 min,
t
R = 2.5 min (buffer A: 0.1% TFA in water,
buffer B: 0.1% TFA, 90% ACN in water).
1
H NMR (500 MHz, (CD 3)2SO) α-anomer: δ
7.69 (d, J = 8.3 Hz, 1H), 4.94 (app s, 1H), 3.78 (m, 1H), 3.62 (m, 1H), 3.50 (m, 2H), 3.37
(m, 1H), 3.10 (app t, J = 9.2 Hz, 1H), 1.83 (s, 3H).
13
C NMR (125 MHz,(CD 3)2SO) β-
anomer: δ 169.39, 90.77, 71.85, 70.52, 70.17, 54.21, 51.60, 22.67.
Compound 2.9 α-1-O-benzyl-N-acetyl-glucosamine (Mayer et al., 2011). The procedure
was adapted from literature.{Sharma:1987up} Commercially
available 2-deoxy-2-N-acetyl-glucopyranose (5.00 g, 22.6 mmol)
was suspended in benzyl alcohol (50 mL) and concentrated HCl
was added (1 mL). The solution was warmed to 75 °C and stirred for 4 h after which time
the reaction was cooled and poured into diethyl ether (400 mL) with vigorous stirring. A
O
AcHN
N
3
HO
HO
OH
O
AcHN
OH
HO
HO
OBn
138
white precipitate was observed and the mixture left at 4 °C for 16 h. The precipitate was
then filtered and washed with diethyl ether (50 mL) to remove remaining benzyl alcohol.
The filtrated was dried and recrystallized in a minimal amount of isopropanol to afford
the product as white solid (2.48 g, 7.98 mmol, 35% yield).
1
H NMR (500 MHz,
(CD
3)2SO): δ 7.82 (d, J = 8.2 Hz, 1H), 7.38-7.28 (m, 5H), 4.71 (d, J = 3.5 Hz, 1H), 4.68
(d, J = 12.5 Hz, 1H), 4.31 (d, J = 12.5 Hz, 1H), 3.80 (q, J = 6.1 Hz, 1H), 3.70-3.64 (m,
2H), 3.55-3.47 (m, 2H), 3.18 (t, J = 9.02 Hz, 1H), 1.83 (s, 3H).
Compound 2.10 α-1-O-benzyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine
(Mayer et al., 2011). Compound 2.9 (2.36 g, 7.58 mmol) was
co-evaporated from toluene and dissolved in anhydrous pyridine (20
mL) under an argon atmosphere. The mixture was then cooled to -20
°C. p-toluenesulfonyl chloride (1.74 g, 9.10 mmol), freshly rescrystallized from CH
2Cl 2,
was dissolved in pyridine (7 mL) and added dropwise over 20 min. The reaction was
stirred at -20 °C for 1 h and the dry ice bath replaced with an ice bath. The reaction was
allowed to warm to room temperature over 16 h. Upon completion, the mixture was
concentrated to remove pyridine and purified over silica gel (9:1:0.5
EtOAc:methanol:water) to afford product (1.85 g, 3.97 mmol, 52% yield).
1
H NMR (500
MHz, CD
3OD): δ 7.85 (d, J = 10.4 Hz, 2H), 7.46 (d, J = 1.1 Hz, 2H), 7.36 (m, 5H), 4.76
(d, J = 4.5 Hz, 1H), 4.65 (d, J = 15 Hz, 1H), 4.45 (d, J = 14.8 Hz, 1H), 4.36 (dd, J = 2.6,
13.6 Hz, 1H), 4.27 (dd, J = 7.3, 13.6 Hz, 1H), 3.88 (dd, J = 4.6, 13.5 Hz, 1H), 3.81-3.78
(m, 1H), 3.71-3.66 (m, 1H), 3.35-3.31 (m, 1H), 2.46 (s, 3H), 1.97 (s, 3H).
O
AcHN
OTs
HO
HO
OBn
139
Compound 2.11 3,4-di-O-acetyl-α-1-O-benzyl-6-O-p-methylbenzenesulfonate-N-acetyl-
glucosamine (Mayer et al., 2011). 2.10 (1.85 g, 3.97 mmol) was
resuspended in pyridine (20 mL) and acetic anhydride (1.12 mL,
11.01 mmol). The reaction was stirred for 3 h at room temperature
after which time the reaction mixture was concentrated and
purified over silica gel (75% EtOAc in hexanes) to afford product in quantitative yield
(2.18 g, 3.97 mmol).
1
H NMR (500 MHz, CDCl 3): δ 7.79 (d, J = 10.3 Hz, 2H), 7.37-7.29
(m, 7H), 5.63 (d, J = 11.9 Hz, 1H), 5.20 (dd J = 11.7, 13.5 Hz, 1H), 4.96 (t, J = 12.1 Hz,
1H), 4.82 (d, J = 4.6 Hz, 1H), 4.67 (d, J = 14.7 Hz, 1H), 4.44 (d, J = 14.8 Hz, 1H), 4.26
(td, J = 4.6, 12.6 Hz, 1H), 4.07 (d, J = 5.0 Hz, 2H), 4.04-4.00 (m, 1H), 2.44 (s, 3H), 1.98
(d, J = 2.6 Hz, 6H), 1.86 (s, 3H).
Compound 2.12 3,4-di-O-acetyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine.
Procedure adapted from published literature (Mayer et al., 2011).
2.11 (969 mg, 1.76 mmol) was resuspended in methanol.
Pd(OH)
2/C (10% Pd) was added and a balloon of H 2 was attached.
The reaction was monitored by TLC (75% EtOAc in hexanes) and
stirred for 48 h to completion. The mixture was then filtered over a pad of celite and the
flow-through evaporated to yield the product (710 mg, 1.55 mmol) that was used in
subsequent reactions with no further characterization.
O
AcHN
OTs
AcO
AcO
OBn
O
AcHN
OTs
AcO
AcO
OH
140
Compound 2.13 Diallyl(3,4-di-O-acetyl-6-O-p-methylbenzenesulfonate-N-acetyl-
glucosamine)-α-1-phosphate. 2.12 (629 mg, 1.37 mmol)
was coevaporated with toluene and resuspended in CH
2Cl 2
(10 mL) under an argon atmosphere.
5-(Ethylthio)-1H-tetrazole (1.07 g, 8.22 mmol) was added
and the reaction stirred for 15 min. Diallyl-N,N’-
diisopropylphosphoramidite (1.00 g, 4.11 mmol) was added dropwise, and the reaction
stirred for 2 h until completed as determined by TLC (5% methanol in CH
2Cl 2). At this
time, the reaction was cooled to -78 °C and freshly recrystallized m-chloroperoxybenzoic
acid was added (1.18 g, 6.85 mmol). The reaction was allowed to proceed for 10 min
after which time the dry ice bath was replaced with an ice bath, and the reaction was
slowly warmed to room temperature over 1 h. Upon completion, the reaction was diluted
with CH
2Cl 2 (50 mL) and washed 2x each with saturated sodium thiosulfate, saturated
sodium bicarbonate, water and brine. The organic layer was then concentrated and
purified over silica gel (35%-45% acetone in hexanes) to afford the product (717 mg,74%
yield over 2 steps).
1
H NMR (500 MHz, CDCl 3): δ 7.70 (d, J = 8.4 Hz, 2H), 7.29 (d, J =
7.8 Hz, 2H), 6.06 (d, J = 9.4 Hz, 1H), 5.93-5.82 (m, 2H), 5.54 (dd, J = 3.3, 6.3 Hz, 1H),
5.32 (ddd, J = 17.1, 12.3, 1.4 Hz, 2H), 5.23 (ddd, J = 10.6, 9.6, 1.2 Hz, 1H), 5.12 (dd, J =
10.9, 9.4 Hz, 1H), 4.96 (dd, J = 10.3, 9.5 Hz, 1H), 4.54-4.49 (m, 3H), 4.25-4.21 (m, 1H),
4.19-4.15 (m, 1H), 4.04 (dd, J = 11.1, 2.6 Hz, 1H), 3.98 (dd, J = 11.1, 5.1 Hz, 1H), 2.39
(s, 3H), 1.94 (s, 3H), 1.93 (s, 3H), 1.86 (s, 3H);
13
C NMR (125 MHz, CDCl 3): δ 171.08,
170.31, 169.06, 145.21, 132.23, 132.12, 132.07, 131.91, 131.86, 129.86, 128.05, 119.06,
O
AcHN
OTs
AcO
AcO
O
P
O
O
O
141
118.93, 95.65, 95.60, 69.73, 69.37, 68.79, 67.67, 66.99, 51.69, 51.63, 22.87, 21.63, 20.58,
20.45;
31
P NMR (500 MHz, CDCl 3): δ -2.77; APCI-HRMS calculated for
C
25H 34NO 13PSNa [M+Na]
+
: 642.1488, found 642.1398.
Compound 2.14 Diallyl(6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-
phosphate. 2.13 (669 mg, 1.08 mmol) was resuspended is
methanol (10 mL). Freshly made NaOMe was added
dropwise until pH 9-10 was reached. The reaction was
monitored by TLC (10% methanol in CH
2Cl 2) and was
determined complete after 1.5 h. Upon completion, the
reaction was quenched with acetic acid and concentrated to afford the crude. Silica gel
chromatography (7% methanol in CH
2Cl 2) yielded the product (282 mg, 57% yield).
1
H
NMR (500 MHz, CD
3OD): 7.79 (d, 2H, J = 8.3 Hz), 7.44 (d, J = 8.5 Hz, 2H), 6.02-5.92
(m, 2H), 5.59 (dd, J = 8.5, 8.5 Hz, 1H), 5.42-5.37 (m, 2H), 5.29-5.26 (m, 2H), 4.60-4.55
(m, 4H), 4.32 (dd, J = 11.0, 1.9 Hz, 1H), 4.21 (dd, J = 11.0, 5.7 Hz, 1H), 3.93-3.89 (m,
1H), 3.86-3.82 (m, 1H), 3.64 (dd, J = 10.8, 8.8 Hz, 1H), 3.36 (t, J = 8.7 Hz, 1H), 2.46 (s,
3H), 1.98 (s, 3H);
13
C NMR (125 MHz, CDCl 3): δ 173.75, 146.59, 134.05, 133.63,
133.58, 133.57, 133.53, 131.02, 129.09, 118.89, 118.87, 97.39, 97.34, 73.35, 71.15,
71.12, 70.22, 69.89, 69.84, 69.79, 69.75, 55.08, 55.01, 22.51, 21.62;
31
P NMR (500 MHz,
CDCl
3): δ -2.51; APCI-HRMS calculated for C 21H 30NO 11PSNa [M+Na]
+
: 558.1169,
found 558.1154.
O
AcHN
OTs
HO
HO
O
P
O
O
O
142
Compound 2.15 Diallyl(6-azido-6-deoxy-N-acetyl-glucosamine)-α-1-phosphate. 2.14
(282 mg, 0.527 mmol) was coevaporated from toluene and
resuspended in N,N’-dimethylformamide (20 mL) under an
argon atmosphere. Sodium azide (172 mg, 2.63 mmol) was
added, and the reaction reaction warmed to 60 °C. The reaction
proceeded for 48 h after which time the reaction was concentrated and purified over silica
gel (7:2:1 EtOAc:methanol:water) to afford the product (151 mg, 71% yield).
1
H NMR
(500 MHz, CD
3OD): 6.07-5.73 (m, 2H), 5.40 (dd, J = 7.4, 3.3 Hz, 1H), 5.27-5.20 (m,
2H), 5.06-5.01 (m, 2H), 4.34-4.31 (m, 2H), 4.27-4.24 (m, 2H), 3.89-3.81 (m, 2H), 3.59
(dd, J = 10.6, 8.9 Hz, 1H), 3.49 (dd, J = 13.2, 2.5 Hz, 1H), 3.36-3.31 (m, 2H), 3.22-3.21
(m, 1H), 1.91 (s, 3H);
13
C NMR (125 MHz, CDCl 3): δ 173.81, 135.99, 135.92, 116.18,
116.15, 95.59, 95.54, 73.63, 72.48, 72.37, 67.33, 67.29, 67.17, 67.13, 55.33, 55.27, 52.63,
22.85;
31
P NMR (500 MHz, CDCl 3): δ 0.66, -1.37.
Compound 2.16 6-azido-6-deoxy-N-acetyl-glucosamine-1-phosphate (Cai et al., 2009).
2.15 (50 mg, 0.123 mmol) was resuspended in 4 mL
methanol:THF (1:1) under an argon atmosphere. p-Toluenesulfinic
acid sodium salt (44 mg, 0.246 mmol) and
Tetrakis(triphenylphosphine)-Palladium(0) (11 mg, 0.095 mmol) were added. The the
reaction was monitored by TLC (3:2:1 N-propanol:acetic acid:water) and determined
complete. The reaction was then evaporated under reduced pressure and purified by silica
gel chromatography (3:2:1 N-propanol:acetic acid:water). The sugar was further purified
O
AcHN
N
3
HO
HO
O
P
O
O
O
O
AcHN
N
3
HO
HO
OPO
3
-2
143
by RP-HPLC over a C18 semi-preparative column (The Nest Group) using a 0% B
isocratic flush over 10 min followed by a 0-50% B linear gradient from 10-20 min and a
second linear gradient 50-0% B 20-30 min,
t
R = 2.5-4 min (buffer A: 0.1% TFA in water,
buffer B: 0.1% TFA, 90% ACN in water).
1
H NMR (500 MHz, D 2O): δ 5.33 (dd, J = 7.3,
3.4 Hz, 1H), 3.94–3.83 (m, 2H), 3.68 (dd, J = 10.5, 9.1 Hz, 1H), 3.64-3.56 (m, 2H),
3.53-3.43 (m, 2H), 1.95 (s, 3H);
13
C NMR (125 MHz, D 2O): δ 174.63, 163.38, 163.10,
162.82, 162.53, 119.78, 117.46, 115.14, 112.82, 93.43, 93.38, 71.34, 70.42, 70.35, 58.59,
53.73, 53.66, 50.67, 33.71, 21.92;
31
P NMR (500 MHz, D 2O): δ -1.65; ESI-MS calculated
for C
8H 14N 4O 8P [M-H]
-
: 323.04, found 325.00.
Cell Culture. COS-7, HEK293, HeLa and MCF7 cells were cultured in DMEM media
(Corning) enriched with 10% fetal bovine serum (HyClone, ThermoScientific).
AmphoPack-293 retroviral packaging cells (Clontech) were cultured in DMEM media
(Corning) enriched with 10% fetal bovine serum (HyClone, ThermoScientific). NIH3T3
and MEF cells were cultured in high-glucose DMEM media (Corning) enriched with
10% fetal calf serum (HyClone, ThermoScientific). H1299 cells were cultured in RPMI
media enriched with 10% fetal bovine serum (HyClone, ThermoScientific). SH-SY5Y
cells were cultured in a 1:1 mixture of DMEM:F12 Medium (Corning) enriched with
10% fetal bovine serum (HyClone, ThermoScientific). All cell lines were maintained in a
humidified incubator at 37 ℃ and 5.0% CO
2.
144
Metabolic Labeling. To cells at 80-85% confluency, media containing Ac 4GlcNAc,
Ac
4GlcNAz, Ac 4GlcNAlk, Ac 36AzGlcNAc, Ac36AlkGlcNAc (1,000 x stock in DMSO),
or DMSO vehicle was added as indicated. For chase experiments, existing media was
replaced with media supplemented with 200 μM Ac
4GlcNAc (Sigma) or 200 μM
Ac
4GlcNAc (Sigma) plus 10 μM Thiamet-G (1,000 x stock in DMSO) as indicated.
Preparation of Nonidet P-40 (NP-40)-Soluble Lysates. The cells were collected by
trypsinization and pelleted by centrifugation at for 4 min at 2,000 x g, followed by
washing 2x with PBS (1 mL). Cell pellets were then resuspended in 100 μL of 1% NP-40
lysis buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with
Complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Roche) for 20 min and
then centrifuged for 10 min at 10,000 x g at 4 ℃. The supernatant (soluble cell lysate)
was collected and the protein concentration was determined by BCA assay (Pierce,
ThermoScientific).
Cu(I)-Catalyzed [3 þ 2] Azide-Alkyne Cycloaddition. Cell lysate (200 μg) was diluted
with cold 1% NP-40 lysis buffer to obtain a desired concentration of 1 μg/μL.
Newly-made click chemistry cocktail (12 μL) was added to each sample
[alkynyl-rhodamine tag (100 μM, 10 mM stock solution in DMSO);
tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (1 mM, 50 mM freshly prepared
stock solution in water); tris[(1-benzyl-1-H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (100
μM, 10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared
145
stock solution in water) for a total reaction volume of 200 μL. The reaction was gently
vortexed and allowed to sit at room temperature for 1 h. Upon completion, 1 mL of ice
cold methanol was added to the reaction, and it was placed at -20 ℃ for 2 h to precipitate
proteins. The reactions were then centrifuged at 10,000 x g for 10 min at 4 ℃. The
supernatant was removed, the pellet was allowed to air dry for 15 min, and then 50 μL
4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) was added to each
sample. The mixture was sonicated in a bath sonicator to ensure complete dissolution,
and 50 μL of 2x SDS-free loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4%
β-mercaptoethanol, pH 6.8) was then added. The samples were boiled for 5 min at 97 ℃,
and 40 μg of protein was then loaded per lane for SDS-PAGE separation (Any Kd,
Criterion Gel, Bio-Rad).
In-Gel Fluorescence Scanning. Following SDS-PAGE separation, gels were scanned on
a Typhoon 9400 Variable Mode Imager (GE Healthcare) using a 532 nm for excitation
and 30 nm bandpass filter centered at 610 nm for detection.
Reverse Transcriptase-PCR. Information about primers used for GALK2 and GAPDH
available upon request. RNA from NIH3T3 cells was isolated using the RNAeasy Kit
(Qiagen). Concentrations of RNA were obtained by UV-Vis. PCR was conducted in an
Eppendorf Mastercycler thermocycler. To a 0.2 mL thermo-walled PCR tube was added
2X Reaction Mix (SuperScript™ One-Step RT-PCR with Platinum® Taq, Invitrogen),
template RNA from NIH3T3 cells (1,000 ng), sense and anti-sense primers for GALK2
146
(10 μM), sense and anti-sense primers for GAPDH (10 μM), water, and Taq enzyme
(SuperScript™ One-Step RT-PCR with Platinum
®
Taq, Invitrogen). The provided PCR
cycle was used according to SuperScript™ One-Step RT-PCR with Platinum® Taq
(Invitrogen ) with an extension time of 32 sec (1 min/kbp). Products were diluted with 6X
sample loading dye (Bio-Rad) and analyzed by electrophoresis on a 5% agarose gel (500
mg agarose in 1X TAE buffer, tris/acetic acid/EDTA, Bio-Rad). The gel was subsequently
visualized using a ChemiDoc XRS+ molecular imager (Bio-Rad).
GalNAc Kinase 2 (GalK2) Assay. Recombinant human GalK2 was prepared as
previously described.(Bourgeaux et al., 2005) Recombinant GalK2 (8 μg mL
−1
) was
incubated in triplicate with GalNAc, GlcNAc or 6AzGlcNAc (40 mM) in 25 μL reaction
buffer (10 mM MgCl
2, 50 mM Tris HCl pH 8.0) containing 1 mg/mL BSA and 5 mM
[
32
P]ɣATP (1000 cpm/nmol) for 60 min at 37 °C. After this time, reactions were
terminated by the addition of water (0.75 mL) and applied to a Dowex 1 x 8 (Cl-) column
(0.7 cm x 3.0 cm). Unreacted starting material was eluted by washing with 2 mL of 25
mM NH
4HCO 3 before eluting the sugar-1-phosphates with 100 mM NH 4HCO 3. The
fractions (2 mL) were counted in a liquid scintillation counter (μBeta, Perkin-Elmer).
Controls without acceptor substrates were treated in the same way. Column profiles were
compared to detect the presence of overlapping radioactive peaks corresponding to
degradation products. If present, these peaks were subtracted from the assay
chromatogram.
147
Expression of phosphoacetylglucosamine mutase (AGM1). Homo sapiens
phosphoacetylglucosamine mutase 3 (AGM1/PGM3) cDNA was obtained from Biovalley
(Marne-la-Vallée, France), amplified by PCR, sequenced and cloned in pTrcHis A
(Invitrogen)). The 6His-tagged PGM3 protein was expressed in Escherichia coli DH5α
(Invitrogen) cultured for 24 h at 18 °C in 2 TY medium supplemented with 1 mM IPTG
and 2 mM MgCl
2. Bacteria were lysed in Y-Per (ThermoScientific) and the lysate diluted
with 5 volumes of 50 mM phosphate buffer pH 8.0, 300 mM NaCl, 10 mM imidazole,
0.1 mM PMSF and 0.1 mM TCEP. After application of the lysate on a HisTrap FF 5mL
column (GE Healthcare), AGM1 was eluted with 250 mM imidazole in 25 mM phosphate
buffer pH 8.0, 150 mM NaCl, 0.1 mM TCEP. AGM1 activity was checked in a coupled
assay by 2 h incubation at 37 °C with GlcNAc-6-phosphate (2 mM), in 75 mM Tris HCl
pH 8.8, 5 mM MgCl
2, containing 0.1 mg BSA and 2 µL of the PGM3 enzyme solution,
and coupling with AGX1 (0.6−6.3 μg mL
−1
, 2.2−22 mU mL
−1
) and yeast inorganic
pyrophosphatase (1.6 μg mL
−1
, 3 U mL
−1
) in the presence of [
3
H]UTP (Perkin-Elmer, 2
mM, 260 cpm nmol
−1
). After AGM1/AGX1/yeast inorganic phosphatase heat
denaturation, calf intestinal alkaline phosphatase (NEB, 80 U mL
−1
) was added in order
to hydrolyze all the UTP and UDP present, and the products were monitored as described
below for AGX1 enzymatic tests (separation on paper chromatography). Finally, the
production of UDP-GlcNAc was estimated by scintillation counting. Under these
conditions, the recombinant AGM1 activity was estimated to 20 nmol h
−1
µg
−1
of the
AGM1 enzyme solution.
148
Phosphoacetylglucosamine mutase (AGM1) Assay. 6AzGlcNAc (2.25 mM final
concentration) was incubated in 100 µL of 75 mM Tris HCl pH 8.8, 5 mM MgCl
2,
containing 0.1 mg BSA and 10 µg of the PGM3 enzyme solution. Incubations were run at
37°C for 1 h 30 min either without cofactor or in the presence of Glc-6P, Glc1,6-diP or
GlcNAc-6P (1.00 mM final concentration). Reactions were stopped by freezing at –20
°C. After thawing, the reaction mixtures were incubated for 1 h at 37 °C with
bicyclo[6.1.0]nonyne-(POE)
3-NH-Dye 495 conjugate (Synaffix, Oss, Netherlands). The
reaction mixtures which contain the azido sugars coupled through their azido moiety to
the BCN fluorescent Dye
495, were then laid onto a 46 x 57 cm sheet of Whatman 3MM
Paper (GE Healthcare) and run for descending chromatography in ethyl acetate/formic
acid/water (70:20:10) for 5 h. After drying, fluorescent spots (Rf ~ 0.8) were cut out and
the products eluted from the paper in 50% methanol. They were further concentrated
under vacuum before mass spectrometry analysis.
UDP-GalNAc pyrophorylase (AGX1) Assay. Recombinant human AGX1 was prepared
as previously described.(Bourgeaux et al., 2005) Recombinant AGX1 (0.6-6.3 μg mL
−1
,
2.2-22 mU mL
−1
) was incubated with GlcNAc-1P or 6AzGlcNAc-1P in 25 μL reaction
buffer (1 mM MgCl
2, 75 mM Tris HCl pH 8.8) containing 1 mg mL
−1
BSA and 2 mM
[
3
H]UTP (260 cpm/nmol) for 10 min at 37 °C. Yeast inorganic pyrophosphatase (1.6 μg
mL
−1
, 3 mU mL
−1
) was also added to inhibit the reverse reaction. Reactions were
terminated by heating for 6 min at 80 °C. To hydrolyze excess UTP and UMP, calf
intestinal alkaline phosphatase (New England Biolabs, 80 U/mL) was added to the
149
reaction mixture and incubated for 2 hours at 37 °C. The reaction was spotted onto
Whatman 3MM chromatography paper and submitted to descending chromatography in
ethyl acetate/formic acid/water (70:20:10) to remove [
3
H]Uridine (R f = 0.25). Spots with
[
3
H]UDP-sugars (R f = 0.01) were cut out of the chromatography paper and counted in a
liquid scintillation counter. Control samples without sugar-1P received the same
treatment in order to deduct background radioactivity. All tests were run in triplicates and
each experiment was repeated three times. K
m and K cat were calculated using the Enzyme
Kinetic Module 1.3 of SigmaPlot 10, from plots obtained with different concentrations of
sugar-1P and different amounts of AGX1.
MTS Assay. NIH3T3 cells were pretreated with 200 μM Ac
36AzGlcNAc or DMSO for
72 hrs prior to plating. NIH-3T3 cells (1 x 10
4
cells) were plated per well in a 96-well,
white bottom dish 24 hours before treatment with 200 μM Ac
4GlcNAc or
Ac
36AzGlcNAc for 16 hours in triplicate. CellTiter 96
®
AQueous Non-Radioactive Cell
Proliferation Assay (Promega, Madison, WI) was used according to the provided
protocol. Absorbance at 490 nm was read using a BioTek Synergy H4 Multi-Mode
Microplate reader.
Flow Cytometry of Cell-Surface Labeling with DBCO-Biotin. NIH3T3 cells grown in
6-well plates at 80-85% confluency were treated with 200 μM Ac
4GlcNAc, Ac 4GlcNAz,
Ac
4GalNAz or Ac 36AzGlcNAc in triplicate for 16 hours at which time media was
removed and cells were gently washed with PBS before being detached from the plate
150
with 1 mM EDTA in PBS. Cells were collected by centrifugation (5 min, 300 x g at 4 ℃)
and were washed three times with PBS (5 min, 300 x g at 4 ℃). Cells were then
resuspended in 200 μL PBS containing DBCO-biotin (Click Chemistry Tools, 60 μM) for
1 h, after which time they were washed three times with PBS (5 min, 300 x g at 4 ℃)
before being resuspended in ice-cold PBS containing fluorescein isothiocynate (FITC)
conjugated avidin (Sigma, 5 μg/mL, 30 mins at 4 ℃). Cells were then washed three times
in PBS (5 min, 300 x g at 4 ℃) before being resuspended in 400 μL PBS for
flow-cytometry analysis. A total of 10,000 cells [dead cells were excluded by treatment
with propdium iodide (2.5 μg/mL in water, 30 mins)] were analyzed on a BD SORP
LSRII Flow Cytometer using the 488 nm argon laser.
FoxO1 Labeling. NIH3T3 cells stably expressing FLAG-tagged FoxO1 were treated
with 200 μM Ac
4GlcNAz, Ac 36AzGlcNAc (1,000x stock in DMSO) or DMSO and
allowed to incubate overnight. After 16 h, cells were washed with PBS, trypsinized and
pelleted. Cell pellets were resuspended in 100 μl of 1% NP-40 lysis buffer [1% NP-40,
150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with Complete Mini, EDTA-free
Protease Inhibitor Cocktail Tablets (Thermo Scientific) for 20 min and then centrifuged at
4 ℃ for 10 min at 10,000 x g. The supernatant was collected and the protein
concentration was determined by BCA assay (Pierce, ThermoScientific). Total cell lysate
(1.5 mg) was diluted as necessary to a final volume of 1 mL with 1% NP-40 buffer with
Complete Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Thermo Scientific).
EZview Red ANTI-FLAG M2 affinity beads (30 μL, Sigma), pre-washed with cold
151
NP-40 buffer 2X followed by cold PBS 2x, were added to each sample. The samples
were placed on a rotator for 2 h at 4 ℃. Beads were collected by centrifugation at 2,000 x
g for 2 min at 4 ℃, and the supernatant was carefully removed. Beads were then washed
with cold PBS by rotating for 5 mins before centrifuging 2 mins at 2,000 x g. The final
PBS wash was carefully removed, and the beads were suspended in 40 μL 4% SDS buffer
and boiled for 5 min at 97 ℃. The appropriate amount of click chemistry cocktail was
added, and the reaction was allowed to proceed for 1 h after which time 30 μL of 2x
loading buffer was added. Samples were boiled for 5 minutes at 97 ℃. Protein samples
(40 μg) were then loaded per lane for SDS-PAGE separation (Any Kd Criterion Gel,
Bio-Rad) and imaged by in-gel fluorescence scanning.
GlyCAM-IgG Labeling. NIH3T3 cells stably expressing GlyCAM-IgG in 6-well dishes
at 80-85% confluency were treated in DMEM with 10% FCS and 200 μM Ac
4GlcNAz,
Ac
36AzGlcNAc (1,000x stock in DMSO) or DMSO for 24 hours. The media from each
sample was collected by centrifugation at 3,000 x g for 10 min at 4 ˚C to remove cell
debris. The supernatant (1mL) was incubated with 50 μL of recombinant protein G
sepharose beads (Invitrogen) in 100 mM TEA pH 8 overnight. Beads were collected by
centrifugation at 2,000 x g for 2 min at 4 ℃. Beads were washed 3x with 1 mL 100 mM
TEA pH 8. GlyCAM-Ig was eluted by addition of 50 μL 4% SDS buffer (4% SDS, 150
mM NaCl, 50 mM TEA pH 7.4) and boiling for 5 min at 97 ˚C. Protein concentration was
determined by BCA assay (ThermoScientific). Final SDS concentration was diluted to
0.5% by addition of 50 mM TEA pH 7.4. The appropriate amount of click chemistry
152
cocktail was added and the reaction was allowed to proceed for 1 h after which time 4x
loading buffer (200 mM Tris HCl, 4% SDS, 40% glycerol, 0.4% bromophenol blue, 1.4%
β-mercaptoethanol, pH 6.8) was added. Samples were boiled for 5 min at 97 ˚C and 50
μg were loaded for SDS-PAGE separation (Any Kd Criterion Gel, Bio-Rad).
Western Blotting. Proteins were separated by SDS-PAGE before being transferred to
PVDF membrane (Bio-Rad) using standard western blotting procedures. All western
blots were blocked in TBST (0.1% Tween-20, 150 mM NaCl, 10mM Tris pH 8.0)
containing 5% non-fat milk for 1 h at rt. The blots were then incubated with the
appropriate primary antibody in blocking buffer for 1 h at rt. The anti-FLAG antibody
(Thermo) and anti-MAb414 antibody (Covance) were used at a 1:5,000 dilution and
1:1,000 for detection of Foxo1A and p62, respectively. The anti-Nedd4 antibody
(Millipore) was used at a 1:10,000 dilution to detect Nedd-4 and the anti-Pyruvate kinase
antibody (Abcam) was used at 1:1,000. The blots were then washed three times in TBST
for 10 min and incubated with the horseradish peroxidase (HRP)-conjugated secondary
antibody for 1 h in blocking buffer at rt. HRP-conjugated anti-mouse, anti-rabbit,
anti-goat and anti-human antibodies (Jackson ImmunoResearch) were used at 1:10,000
dilutions. After being washed three more times with TBST for 10 min, the blots were
developed using ECL reagents (Bio-Rad) and the ChemiDoc XRS+ molecular imager
(Bio-Rad).
153
Biotin Enrichment and On-bead Trypsinolysis. NIH3T3 cell-pellets labeled with
Ac
36AzGlcNAc, Ac 3GlcNAz, Ac 3GalNAz or Ac 4GlcNAc for 16 hours were resuspended
in 200 μL H
2O, 60 μL PMSF in H 2O (250 mM), and 500 μL 0.05% SDS buffer (0.05%
SDS, 10 mM TEA pH 7.4, 150 mM NaCl) with Complete Mini protease inhibitor
cocktail (Roche Biosciences). To this was added 8 μL Benzonase (Sigma), and the cells
were incubated on ice for 30 min. Then, 4% SDS buffer (2000 μL) was added, and the
cells were briefly sonicated in a bath sonicator followed by centrifugation (20,000 x g for
10 min at 15 °C). Soluble protein concentration was normalized by BCA assay (Pierce,
ThermoScientific) to 1 mg/mL, and 10 mg of total protein was subjected to the
appropriate amount of click chemistry cocktail containing alkyne-PEG3-biotin (5 mM,
Click Chemistry Tools) for 1 h, after which time 10 volumes of ice-cold MeOH were
added. Precipitation proceeded 2 hours at -20 °C. Precipitated proteins were centrifuged
at 5,200 x g for 30 min at 0 ℃ and washed 3 times with 40 mL ice-cold MeOH, with
resuspension of the pellet each time. The pellet was then air-dried for 1 h. To capture the
biotinylated proteins by streptavidin beads, the air-dried protein pellet was resuspended in
2 mL of resuspension buffer (6 M urea, 2 M thiourea, 10 mM HEPES pH 8.0) by bath
sonication. To cap cysteine residues, 100 μl of freshly-made TCEP (200 mM stock
solution, Thermo) was then added and the mixture incubated for 30 min, followed by 40
μl of freshly prepared iodoacetamide (1 M stock solution, Sigma) and incubation for a
further 30 min in the dark. Steptavadin beads (250 μL of a 50% slurry per sample,
Thermo) were washed 2x with 1 mL PBS and 1x with 1 mL resuspension buffer and
resuspended in resuspension buffer (200 μL). Each sample was combined with
154
streptavadin beads and incubated on a rotator for 2 h. These mixtures were then
transferred to Mini Bio-Spin
®
columns (Bio-Rad) and placed on a vacuum manifold.
Captured proteins were then washed with agitation 5x with resuspension buffer (10 mL),
5x PBS (10 mL), 5x with 1% SDS in PBS (10 mL), 30x with PBS (1 mL per wash,
vacuum applied between each wash), and 5x 2M urea in PBS (1 mL per wash, vacuum
applied between each wash). Beads were then resuspended in 2 M urea in PBS (1 mL),
transferred to screw-top tubes, and pelleted by centrifugation (2000 x g for 2 min). At this
time, 800 μL of the supernatant was removed, leaving a volume of 200 μL. To this
bead-mixture was added 2 μL of CaCl
2 (200 mM stock, 1 mM final concentration) and 2
μL of 1 mg/mL sequence grade trypsin (Promega) and incubated at 37 ℃ for 18 hours.
The resulting mixtures of tryptic peptides and beads were transferred to Mini Bio-Spin
®

columns (Bio-Rad) and the eluent was collected by centrifugation (1,000 x g for 2 min).
Any remaining peptides were eluted by addition of 100 μL of 2 M urea in PBS followed
by centrifugation as immediately above. The tryptic peptides were then applied to C18
spin columns (Pierce) according to manufacturer's instructions, eluted with 70%
acetonitrile in H
2O, and concentrated to dryness on a speedvac.
LC-MS Proteomics Analysis. Peptides were desalted on a trap column following
separation on a 12cm/75um reversed phase C18 column (Nikkyo Technos Co., Ltd.
Japan). A 3 hour gradient increasing from 10% B to% 45% B in 3 hours (A: 0.1% Formic
Acid, B: Acetonitrile/0.1% Formic Acid) was delivered at 150 nL/min. The liquid
chromatography setup (Dionex, Boston, MA, USA) was connected to an Orbitrap XL
155
(Thermo, San Jose, CA, USA) operated in top-5-mode. Acquired tandem MS spectra
(CID) were extracted using ProteomeDiscoverer v. 1.3 (Thermo, Bremen, Germany) and
queried against the human Uniprot protein database using MASCOT 2.3.02
(Matrixscience, London, UK). Peptides fulfilling a Percolator calculated 1% false
discovery rate threshold were reported. All LC-MS/MS analysis were carried out at the
Proteomics Resource Center at The Rockefeller University, New York, NY , USA.
156
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Chapter 3. A New Chemical Reporter of O-GlcNAc Reveals
Modification of The Apoptotic Caspases That Blocks The
Cleavage/Activation of Caspase-8
†
Introduction
O-GlcNAc modification is found on serine and threonine side-chains of proteins
throughout the cytosol, nucleus, and mitochondria of animal and plant cells (Figure
3-1A).(Zachara and Hart, 2002) Unlike many forms of cell surface glycosylation, it only
consists of the addition of the single monosaccharide N-acetyl-glucosamine that is not
elaborated by any additional carbohydrates. O-GlcNAcylation is also dynamic through
action of the enzyme O-GlcNAc transferase (OGT) that adds the modification and
subsequent removal by the enzyme O-GlcNAcase (OGA).(V ocadlo, 2012) Genetic
experiments have demonstrated that O-GlcNAcylation is required for development in
mice(O'Donnell et al., 2004; Shafi et al., 2000; Yang et al., 2012) and Drosophila(Sinclair
et al., 2009) and for survival of mammalian cells in culture. Additionally, changes in the
overall cellular levels of this modification directly contribute to a variety of diseases.
O-GlcNAcylation levels are consistently elevated in all types of cancer when compared
to healthy tissue.(Ma and V osseller, 2013) In contrast, genetic loss of OGT in mouse
neurons results in hyperphosphorylated tau,(O'Donnell et al., 2004) and a combination of
†
Anna R. Batt, Balyn W. Zaro,

Narek Darabedian, Nicholas P. Marotta,

Caroline K. Brennan, and Arya
Amirhekmat (University of Southern California) contributed to the work presented in this chapter.
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biochemical and pharmacological experiments has demonstrated that O-GlcNAcylation
blocks the aggregation of the neurodegenerative disease associated proteins tau(Yuzwa et
al., 2008; 2012) and α-synuclein.(Marotta et al., 2012; 2015) The general role for
O-GlcNAcylation in the modulation of cell survival during stress potentially links these
two disease observations. The overall levels of O-GlcNAcylation are significantly
elevated by a variety of cellular stressors in both culture and in vivo models, and these
increased modification levels promote the survival of both cells and tissue.(Darley-Usmar
et al., 2012; Groves et al., 2013) Notably, genetic and biochemical experiments have
implicated O-GlcNAcylation as a inhibitor of apoptosis. For example, specific genetic
deletion of OGT in T-cells caused a large amount of apoptosis in both CD4+ and CD8+
cells,(O'Donnell et al., 2004) and treatment of pancreatic cancer cell-lines with a small
molecule inhibitor of OGT resulted in the induction of apoptosis.(Ma et al., 2013)
However, the previously identified roles for O-GlcNAcylation in apoptosis that could
explain these results are somewhat indirect: O-GlcNAcylation can drive the expression of
heat-shock proteins,(Zachara, 2004) glycosylated phosphofructokinase and
glucose-6-phosphate dehydrogenase have altered activities, resulting in the production of
NADPH,(Yi et al., 2012) and O-GlcNAcylated NFκB has increased transcriptional
activity.(Ma et al., 2013)
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Figure 3-1. O-GlcNAcylation and the major apoptotic caspases. (A) O-GlcNAcylation is the
reversible addition of the monosaccharide N-acetyl-glucosamine to serine and threonine
side-chains of proteins in the cytosol, nucleus and mitochondria. (B) Apoptosis is carried out by
several caspase proteases, including 3, 8 & 9. The caspases are translated as inactive zymogens
(pro-caspases) that are activated by cleavage at specific sites, directly after certain aspartic acid
(D) residues. Caspases-8 & -9 activate themselves and subsequently activate caspase-3.
While these pathways certainly contribute to cell survival, we were interested in
determining if any of the direct components of the apoptotic machinery are affected by
O-GlcNAcylation as a more immediate mechanism to inhibit cell death. Apoptosis in
mammalian cells is regulated by the activation of a family of cysteine proteases termed
caspases.(Fuchs and Steller, 2015; Li and Yuan, 2008) These enzymes are translated as
inactive zymogens (pro-caspases), preventing the uncontrolled activation of cell death.
Upon certain stimuli the caspases are activated by proteolysis in two different pathways
(Figure 3-1B). The intrinsic pathway begins with release of mitochondrial proteins,
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including cytochrome c, into the cytosol. This results in the formation of a protein
complex that contains multiple copies of pro-caspase-9. The proteolytic activity of
caspase-9 is increased in this complex, resulting in its self-cleavage and activation.
Caspase-9 then cleaves and activates the effector caspases, including caspase-3, which
then cleave hundreds of specific substrates resulting in cell death.(Crawford and Wells,
2011) The second pathway, called the extrinsic pathway, is typically initiated from
outside the cell through the engagement of death receptors by appropriate ligands. This
leads to a similar activation and cleavage of caspase-8 that will then activate the effector
caspases. The caspase family members have different substrate preferences but all require
an aspartic acid (D) residue immediately N-terminal to the cleavage site and prefer small
amino acids immediately C-terminal.(Pop and Salvesen, 2009) In addition to the
caspases, there are other proteins that play key roles in apoptosis, including numerous
receptors, scaffolding proteins, and inhibitors and activators of the pathway.
Here, we describe a chemical-proteomics approach that enabled the discovery of
O-GlcNAcylation on all three of the major apoptotic caspases (3, 8, and 9). To make this
discovery, we developed a new, metabolic chemical reporter (MCR), termed
Ac 36AlkGlcNAc, that shows selectivity for labeling O-GlcNAcylated proteins in
mammalian cells. This MCR builds upon our previously published reporter
Ac 36AzGlcNAc,(Chuh et al., 2014) but displays improved detection capabilities because
of a superior a signal-to-noise ratio due to the reverse orientation of the copper-catalyzed
azide-alkyne cycloaddition CuAAC chemistry.(Speers and Cravatt, 2004) When
combined with the copper(I)-catalyzed azide-alkyne cycloaddition, 6AlkGlcNAc enables
the visualization of labeled proteins in as little as one hour and the proteomic
identification of 469 potentially O-GlcNAcylated proteins in embryonic mouse fibroblast
cells, including caspases-3 and -8. We subsequently used biochemical methods to confirm
O-GlcNAcylation of all three of the major apoptotic caspases in both mouse and human
cell lines. We then explored the consequences of modification on caspase-8. Using
small-molecule inhibitors to raise and lower global O-GlcNAcylation levels, we find that
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they are inversely related to the kinetics of caspase-8 activation during apoptosis. Finally,
we demonstrate that caspase-8, a key initiator of the extrinsic apoptotic pathway, is
modified at its cleavage/activation sites and show that these modifications inhibit
cleavage in vitro and potentially in living cells. Collectively, these results provide an
advanced MCR for the discovery of potentially O-GlcNAcylated proteins and provide,
for the first time, a potential role for O-GlcNAc in the direct inhibition of caspase
activation. We predict that this caspase modification may be a function of
O-GlcNAcylation in both development and human disease.
Results
Development of a selective and robust metabolic chemical reporter for the analysis of
O-GlcNAcylated proteins.
We and others have demonstrated that the majority of O-GlcNAcylation MCRs are not
selective for O-GlcNAc and label other types of glycoproteins. While this does not
preclude their use for the characterization of proteins, it does limit the number of
potentially O-GlcNAcylated proteins that can be identified in a proteomics experiment, as
the enrichment of abundant cell-surface glycoproteins can mask the presence of
O-GlcNAcylated proteins.(Chuh et al., 2014) We serendipitously overcame this
selectivity issue with our previously published MCR (6AzGlcNAc, Figure 3-2A), which
was excluded from cell-surface glycoproteins but labeled O-GlcNAcylation
substrates.(Chuh et al., 2014) We chose to explore a 6-modified MCR because
UDP-6AzGlcNAc is a substrate for OGT in vitro(Mayer et al., 2011) and because a
small-molecule glycoside of 6AzGlcNAc can be hydrolyzed by OGA.{Kim:2017cy}
Notably, we also demonstrated in vitro that Ac
36AzGlcNAc can bypass the canonical
metabolic salvage pathway by transformation to the 6AzGlcNAc-1-phosphate by the
enzyme phosphoacetylglucosamine mutase (AGM1).(Chuh et al., 2014) As mentioned
above, azido-probes in combination alkyne-tags under CuAAC has increased background
signal due to non-specific reactivity of alkyne-tags with proteins,(Speers and Cravatt,
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2004) which reduces the number of proteins that can be identified with confidence.
Therefore, we synthesized here the analogous alkyne-bearing MCR Ac 36AlkGlcNAc
(Figure 3-2A, Scheme 3-1) in 8 synthetic steps from commercially-available material.
With this potential MCR in hand, we first compared it to Ac 36AzGlcNAc, by treating
NIH3T3 cells with either reporter or DMSO vehicle for 16 h. Analysis by in-gel
fluorescence, following lysis and CuAAC with the appropriate fluorescent tag,
demonstrated that Ac 36AlkGlcNAc labels a qualitatively similar spectrum of proteins as
Ac 36AzGlcNAc (Figure 2B) but with notably improved signal-to-noise (Figure 3-2B,
High “exposure”).
Figure 3-2. Characterization of the metabolic chemical reporter Ac 36AlkGlcNAc. (A)
Metabolic chemical reporters (MCRs) of O-GlcNAcylation. Treatment of cells with
Ac36AzGlcNAc results in the incorporation of an azide functionality onto O-GlcNAcylated
proteins, which can be subsequently reacted with alkyne tags using bioorthogonal chemistry.
Ac36AlkGlcNAc was hypothesized to function in the same way but with improved
signal-to-noise. (B) Ac36AlkGlcNAc has improved signal-to-noise compared to Ac 36AzGlcNAc.
NIH3T3 cells were treated with ether DMSO vehicle, Ac36AzGlcNAc (200 μM), or
Ac36AlkGlcNAc (200 μM) for 16 h, followed by CuAAC with the appropriate fluorescent tag
and analysis by in-gel fluorescence scanning. The data is representative of two biological
replicates.
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Scheme 3-1. Synthesis of Ac 36AlkGlcNAc. Reagents: (a) dimethoxypropane, p-toluenesulfonic
acid, 1,4-dioxane, 80 °C, 24 h, 63%; (b) 80% acetic acid, 40 °C, 2 h, 81%; (c) methanesufonyl
chloride, pyridine, -20 °C, 2 h, 73%; (d) NaCN, tetrabutylammonium bromide, DMF, 16 h, 86%;
(e) i) diisobutylaluminum hydride, CH2Cl 2, -78 °C, 1 h; ii) 1 M HCl:MeOH (1:1), 1 h, 62%; (f)
dimethyl-(1-diazo-2-oxopropyl)-phosphonate, K2CO 3, MeOH, 3 h, 75%; (g) i) 2.5 M HCl, reflux,
2 h; ii) Ac2O, dimethylaminopyridine, pyridine, 16 h, 58%.
We next determined whether it is able to label proteins in different mammalian cell-lines.
Accordingly, a small panel of cells was treated with either Ac
36AlkGlcNAc (200 μM) or
DMSO vehicle for 16 h. The cells were then lysed and subjected to CuAAC conditions
with azido-rhodamine. In-gel fluorescence scanning showed a diversity of modified
proteins in all the cell-lines tested (Figure 3-2), demonstrating the general applicability of
Ac
36AlkGlcNAc to different mammalian cell-lines. We also determined the kinetics of
Ac
36AlkGlcNAc labeling and compared these results to Ac 36AzGlcNAc, as it is our
other selective MCR for O-GlcNAcylation. NIH3T3 cells were treated with
Ac
36AlkGlcNAc (200 μM) or Ac 36AzGlcNAc (200 μM) for different lengths of time,
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followed by CuAAC and analysis by in-gel fluorescence. Notably, Ac 36AlkGlcNAc
displayed better labeling kinetics due to both better overall incorporation levels and lower
signal to noise (Figure 3-2). This allows O-GlcNAcylated proteins to be robustly labeled
in as little as 30 to 60 min (Figure 3-2), which may enable a range pulse experiments to
isolate newly O-GlcNAcylated proteins in the future. Next, we compared
Ac
36AlkGlcNAc to Ac 4GlcNAlk, since they both take advantage of the same CuAAC
orientation (i.e., alkyne probe and azide tag). NIH3T3 cells were first treated with
Ac
36AlkGlcNAc (200 μM) or Ac 4GlcNAlk (200 μM) for 24 h. Aliquots of the cells were
collected after different lengths of time and analyzed by CuAAC with azido-rhodamine
and in-gel fluorescence. As expected, both MCRs show time-dependent labeling of cells;
however, Ac
4GlcNAlk labels the cells faster than Ac 36AlkGlcNAc (Figure 3-2). These
data may reflect a difference in the metabolism of the two MCRs to the corresponding
UDP-sugar donors, and suggests that there may be a tradeoff between labeling selectivity
and kinetics. We also examined the reversibility of Ac
36AlkGlcNAc-labeling by
performing a pulse-chase experiment. NIH3T3 cells were treated with either
Ac
36AlkGlcNAc (200 μM) or Ac 4GlcNAlk (200 μM) for 16 h. At this time, the cells
were washed and fresh media containing Ac
4GlcNAc (200 μM) was added. Cells were
collected after different lengths time and analyzed by in-gel fluorescence after CuAAC
(Figure 3-2). Fluorescent signal from both of the MCRs was steadily lost at qualitatively
the same rate, consistent with our previous results that these classes of MCR are
enzymatically removed by OGA(Chuh et al., 2014). Finally, we compared the
dose-dependence of Ac
36AlkGlcNAc labeling. Again, NIH3T3 cells were treated with
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different concentrations of either Ac 36AlkGlcNAc or Ac 4GlcNAlk for 16 h, followed by
CuAAC and in-gel fluorescence (Figure 3-2). Notably, Ac
36AlkGlcNAc-dependent signal
reached a maximum at ~200 μM concentration, while labeling by Ac
4GlcNAlk did not
saturate under these conditions. Again this suggests that their is likely a trade-off between
O-GlcNAc selectivity and overall labeling levels.
Figure 3-3. Ac 36AlkGlcNAc can label proteins in a range of mammalian cell lines. The
indicated cell lines were treated with DMSO vehicle or Ac36AlkGlcNAc (200 μM) for 16 h,
followed by CuAAC with azido-rhodamine and analysis by in-gel fluorescence scanning. The
data is representative of two biological replicates.
We next determined whether Ac 36AlkGlcNAc is able to label proteins in different
mammalian cell-lines. Accordingly, a small panel of cells was treated with either
Ac
36AlkGlcNAc (200 μM) or DMSO vehicle for 16 h. The cells were then lysed and
subjected to CuAAC conditions with azido-rhodamine. In-gel fluorescence scanning
showed a diversity of modified proteins in all the cell-lines tested (Figure 3-3),
demonstrating the general applicability of Ac
36AlkGlcNAc to different mammalian
cell-lines. We then characterized the dose-dependence and kinetics of Ac
36AlkGlcNAc
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labeleing and compared it directly to our two previously published MCRs,
Ac
36AzGlcNAc(Chuh et al., 2014) and Ac 4GlcNAlk(Zaro et al., 2011b) (Figure 3-4 and
Figure 3-5). These data show that Ac
36AlkGlcNAc labels proteins with faster kinetics
than Ac
36AzGlcNAc (Figure 3-4A), which combined with the low background signal
enables proteins to be robustly labeled in as little as 1 h (Figure 3-4B). Additionally,
Ac
36AlkGlcNAc is incorporated more efficiently than Ac 36AzGlcNAc (Figure 3-4C).
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Figure 3-4. Ac 36AlkGlcNAc displays improved labeling compared to Ac 36AzGlcNAc. (A)
Ac36AlkGlcNAc labeling is faster that the corresponding modification by Ac 36AzGlcNAc.
NIH3T3 cells were treated with Ac36AlkGlcNAc (200 μM) or Ac 36AzGlcNAc (200 μM) for the
indicated lengths of time. The corresponding lysates were then subjected to CuAAC with the
appropriate rhodamine-tag and analyzed by in-gel fluorescence scanning. (B) Treatment with
Ac36AlkGlcNAc enables protein-labeling in less than 1 hour. NIH3T3 cells were treated with
Ac36AlkGlcNAc (200 μM) for the indicated lengths of time, followed by CuAAC with
azido-rhodamine and in-gel fluorescence scanning. The data in both panels is representative of
two biological replicates. (C) Ac36AlkGlcNAc labeling is more efficient than Ac 36AzGlcNAc
incorporation. NIH3T3 cells were treated with the indicated concentrations of Ac36AlkGlcNAc or
Ac36AzGlcNAc for 16 h, followed by CuAAC with the appropriate rhodamine-tag and in-gel
fluorescent scanning.
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Ac 36AlkGlcNAc labeled cells with slightly slower kinetics compared to Ac 4GlcNAlk
(Figure 3-5A) and at lower overall levels (Figure 3-5B). Finally, we looked at the
turnover of Ac
36AlkGlcNAc-dependent labeling in a pulse-chase experiment (Figure
3-5C). The associated fluorescent signal was steadily lost over the course of 48 h at a
comparable rate to Ac
4GlcNAlk-labeling. While some of this signal loss is undoubtably
due to protein degradation, the similarity in the kinetics suggests that 6AlkGlcNAc, like
GlcNAlk and 6AzGlcNAc, is also a substrate for OGA.
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Figure 3-5. Global Ac 36AlkGlcNAc labeling is less efficient compared to Ac 4GlcNAlk. (A)
Protein labeling by Ac36AlkGlcNAc is slower than Ac 4GlcNAlk. NIH3T3 cells were treated with
200 μM Ac36AlkGlcNAc or Ac 4GlcNAlk for the indicated times before modified proteins were
subjected to CuAAC with azido-rhodamine and in-gel fluorescent scanning. (B) Ac36AlkGlcNAc
labeling saturates at lower concentrations than Ac4GlcNAlk modification. NIH3T3 cells were
treated with the indicated concentrations of Ac36AlkGlcNAc or Ac 4GlcNAlk for 16 h, followed
by CuAAC with azido-rhodamine and in-gel fluorescent scanning. (C) Ac36AlkGlcNAc and
Ac4GlcNAlk are removed from proteins at similar rates. NIH3T3 cells were treated with 200 μM
Ac36AzGlcNAc or Ac 4GlcNAz for 16 hours at which time media was exchanged for fresh media
containing 200 μM Ac4GlcNAc. Cells were harvested at the indicated times and modified proteins
were subjected to in-gel fluorescent scanning following CuAAC with azido-rhodamine. The data
in all panels is representative of two biological replicates.
We then examined the selectivity of Ac 36AlkGlcNAc labelling of different classes of
glycoproteins by first using two reporter proteins: GlyCAM-IgG, which is a secreted
fusion protein that contains both N-linked and mucin O-linked cell-surface glycans, and
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FoxO1 that is an O-GlcNAcylated transcription factor. As expected, treatment with
Ac
4GlcNAlk as a control resulted in labeling of both proteins (Figure 3-6A), as this MCR
is not excluded from cell-surface glycoproteins. However, when these cells were treated
with Ac
36AlkGlcNAc, we observed no labeling of GlyCAM-IgG but robust labeling of
FoxO1 (Figure 3-6A), indicating that Ac
36AlkGlcNAc may be selectively-labeling
potential O-GlcNAcylation substrates. To confirm these results, we also performed an
analysis of the entire repertoire of cell surface glycans using flow cytometry by taking
advantage of a copper-chelating azido biotin tag that enables CuAAC on the surface of
living cells.(Uttamapinant et al., 2012) More specifically, NIH3T3 cells were treated with
either Ac
36AlkGlcNAc or the non-selective MCRs Ac 4GlcNAlk or Ac 4ManNAlk.
Analysis of cell-surface labeling by flow cytometry demonstrated that Ac
36AlkGlcNAc
treatment resulted in dramatically less global cell-surface labelling (Figure 3-6B),
consistent with its selectivity towards FoxO1 labeling.
179
Figure 3-6. Ac 36AlkGlcNAc is selective for O-GlcNAcylation. (A) Ac 36AlkGlcNAc is
selective for an O-GlcNAcylated reporter protein. NIH3T3 cells expressing either secreted
GlyCAM-IgG or nucleocytoplasmic FoxO1 were treated with DMSO vehicle, Ac36AlkGlcNAc
(200 μM), or Ac4GlcNAlk (200 μM) for 24 h. At this time the proteins were enriched from the
media or soluble cell lysate, respectively, subjected to CuAAC with azido-rhodamine and
analyzed by in-gel fluorescence scanning. The data is representative of two biological replicates.
(B) Ac36AlkGlcNAc is excluded from cell surface glycosylation. NIH3T3 cells were treated with
Ac4GlcNAlk, Ac 4ManNAlk, or Ac 36AlkGlcNAc (all at 200 μM) for 16 h. The cells were then
harvested and subjected to CuAAC with picolyl azido-biotin. After incubation with FITC-biotin,
cell-surface labeling was measured by flow cytometry. Error bars represent ±s.e.m. from the mean
of biological replicates (n = 3), and statistical significance was calculated using a two-tailed
Student’s t-test.
Treatment with Ac 36AlkGlcNAc is not toxic and does not induce apoptosis in mammalian
cells.
Finally, we set out to determine if treatment with MCRs reduces cellular proliferation or
induces apoptosis. To test these possibilities, NIH3T3 cells were treated in triplicate with
either Ac
36AlkGlcNAc, Ac 36AzGlcNAc, Ac4GlcNAlk before analysis using two
different, commercially-available assays: an MTS assay to measure general
cellular-proliferation and a apoptosis assay that measures the catalytic activity of
180
caspase-3 and -7 (Figure 3-7A). Although there appeared to be a small decrease in the
proliferation of the cells, none of the differences were statistically significant and no
induction of apoptosis was observed. We next repeated this experiment in HeLa cells
under Ac
36AlkGlcNAc treatment and observed similar results, with no significant change
in cellular proliferation and no induction of apoptosis (Figure 3-7B). Together these data
demonstrate that Ac
36AlkGlcNAc has the potential to be a selective probe for the
identification of O-GlcNAcylated proteins that is relatively benign to cells.
Figure 3-7. Treatment with Ac 36AlkGlcNAc is not toxic to mammalian cells. (A) NIH3T3
cells were treated with DMSO vehicle or the indicated MCR (200 μM) for 16 h before analysis
by commercially available MTS (Celltiter 96 Aqueous, Non-Radioactive Cell-Proliferation Assay,
Promega) or apoptosis (Caspase-Glo 3/7 Assay, Promega) assays. (A) HeLa cells were treated
with DMSO or Ac36AlkGlcNAc (200 μM) for 16 h before analysis as in (A). Error bars represent
±s.e.m. from the mean of biological replicates (n = 3), and statistical significance was calculated
using a two-tailed Student’s t-test.
Identification of potentially O-GlcNAcylated proteins using Ac 36AlkGlcNAc.
Next, we applied Ac
36AlkGlcNAc to identify potentially modified proteins and
simultaneously compared it to Ac
4GlcNAlk in order to uncover any obvious benefits of
the increased selectivity. Immortalized mouse embryonic fibroblasts (MEFs) were treated
181
in triplicate with either Ac 36AlkGlcNAc, Ac4GlcNAlk, or Ac 4GlcNAc as a control. The
cells were lysed and subjected to CuAAC conditions with an azide-bearing biotin tag.
The proteomes were then reduced, alkylated, and subjected to biotin-enrichment using
streptavidin-conjugated beads. After extensive washing to remove unlabeled proteins,
on-bead trypsinolysis afforded peptides that were analyzed using LC-MS/MS. Labeled
proteins were identified as those that met threshold criteria for enrichment based on
spectral counting, resulting in 469 proteins identified as potential O-GlcNAcylated
proteins that were labeled by Ac
36AlkGlcNAc (Figure 3-8A) and 433 proteins labeled by
Ac
4GlcNAlk (Figure 3-8A), with many of the proteins showing overlap in the two data
sets. Additionally, a large fraction of the identified proteins have been characterized as
being potentially O-GlcNAcylated in past proteomic experiments. Although spectral
counting is inherently a semi-quantitative measurement of protein enrichment, as larger
proteins will produce more peptides, we used it because it allowed us to directly compare
the identified proteins in this study to our previously published data on azide-bearing
MCRs.(Chuh et al., 2014) We first compared the total number of proteins that could be
identified at a statistically-significant level between the different MCRs (Figure 3-8B).
Not surprisingly the alkyne-MCRs outperformed the azide counterparts because of their
improved signal-to-noise ratios, and Ac
36AlkGlcNAc enabled the identification of more
potentially O-GlcNAcylated proteins compared to Ac
4GlcNAlk. We then collated the
overlap between the proteins identified by Ac
36AzGlcNAc, Ac36AlkGlcNAc, and
Ac
4GlcNAlk and found significant overlap between the datasets (Figure 3-8C). We also
annotated the data based on the sub-cellular localization of the identified proteins as
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either intracellular or transmembrane, and therefore potential O-GlcNAc substrates, or
extracellular/lumenal. Proteins without defined localizations were omitted. In our
experiments, Ac
36AlkGlcNAc enabled the identification of the most intracellular proteins
(Figure 3-8D), which have the opportunity to be O-GlcNAcylated. Importantly, the
previous proteomic identification of proteins labeled with azide-containing MCRs was
performed in the same cell line, at identical concentrations and treatment times. Notably,
the apoptotic caspases-3 and -8 were identified as potential O-GlcNAc substrates from
the Ac
36AlkGlcNAc-treated but not Ac 4GlcNAlk-treated cells (Figure 3-8A).
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Figure 3-8. Identification of O-GlcNAcylated proteins, including caspase-3 and -8, using
Ac36AlkGlcNAc. (A) Mouse embryonic fibroblasts were treated with Ac 36AlkGlcNAc,
Ac4GlcNAlk, or Ac 4GlcNAc (all at 200 μM) for 16 h. At this time, the corresponding cell-lysates
were subjected to CuAAC with azido-biotin, incubation with streptavidin-coated beads, and
trypsinolysis. Proteins identified by LC-MS/MS are graphically presented as total number of
positive minus total number of control spectral counts. Three known O-GlcNAcylated proteins
and identified caspases are annotated. (B) Number of proteins identified as being enriched to a
statistically significant amount by MCRs in this study (in green) and a previously published study
(in black). (C) Ven diagram showing the overlap in the proteins identified using Ac36AzGlcNAc,
Ac36AlkGlcNAc, and Ac 4GlcNAlk. (D) Graphical representation of the enriched proteins from
this study (in green) and a previously published experiment (in black) based on their annotated
localization. In all experiments, identical cell lines, MCR concentrations, and treatment times
were used.
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To visualize the enrichment of these caspases and three known O-GlcNAcylated proteins,
the same cells were treated with Ac
36AlkGlcNAc as above, and the labeled proteins were
enriched and selectively eluted using a cleavable azido-azo-biotin tag. Western blotting
confirmed the enrichment of both caspases and the known O-GlcNAcylated substrates
NEDD4-1, pyruvate kinase (PK), and nucleoporin 62 (nup62) (Figure 3-10A).
Importantly, the abundant, non-O-GlcNAcylated protein β-actin was not enriched,
demonstrating the potential O-GlcNAc-selective nature of our enrichment. To confirm
that caspases-3 and -8 are legitimately O-GlcNAcylated, we first used chemoenzymatic
labeling (Figure 3-9).(Clark et al., 2008)
Figure 3-9. Chemoenzymatic labeling of O-GlcNAc modifications. Cell lysates are treated
with both synthetic UDP-GalNAz and the recombinant enzyme GalT(Y298L), resulting in
formation of an azide-containing disaccharide. This disaccharide can then be subjected to
CuAAC with alkyne tags for the installation of visualization or affinity probes.
This method detects endogenous O-GlcNAcylated proteins by using an engineered
galactosyltransferase, GalT(Y289L), to modify O-GlcNAc modifications with the
azide-bearing monosaccharide N-azidoacetyl-galactosamine. The azide of the resulting
disaccharide can then be detected using CuAAC and alkyne visualization- or
affinity-tags. Lysates from MEFs were subjected to chemoenzymatic labeling followed
185
by CuAAC with an alkyne-azo-biotin tag (Click Chemistry Tools #1042) to enable
enrichment of O-GlcNAcylated proteins. Western blotting after elution from streptavidin
beads confirmed that caspase-3 and -8 are indeed endogenously O-GlcNAcylated (Figure
3-10B). We then stably transformed NIH3T3 cells with FLAG-tagged caspase-3 or
caspase-8. Notably, these proteins contained mutations at their active sites to prevent the
autoinduction of caspase activity. Immunoprecipitation of these proteins with anti-FLAG
beads was performed, followed by a second round of enrichment with the lectin wheat
germ agglutinin (WGA) that binds to O-GlcNAc modifications. Western blotting against
the caspases showed WGA enrichment of a fraction of the protein, further indicating that
these proteins are endogenously O-GlcNAcylated (Figure 3-10C). The stable cells were
then treated with Ac
36AlkGlcNAc or DMSO vehicle, followed by immunoprecipitation
of the individual caspases using an anti-FLAG antibody. Subsequent CuAAC with
azido-biotin, followed by analysis by streptavidin blotting confirmed the MCR labeling of
both caspase -3 and -8 (Figure 3-10D). Together these data confirm that caspases-3 and -8
are bona fide O-GlcNAcylated proteins in mouse cells.
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Figure 3-10. Caspases-3 and -8 are genuine O-GlcNAcylated proteins in mouse cells. (A)
Confirmation of Ac36AlkGlcNAc labeling of known O-GlcNAcylated proteins and caspase-3 and
-8. Mouse embryonic fibroblasts were treated with either Ac36AlkGlcNAc (200 μM) or DMSO
for 16 h, followed by CuAAC with a cleavable azido-biotin tag. After enrichment on streptavidin
beads, the labeled proteins were eluted and visualized by Western blotting. The non-glycosylated
protein β-actin is a negative control. (B) Chemoenzymatic labeling identifies caspases-3 and -8 as
O-GlcNAcylated proteins. MEF cell-lysates were subjected to chemoenzymatic modification of
endogenous O-GlcNAcylation sites and then CuAAC with alkyne-azo-biotin. Western blotting
after enrichment and elution confirmed O-GlcNAcylation of caspases-3, 8, and the positive
control Nup62. (C) The WGA lectin enriches caspase-3 and -8. Cell lysates from NIH3T3 cells
that stably express FLAG-tagged caspase-3 or -8 were subjected to two rounds of affinity
enrichment, anti-FLAG followed by WGA lectin, before analysis by Western blotting. (D)
Reverse confirmation of Ac36AlkGlcNAc labeling of caspase-3 and -8. NIH3T3 cells that stably
express FLAG-tagged caspase-3 or -8 were treated with either either Ac36AlkGlcNAc (200 μM)
or DMSO for 16 h, followed by immunoprecipitation with an anti-FLAG antibody, CuAAC with
an azido-biotin tag and analysis by streptavidin and Western blotting. The data in all panels is
representative of two biological replicates.
The major apoptotic caspases are O-GlcNAcylated in human cancer cell-lines.
To confirm caspase O-GlcNAcylation in human cells, we first treated three cancer
cell-lines - H1299 (lung), HeLa (cervical), and MCF7 (breast) - with Ac
36AlkGlcNAc
and enriched the labeled proteins using the cleavable azido-azo-biotin tag. Western
blotting after elution demonstrated MCR labeling of caspase-3 and -8, as well as
187
caspase-9, which we did not detect in our proteomics experiment (Figure 3-11A).
Notably, the non-O-GlcNAcylated protein β-actin was again largely unenriched
compared to inputs in any of the cell lines and caspase-3 was absent from MCF7 cells as
this cell line is caspase-3 null. We then used chemoenzymatic labeling followed by
biotin-enrichment and found that all three caspases are endogenously O-GlcNAcylated in
H1299 cells (Figure 3-11B). We then turned our focus to caspase-8, the initiator caspase
of the exogenous apoptosis pathway, and used chemoenzymatic enrichment to show that
caspase-8 is O-GlcNAcylated in all three cancer cell lines.
Figure 3-11. The apoptotic caspases are O-GlcNAcylated in human cancer cell-lines. (A)
Metabolic chemical reporter enrichment. H1299, HeLa, and MCF7 cells were treated with either
DMSO vehicle or Ac36AlkGlcNAc (200 μM) for 16 h. Cell lysates were then subjected to
CuAAC with the appropriate cleavable biotin-tag, followed by enrichment on streptavidin-coated
beads. After extensive washing and elution (Na2S 2O 4), Western blotting revealed the modification
of caspases-3, -8, and -9. (B) Chemoenzymatic modification and enrichment. H1299 cell-lysates
were subjected to chemoenzymatic modification of endogenous O-GlcNAcylation sites and then
CuAAC with alkyne-azo-biotin. Western blotting after enrichment and elution confirmed
O-GlcNAcylation of caspases-3, 8, and -9. In A and B, enrichment of the known O-GlcNAcylated
protein nucleoporin 62 (Nup62) served as a positive control, and the non-glycosylated protein β-
actin was a negative control.
188
Changes in O-GlcNAcylation levels directly affect the kintetics of caspase-8
cleavage/activation.
We next investigated the consequences of O-GlcNAcylation on caspase-8 biochemistry
by first determining whether the levels of its modification change during apoptosis.
During the initiation of extrinsic apoptosis, monomers of pro-caspase-8 are brought into
close proximity where they are able to cleave themselves to generate the fully active
protease, which then goes on to activate the effector caspases, such as caspase-3 (Figure
3-12A). Typically, this happens through first cleavage after aspartic acid residues (D374
and D394) between the large and small subunits of the catalytic domain, removing the
“activation loop,” followed by cleavage after at least one other aspartic acid residue
(D210, D216, and/or D223) which removes the pro-domain.(Chang et al., 2003)
Accordingly, HeLa cells were treated with tumor necrosis factor alpha (TNFα) to induce
caspase-8-mediated apoptosis, which also caused a very slight increase in the global
levels of O-GlcNAcylation. Western blotting with two different caspase-8 antibodies
clearly showed the time dependent cleavage/activation of caspase-8 based on the
appearance of a lower molecular-weight, activated species (p43/41, Figure 3-12B).
Chemoenzymatic labeling was used to enrich the endogenously O-GlcNAcylated fraction
of caspase-8 in this experiment. This modified protein showed less loss (i.e., cleavage) of
pro-caspase-8 and very little of the activated p43/41 species (Figure 3-12B), indicating
that O-GlcNAcylation of caspase-8 inhibits the cleavage and activation of caspase-8.
Given this result, we next treated HeLa cells with either DSMO, 5SGlcNAc or ThiametG
to alter O-GlcNAcylation levels. These cells were then treated with TNFα and the
189
cleavage and activation of caspase-8 and caspase-3 was visualized by Western blotting
(Figure 3-12C). In all the biological replicates, treatment with 5SGlcNAc resulted in
faster kinetics of caspase activation, while Thiamet-G gave slower rates of activation,
when compared to DMSO treated cells. Importantly, quantitation of the percentage of
cleaved caspase-8 (p43/41 plus p18) from these replicates confirms a statistically
significant difference in the rate of caspase-8 activation that inversely depends on
O-GlcNAcylation levels (Figure 3-12D). Importantly, we did not observe a change in the
overall levels of caspase-8 upon inhibitor treatment. Additionally, the activation of
caspase-8 is through direct engagement of death receptors by TNFα, indicating that the
effects of O-GlcNAcylation are in this pathway, and these observed effects must occur
upstream of NFκB transcriptional activity, as the cells were also treated with
cycloheximide to prevent new protein synthesis. This is in contrast to activation of
apoptosis by a stimulus like cellular stress that could be affected by the established roles
for O-GlcNAc in cellular metabolism and heat shock response.
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Figure 3-12. O-GlcNAcylation levels affect caspase-8 cleavage/activation. (A) The extrinsic
apoptosis pathway. Upon certain signals, inactive, full-length caspase-8 can be recruited to
cell-death receptors where it is oligomerized. Dimers of inactive caspase-8 have increased
activity and self-cleave after aspartic acid residues 374 and 384, removing the activation loop and
yielding activated enzyme (p43/p41 and p10 fragments). The p43/41 fragment can then be further
cleaved to release the death effector domain (DED), resulting in the formation of a p18 fragment.
Active caspase-8 will then go on to cleave the effector caspases, like caspase-3. (B) The
endogenously O-GlcNAcylated fraction of pro-caspase-8 has less cleavage during apoptosis.
HeLa cells were treated with either vehicle or TNFα (10 ng mL
-1
) and cycloheximide (1 μg mL
-1
)
to induce apoptosis. After 6 h, the O-GlcNAcylated proteins were enriched using
chemoenzymatic modification and caspase-8 was visualized by Western blotting. (C) Altering
O-GlcNAcylation levels changes the kinetics of caspase-8-mediated apoptosis. HeLa cells were
treated with TNFα (10 ng mL
-1
) and cycloheximide (CHX, 1 μg mL
-1
) to induce apoptosis in the
presence or absence of either 5SGlcNAc (OGT inhibitor, 100 μM, 16 h pretreatment) or
Thiamet-G (OGA inhibitor, 5 μM, 20 h pretreatment). The kinetics of caspase activation were
then visualized after the indicated times by Western blotting. Active caspases were distinguished
from their corresponding zymogen form by molecular-weight shift. The different amounts of each
caspase species was quantitated and normalized to the total caspase signal in each lane. See
Figure S10 for the other biological replicates. (D) Quantitation of the percentage of active
caspase-8 (p43/41 plus p18) from the experiments in (C) and Figure S10 (SI). Error bars represent
±s.e.m. from the mean of biological replicates (n = 2 for 5SGlcNAc and Thiamet-G, n = 3 for
DMSO), and statistical significance was calculated using a two-tailed Student’s t-test.
Caspase-8 is O-GlcNAcylated near its cleavage/activation sites.
As mentioned above, the caspases are activated upon self- or cross-cleavage after
particular aspartic acid residues. Notably, at or near the cleavage sites required for
191
activation of all three of these caspases are serine (S) or threonine (T) residues that could
be O-GlcNAcylated, which might effect the efficiency of cleavage/activation.
Importantly, these cleavage sites lie in flexible activation loops, and OGT has a strong
preference for unstructured regions of its substrate proteins.(Gloster, 2014; Liu et al.,
2014; Pathak et al., 2015) Here, we focused on these sites (
374
DSE
376
and
384
DLSS
387
;
cleavage after D during activation) in caspase-8 (schematized in Figure 3-13).(Boatright
et al., 2003; Chang et al., 2003) Specifically, H1299 cells were co-transfected in duplicate
with constructs encoding either HA-tagged wild-type or a FLAG-tagged mutant, either
caspase-8(S375A), caspase-8(S386,387A), or caspase-8(AAA) (
374
DAE
376
and
384
DLAA
387
). Importantly, all of these proteins also contained an active-site mutation (C
to A) to prevent the autoinduction of apoptosis observed in some caspase-8
overexpression experiments. The entire pool of endogenously O-GlcNAcylated proteins
from these cells were enriched using chemoenzymatic labeling, followed by CuAAC with
the cleavable biotin tag, enrichment, and elution as above. Western blotting showed that
all three mutants were enriched at significantly lower amounts (normalized to inputs)
compared to the wild-type protein (Figure 3-13). Strikingly, this effect was additive and
caspase-8(AAA) was only enriched at ~10% compared to the wild-type protein,
indicating that essentially all of the O-GlcNAcylation occurs at the cleavage sites.
192
Figure 3-13. Caspase-8 is O-GlcNAcylated near its cleavage/activation sites. Schematic of
wild-type caspase-8 containing two cleavage sites (
374
DSE
376
and
384
DLSS
387
). O-GlcNAcylated
proteins from H1299 cells transfected with both HA-tagged wild-type or FLAG-tagged versions
of the indicated mutant caspase-8 proteins were enriched using chemoenzymatic transfer and
visualized by Western blotting. The HA and FLAG blots were visualized simultaneously to enable
a quantitative comparison between the two blots. Results are the average of two separate
biological experiments that were imaged simultaneously (see Figure S13(SI)). Error bars
represent ± SEM, and statistical significance was calculated using a two-tailed Student’s t-test.
O-GlcNAcylation blocks caspase-8 cleavage/activation in vitro and potentially in living
cells.
To determine if O-GlcNAcylation can directly affect caspase-8 self-cleavage/activation,
we first performed an in vitro cleavage assay. Because caspase-8 proteolyzes itself during
activation, it is able to cleave peptides that correspond to these cleavage sites in vitro. We
synthesized peptides surrounding the cleavage site that were either unmodified (1 and 3)
or O-GlcNAcylated (2 and 4, Figure 3-16A). These peptides were then incubated in
triplicate with recombinant, active human caspase-8 before analysis using reverse-phase
HPLC (RP-HPLC) which separated any cleavage product from the starting peptide, and
193
the absolute identity of the peptides was confirmed by ESI-MS. Caspase-8 was able to
readily cleave both of the unmodified peptides. In contrast, O-GlcNAcylation almost
completely prevented the formation of the cleavage product, with the unmodified
peptides 1 and 3 cleaving approximately 28- and 16-times more efficiently than the
O-GlcNAcylated peptides 2 and 4, respectively (Figure 3-16A). This leads to a 20-times
reduction in the relative removal of the activation loop (Figure 3-16B), indicating that
O-GlcNAcylation at these sites would inhibit the self-activation of caspase-8 during cell
death. We next wanted to test if O-GlcNAcylation at these sites could affect caspase-8
cleavage in living cells. During apoptosis in cells both dimerization and cleavage have
been shown to be required for the generation of the fully active caspase-8 enzyme.(Oberst
et al., 2010) To determine the consequences of O-GlcNAcylation, we planned to test this
cleavage using both a mutant that cannot be O-GlcNAcylated (i.e., serine to alanine) and
a mutant that could approximate the steric bulk of O-GlcNAc (serine to tryptophan). We
chose tryptophan as a surrogate because there is no way to site-specifically incorporate
O-GlcNAc using artificial amino acids. To test if these mutations effected caspase-8
cleavage, we first synthesized the corresponding serine to alanine (StoA) and serine to
tryptophan (StoW) peptides (5-8) and repeated the in vitro cleavage assay described
above. In this experiment, we found the native peptide 1, containing
374
DSE
376
, to be fully
cleaved by caspase-8. This increased cleavage compared to the previous assay could be
due to the addition of DMSO, which was required to solubilize the mutant peptides.
Consistent with previous experiments,{Stennicke:2000vo} we found that substitution of
the serine at position P1’ in this peptide by either alanine (5,
374
DAE
376
) or tryptophan (6,
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374
DWE
376
) inhibited the cleavage of this peptide, although the inhibition by StoW was
more pronounced. To our knowledge the effects of mutations at the P2’ and P3’ positions
on caspase-8-mediated cleavage have never been systematically tested. Native peptide 2,
containing
384
DLSS
387
, again showed notable cleavage, while the alanine peptide (7,
384
DLAA
387
) showed increased cleavage and the tryptophan peptide (8,
384
DLWW
387
)
displayed less. We then quantitated this data to again determine the efficiency of removal
of the activation loop (Figure 3-16C). We found that the alanine mutations had an overall
small inhibitory effect on caspase-8 activation loop cleavage by 1.2-times compared to
the corresponding serine residues, indicating that the effect of the mutation alone would
be to slow caspase-8 activation slightly. The tryptophan mutations inhibited cleavage of
the loop by 3-times, making these mutations a reasonable, but less-effective, analog of
O-GlcNAcylation for the inhibition of caspase-8 cleavage.
With the peptide mutations characterized in vitro, we set out to test their consequences on
caspase-8 activation in living cells. It has been observed that transient transfection of
caspase-8 can lead to spontaneous dimerization and cleavage/activation of the protein,
presumably due to high cellular concentrations (Figure 3-16D). This feature can be used
to directly test any differences in self-activation caspase-8, without complications from
upstream receptor-mediated activation steps. Unfortunately, the endogenous caspase-8 in
mammalian cells can participate in this process, so we first used used the CRISPR genetic
engineering system to generate several clones of a caspase-8 null HeLa cell-line (Figure
3-14).
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Figure 3-14. Generation of caspase-8-null HeLa cells. Clonal populations of cells that
underwent CRISPR-mediated genetic engineering were analyzed by Western blotting, which
showed no detectable caspase-8 protein. DNA from clone #1 was sequenced and aligned with
wild-type sequence.
With these cells in-hand, we next confirmed that overexpression of caspase-8 would
cause its auto-dimerization. To accomplish this goal, caspase-8 null cells were
co-transfected with wild-type HA- and FLAG-tagged caspase-8 constructs bearing a
active-site mutations to prevent auto-activation. Importantly, we first confirmed that the
antibodies to these tags do not cross-react (Figure 3-15A). Immunoprecipitation with an
anti-FLAG antibody resulted in co-enrichment of the HA-tagged caspase-8 and vice
versa, demonstrating that we were indeed inducing dimerization of caspase-8 upon
overexpression (Figure 3-15B). To determine if global changes in O-GlcNAcylation
could affect this process, we performed the same immunoprecipitation in triplicate in
cells that were treated with either 5SGlcNAc, DMSO, or Thiamet-G to change the overall
O-GlcNAcylation levels (Figure 3-15C). Notably, the changes in O-GlcNAcylation had
no significant effect on the co-immunoprecipitation, indicating it does not alter the ability
of caspase-8 to dimerize during its activation.
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Figure 3-15. Characterization of caspase-8 dimerization. (A) The anti-FLAG and anti-HA
antibodies do not cross-react and can be effectively “stripped.” HeLa cells were separately
transfected with plasmids encoding HA- and FLAG-tagged caspase-8 before analysis by Western
blotting. (B) Caspase-8 auto-dimerizes under overexpression in HeLa cells. Caspase-8 null HeLa
cells were co-transfected with plasmids encoding HA- and FLAG-tagged, inactive caspase-8.
Anti-HA, Anti-FLAG, or anti-mouse IgG were used to immunoprecipitate proteins from these
cells, followed by analysis by Western blotting. The data is representative of two biological
replicates. (C) Changing O-GlcNAcylation levels does not affect the dimerization of caspase-8.
Caspase-8 null HeLa cells were co-transfected with plasmids encoding HA- and FLAG-tagged,
inactive caspase-8, before being treated with 5SGlcNAc (200 μM), DMSO vehicle, or Thiamet-G
(20 μM) to alter O-GlcNAcylation levels. An anti-FLAG immunoprecipitation was then
performed before visualization with an anti-HA antibody, stripping of the blot, and visualization
with an anti-FLAG antibody.
With these control experiments completed, we next tested the effects of mutating the
activation loop from serine to alanine (caspase-8(AAA)) to prevent O-GlcNAcylation or
from serine to tryptophan (caspase-8(WWW)) to mimic O-GlcNAc. Accordingly,
caspase-8 null cells were transiently transfected with wild-type caspase-8,
caspase-8(AAA), or caspase-8(WWW). In contrast to previous experiments, these
caspase-8 enzymes do not contain active site mutations. Cells were collected after
different lengths of time post-transfection, and analysis by Western blotting showed that
at relatively short times (< 36 h) this results in a “pulse” of cleaved/activated caspase-8
that is then removed, potentially by death of the transfected cell population (Figure
3-16F). Using these conditions, side-by-side comparison of wild-type caspase-8,
caspase-8(AAA), and caspase-8(WWW) by Western blotting showed that the
caspase-8(AAA) mutant self-cleaved/activated with faster kinetics than wild-type protein
197
(Figure 3-16F). Specifically, cleavage of caspase-8(AAA) could be detected in as little as
4 h post-transfection, essentially as soon as significant amounts of protein were
detectable. In contrast, wild-type caspase-8 also self-activated but did not reach peak
activation until 12 h post-transfection. This cellular result is notable because our in vitro
peptide data suggested that caspase(AAA) should self-cleave/activate with similar, if not
slightly slower, kinetics compared to wild-type protein. Finally, caspase-8(WWW)
self-activated with the slowest kinetics with a peak at 16 h post-transfection, consistent
with our peptide data and tryptophan acting as an O-GlcNAc surrogate. These data
indicate that O-GlcNAcylation of caspase-8 can change the kinetics of its activation
living cells by serving as a steric hinderance to caspase-8 self-cleavage. However, we
cannot completely rule out some differences in the cellular experiment, such as the
activation of executioner caspases that then feed-back to activate our overexpressed
caspase-8. Importantly, based on our peptide data, bona fide O-GlcNAcylation should
have an even more pronounced inhibitory effect on cleavage than mutation to tryptophan.
198
Figure 3-16. O-GlcNAcylation of the caspase-8 cleavage sites affects its cleavage/activation.
(A) O-GlcNAcylation inhibits the self-cleavage of caspase-8 in vitro. The indicated unmodified or
O-GlcNAcylated peptides (2 mM) were incubated with recombinant, active caspase-8 for 24 h
and the percent cleavage was determined using reverse-phase high-performance liquid
chromatography (RP-HPLC). Results are the mean ±s.e.m of three separate biological
experiments (see Figure S14 (SI)). (B) Comparison of the relative efficiency of caspase-8
activation loop cleavage between unmodified and O-GlcNAcylated peptides. The individual
effects of O-GlcNAcylation shown in (A) were averaged to obtain the overall loop cleavage
efficiency. (C) Mutation of the O-GlcNAcylation sites in caspase-8

to alanine or tryptophan
inhibits overall caspase-8 activation loop cleavage in vitro, with tryptophan having a larger
inhibitory effect. The individual effects of the alanine and tryptophan mutations (shown in Figure
S17 (SI)) were averaged to obtain overall loop cleavage efficiency. (D) Activation of caspase-8 by
transient overexpression. Upon overexpression, caspase-8 will self-dimerize and activate itself
without a requirement for cell-death signaling. (E) Changing O-GlcNAcylation levels does not
affect the dimerization of caspase-8. Caspase-8 null HeLa cells were co-transfected with plasmids
encoding HA- and FLAG-tagged, inactive caspase-8, before being treated with 5SGlcNAc (200
μM), DMSO vehicle, or Thiamet-G (20 μM) to alter O-GlcNAcylation levels. An anti-FLAG
immunoprecipitation was then performed before visualization with an anti-HA antibody, stripping
of the blot, and visualization with an anti-FLAG antibody. The amount of enriched HA-tagged
protein relative to enriched FLAG-tagged protein was then quantified. Results are the average of
three separate biological experiments that were imaged simultaneously (Figure S19B (SI)). Error
199
bars represent ±s.e.m. of three biological experiments, and statistical significance was calculated
using a two-tailed Student’s t-test. (F) Caspase-8 with its O-GlcNAc sites mutated to alanine
(caspase-8(AAA)) activates faster than wild-type protein, and mutation to tryptophan
(caspase-8(WWW)) slows the rate of cleavage. Caspase-8 null HeLa cells were transiently
transfected with plasmids encoding either wild-type caspase-8, caspase-8(AAA), or
caspase(WWW) for 6 h. After the indicated lengths of time, cells were collected. Analysis by
Western blotting shows auto-activation of all proteins. The different amounts of each caspase
species was quantitated and normalized to the total caspase signal in each lane. The data is
representative of two biological experiments.
Discussion
Despite the observations that increased O-GlcNAcylation inhibits apoptosis and promotes
survival in cell culture and tissues undergoing stress, there have been relatively few
mechanistic explanations for this phenotypic association. Here, we demonstrate that the
proteins that directly carry out apoptosis, the caspases, are themselves O-GlcNAc
modified. In order to make this discovery, we created a new metabolic chemical reporter
(MCR), Ac
36AlkGlcNAc, which we show has significantly improved signal-to-noise
compared to our previous probe Ac
36AzGlcNAc(Chuh et al., 2014) while maintaining its
selectivity for intracellular proteins over cell surface glycoproteins (Figures 2 and 3). This
is most likely due to the improved orientation of the CuAAC reaction that reduces
background labeling but could also be a result of improved metabolic stability of the
alkyne- versus azide-MCR. Notably, treatment of cells with Ac
36AlkGlcNAc allowed for
the visualization of modified proteins in as little as 15 min and a large amount of labeling
in only 1 h, which could enable the specific identification of newly O-GlcNAcylated
proteins in response to cellular stimuli (e.g., signaling or stress). Like Ac
36AzGlcNAc,
our new MCR cannot be metabolized to the UDP-6AlkGlcNAc by the canonical GlcNAc
salvage pathway in mammalian cells, and we believe that it can be converted to
200
6AlkGlcNAc-1-phosphate by the action of the enzyme phosphoacetylglucosamine
mutase (AGM1), which we previously demonstrated can phosphorylate 6AzGlcNAc. It is
also possible that there are uncharacterized small-molecule kinases that can generate
these 1-phosphate sugars. In either case, a wide range of cell-lines are strongly labeled by
Ac
36AlkGlcNAc treatment, indicating that this MCR will be broadly applicable. By
applying Ac
36AlkGlcNAc to MEF cells in an unbiased proteomics experiment, we were
able to identify caspase-3 and -8 as being potentially O-GlcNAcylated. A parallel
proteomics experiment using our non-selective MCR, Ac
4GlcNAlk,(Zaro et al., 2011b)
did not identify these caspases, demonstrating the utility of our new compound in
discovery-based experiments. We then confirmed the MCR-labeling of caspase-3 and -8,
as well as caspase-9, in three different cancer cell types. Notably, caspase-9 was not
identified in our proteomics experiment, suggesting that applying Ac
36AlkGlcNAc in a
quantitative proteomics experiment (e.g., SILAC labeling) may uncover additional
interesting O-GlcNAcylated proteins. Critically, we were able to use chemoenzymatic
labeling and/or WGA enrichment of O-GlcNAcylated proteins to independently confirm
the endogenous O-GlcNAc modification of the caspases (Figures 5 and 6). This type of
independent confirmation is a key first step after MCR-based identification of any
O-GlcNAcylated protein, as treatment with metabolic reporters may alter the normal
modification state of the cell.
We next chose to focus on caspase-8 given its key roles in development, immunology,
and human disease.{Salvesen:2014gv} We first used endogenous co-immunoprecipitation
201
to caspase-8 to demonstrated that OGT and caspase-8 can form a physiologically relevant
complex in cells and that OGT has the opportunity to O-GlcNAc modify caspase-8. By
taking advantage of chemoenzymatic labeling of endogenous O-GlcNAc modifications,
we then showed that the O-GlcNAcylated fraction of caspase-8 is resistant to
cleavage/activation upon induction of apoptosis by TNFα. It is possible that
O-GlcNAcylation affects the ability of antibodies to recognize caspase-8; however, we
observe a difference in the amount of cleaved caspase compared to full length and not an
overall reduction in signal in the Western blot, so we think this is unlikely. We also
demonstrated that changing the levels of O-GlcNAcylation can affect the
activation/cleavage of caspase-8 during the initiation of apoptotic signaling by TNFα.
Using site-directed mutagenesis and chemoenzymatic detection, we found that the vast
majority of O-GlcNAcylation on caspase-8 occurs at serine residues near the two
cleavage sites required for full caspase-8 activation during apoptosis. We also
demonstrated that O-GlcNAcylation of these sites inhibits their cleavage in vitro, which
has been shown to be required for the full activation of caspase-8 during
apoptosis,(Oberst et al., 2010) and we used caspase-8 mutants to indicate that
O-GlcNAcylation of these sites can change the kinetics of caspase-8 cleavage/activation
in living cells. We believe that this is certainly not the only O-GlcNAc modification that
causes the phenotypic association between elevated O-GlcNAcylation levels and cell
survival, but our results suggest that caspase-8 O-GlcNAcylation could contribute to the
anti-apoptotic function of O-GlcNAc in cells. The exact stoichiometry of the
modification and the specific biological contexts where caspase-8 O-GlcNAcylation has a
202
significant effect on overall apoptosis remain to be discovered, a line of investigation we
are currently undertaking. Unfortunately, we have found that the generation of cell-lines
that stably express caspase-8 mutants, like caspase-8(AAA), is difficult due to variable
expression levels that result in auto-activation of apoptosis. Additionally, transient
transfection of caspase-8 can be effected by the presence of OGT and OGA inhibitors in
the media. To overcome these issues, we are currently exploring the use of CRISPR-Cas9
to introduce mutants into the caspase-8 gene at the chromosome level. Intriguingly, all of
the apoptotic caspases in humans and mice, two out of the three caspases in Drosophila,
and ced-3 in C. elegans have serine or threonine residues at or near their cleavage sites.
However, the O-GlcNAcylation sites we identified here on caspase-8 are only conserved
in mammals. Additionally, analysis of all of the known caspase cleavage-sites in substrate
proteins{Fridman:2013jt} shows that serine or threonine is the second most favored
amino-acid at all of the surrounding eight residues. Many known caspase substrates have
also been identified as being O-GlcNAcylated in a variety of proteomics experiments,
although the specific sites of modification are largely unknown. Additionally,
phosphorylation of serine/threonine residues near caspase cleavage sites has been shown
to alter proteolysis rates.{Tozser:2003gw, DegliEsposti:2003hg, Dix:2012ee} This raises
the possibilities that O-GlcNAcylation of caspases is an evolutionarily conserved
mechanism for modulating caspase activation and that this modification may similarly
change substrate proteolysis. We are currently exploring both of these avenues of
research. In conclusion, our new MCR (Ac
36AlkGlcNAc) enabled the discovery of
caspase O-GlcNAcylation, and our data show that this modification can inhibit caspase-8
203
cleavage/activation, which may play a role in the regulation of programmed cell death in
development and disease.
Materials and Methods
Chemical Synthesis.
Known chemical reporters Ac
4GlcNAz(Saxon et al., 2002) and Ac 36AzGlcNAc (Saxon et
al., 2002), were synthesized according to literature procedures. The fluorescent detection
tag alk-rho(Charron et al., 2009) and the OGA inhibitor Thiamet-G(Yuzwa et al., 2008)
were also synthesized in lab according to literature procedures. Ac
36AzGlcNAc and
alkyne-azo-biotin were synthesized according to literature procedures as described below.
Dimethyl-(1-diazo-2-oxopropyl)-phosphonate (Bessman reagent): Sodium hydride (0.072
g, 3.01 mmol) was dissolved in benzene (12.8 mL) and tetrahydrofuran (2.2 mL) under a
nitrogen atmosphere and cooled to 0 °C. Dimethyl acetyl-methyl phosphonate (0.500 g,
3.01 mmol) was added dropwise and reaction let stir 90 min upon which time p-toluene
sulfonyl azide (11-14% in toluene, 4.7 mL, 3.01 mmol) was added and reaction was
allowed to warm to room temperature over 1 h. Upon completion, the mixture was
filtered to remove solids and concentrated by vacuum. The resulting crude oil was
purified by column chromatography (70% ethyl acetate in hexanes) to afford the product
as a slightly yellow oil (0.548 g, 95%).
1
H NMR (500 MHz, CDCl 3): δ (ppm) 3.82 (d, J =
12.0 Hz, 6H), 2.24 (s, 4H).
204
Compound 3.1 2-Acetamido-2-deoxy-3,4,5,6-di-O-isopropylidine-aldehydo-D-glucose
dimethyl acetal (Gloster et al., 2011): Commercially available 2-
deoxy-2-N-acetyl-glucopyranose (10.0 g, 45.0 mmol) was suspended
in 1,4 dioxane (100 mL) at 80 °C under nitrogen.
2,2-dimethoxypropane (40 mL) and p-toluene sulfonic acid
monohydrate (1.51 g, 7.87 mmol) were added and the reaction was stirred for 24 h. The
reaction was cooled and neutralized with sodium bicarbonate. After filtration the mixture
was concentrated by vacuum and purified by column chromatography (2%
methanol:CH
2Cl 2) to afford the product as a light brown thick syrup (9.00 g, 63%).
1
H
NMR (400 MHz, CDCl
3): δ (ppm) 5.84 (d, J = 9.9 Hz, 1H), 4.48-4.44 (m, 1H), 4.36 (d, J
= 5.9 Hz, 1H), 4.24 (dd, J = 8.1, 1.4 Hz, 1H), 4.18-4.00 (m, 2H), 3.96 (dd, J = 8.3, 4.6
Hz, 1H), 3.62 (t, J = 7.8 Hz, 1H), 3.41 (s, 3H), 3.37 (s, 3H), 2.02 (s, 3H), 1.47 (s, 3H),
1.39 (s, 3H), 1.37 (s, 3H), 1.33 (s, 3H).
Compound 3.2 2-Acetamido-2-deoxy-3,4-O-isopropylidine-aldehydo-D-glucose dimethyl
acetal (Gloster et al., 2011): Compound 3.1 (8.37 g, 24.1 mmol) was
dissolved in 80% acetic acid in water and was let stir for 2 h at 40 °C.
Upon completion, the reaction was concentrated by vacuum to remove
acid. The resulting mixture was coevaporated with toluene to remove
any residual acid. The resulting oil was purified by column chromatography using a
gradient starting at 40% acetone and increasing to 65% acetone in hexanes to afford the
product as a yellow syrup (6.03 g, 81%).
1
H NMR (400 MHz, CDCl 3): δ (ppm) 6.08 (d, J
MeO OMe
NHAc
O
O
O
O
MeO OMe
NHAc
O
O
OH
OH
205
= 9.5 Hz, 1H), 4.45 (d, J = 6.7 Hz, 1H), 4.38-4.34 (m, 1H), 4.24 (dd, J = 8.4, 1.4 Hz, 1H),
3.78-3.57 (m, 4H), 3.39 (s, 3H), 3.30 (s, 3H), 2.05 (s, 3H), 1.36 (s, 3H) 1.35 (s, 3H).
Compound 3.3 2-Acetamido-6-O-mesyl-2-deoxy-3,4-O-
isopropylidene-aldehydo-D-glucose dimethyl acetal: Compound 3.2
(2.50 g, 8.14 mmol) was dissolved in anhydrous pyridine under a
nitrogen atmosphere and was cooled to -20 °C. Methanesulfonyl
chloride (0.801 mL, 10.3 mmol) was added dropwise. Reaction was stirred for 2 h upon
which time methanol (10 mL) was added and reaction was allowed to warm to rt. The
reaction was concentrated by vacuum and purified by column chromatography (2%
methanol:CH
2Cl 2) to afford the product as a pale yellow oil (2.25 g, 73%).
1
H NMR (400
MHz, CDCl
3): δ (ppm) 5.96 (d, J = 9.0 Hz, 1H), 4.50 (d, J = 6.8 Hz, 1H), 4.43 (dd, J =
10.9, 4.7 Hz, 1H), 4.35-4.31 (m, 1H), 4.28 (dd, J = 10.7, 4.9 Hz, 1H), 4.17 (dd, J = 8.6,
1.7 Hz, 1H), 3.83-3.79 (m, 1H), 3.57 (t, J = 8.6 Hz, 1H), 3.40 (s, 3H), 3.30 (s, 3H), 3.09
(s, 3H), 2.05 (s, 3H), 1.36 (s, 3H), 1.35 (s, 3H).
13
C NMR (125 MHz, CDCl 3): δ (ppm)
171.37, 109.31, 102.33, 78.77, 75.04, 72.07, 71.35, 72.07, 54.87, 52.33, 50.32, 37.49,
26.88, 23.15. ESI/APCI-HRMS calculated for C
14H 27NO 9SNa [M+Na]
+
: 408.1299, found
408.1290.
Compound 3.4 2-Acetamido-6-cyano-2-deoxy-3,4-O-isopropylidene-
aldehydo-D-glucose dimethyl acetal: To a stirred solution of 3.3 (1.95
g, 5.06 mmol) in N,N’-dimethylformamide under a nitrogen
atmosphere, was added sodium cyanide (0.680 g, 13.7 mmol) and
MeO OMe
NHAc
O
O
OH
OMs
MeO OMe
NHAc
O
O
OH
C N
206
tetra-n-butylammonium bromide (0.081 g, 0.253 mmol) at 40 °C. The reaction was let
stir 16 h upon which time it was filtered to remove solids and concentrated by vacuum.
The resulting brown syrup was purified by column chromatography 40% acetone in
hexanes to afford the product as a pale yellow solid (1.38 g, 86%).
1
H NMR (500 MHz,
CDCl
3): δ (ppm) 5.89 (d, J = 8.8 Hz, 1H), 4.46 (d, J = 6.7 Hz, 1H), 4.28-4.24 (m, 1H),
4.08 (dd, J = 8.5, 1.9 Hz, 1H), 3.80-3.75 (m, 1H), 3.45 (t, J = 8.6 Hz, 1H), 3.34 (s, 3H),
3.24 (s, 2H), 2.85 (d, J = 29.3 Hz, 1H), 2.70 (dd J = 16.7, 3.6 Hz, 1H), 2.49 (dd, J = 16.7,
6.6 Hz, 1H), 2.02 (s, 3H), 1.30 (s, 3H), 1.29 (s, 3H).
13
C NMR (125 MHz, CDCl 3): δ
(ppm) 171.53, 117.23, 109.25, 102.16, 78.82, 69.31, 55.03, 52.28, 50.51, 49.51, 45.02,
26.86, 24.88, 23.27. ESI/APCI-HRMS calculated for C
14H 24N 2O 6Na [M+Na]
+
: 339.1527,
found 339.1526.
Compound 3.5 2-Acetamido-6-aldehydo-2-deoxy-3,4-O-
isopropylidene-aldehydo-D-glucose dimethyl acetal: A stirred solution
of 19 (0.269 g, 0.850 mmol) in anhydrous CH
2Cl 2 (4 mL) under a
nitrogen atmosphere was cooled to -78 °C. Diisobutylaluminum
hydride 1M in CH
2Cl 2 (3.74 mL, 3.74 mmol) was added portionwise (1.1 eq every 20
min) at which time 10 mL methanol:1 M HCl (1:1) was added and reaction stirred one
hour at rt. The reaction was diluted with CH
2Cl 2 and poured over dilute sodium
bicarbonate. The mixture was extracted 3X with CH
2Cl 2. The organic layer was washed
with brine and subsequently dried over sodium sulfate, filtered, and concentrated. The
resulting crude mixture was purified by column chromatography (6:3.5:0.5) toluene:ethyl
O O
NHAc
O
O
OH
CHO
207
acetate:water to afford the product as a clear oil (0.135 g, 62%).
1
H NMR (400 MHz,
CDCl
3): δ (ppm) 9.83 (t, J = 1.7 Hz, 1H), 5.90 (d, J = 9.3 Hz, 1H), 4.46 (d, J = 6.7 Hz,
1H), 4.38-4.32 (m, 1H), 4.24-4.17 (m, 2H), 3.53 (dd, J = 8.4, 7.4 Hz, 1H), 3.41 (s, 3H),
3.31 (s, 3H), 2.82 (ddd, J = 17.0, 3.3, 1.4 Hz, 1H), 2.60 (ddd, J = 17.0 Hz, 9.0, 2.0 Hz,
1H), 2.06 (s, 3H), 1.37 (s, 3H), 1.36 (s, 3H).
13
C NMR (125 MHz, CDCl 3): δ (ppm)
201.32, 170.74, 109.17, 102.59, 78.78, 77.92, 68.32, 55.03, 52.49, 50.16, 48.59, 26.97,
23.32.
Compound 3.6 2-Acetamido-6-alkyne-2-deoxy-3,4-O-isopropylidene-
aldehydo-D-glucose dimethyl acetal : To a solution of 3.5 (0.189 g,
0.592 mmol) in anhydrous methanol (3 mL) under nitrogen was added
potassium carbonate ( 0.163 g, 1.18 mmol). Freshly prepared
dimethyl-(1-diazo-2-oxopropyl)-phosphonate (0.136 g, 0.711 mmol) in methanol was
added dropwise and the reaction was stirred for 3 h at rt. Upon completion, the reaction
was diluted with CH
2Cl 2 and poured over dilute sodium bicarbonate. The mixture was
extracted 3X with CH
2Cl2 . The organic layer was washed with brine, dried over sodium
sulfate, filtered and concentrated by vacuum. The resulting crude mixture was purified by
column chromatography by first flushing 1L of 20% ethyl acetate in toluene before
switching to (6:3.5:0.5) toluene:ethyl acetate:methanol to elute the product.
Concentration of desired fractions by vacuum afforded the product as a clear oil (0.141 g,
75%).
1
H NMR (400 MHz, CDCl 3): δ (ppm) 5.86 (d, J = 8.3 Hz, 1H), 4.48-4.41 (m, 2H),
4.42 (dd, J = 8.3, 1.3 Hz, 1H), 3.76-3.70 (m, 1H), 3.59 (t, J = 8.3 Hz, 1H), 3.41 (d, J = 1.7
O O
NHAc
O
O
OH
208
Hz, 3H), 3.33 (s, 3H), 2.65 (ddd, J = 16.9, 3.7, 2.36 Hz, 1H), 2.44 (ddd, J = 16.9, 6.4, 2.7
Hz, 1H), 2.05 (s, 3H), 1.37 (s, 6H).
13
C NMR (150 MHz, CDCl3): δ (ppm) 170.51,
109.00, 102.62, 79.90, 78.29, 77.80, 54.72, 52.54, 50.00, 26.96, 26.92, 25.01, 23.25.
ESI/APCI-HRMS calculated for C
15H 25NO 6Na [M+Na]
+
: 338.1574, found 338.1573.
Compound 3.7 1,3,4-Tri-O-acetyl-2-deoxy-2-N-acetyl-6-deoxy-alkynyl-glucopyranose
(Ac 36AlkGlcNAc): Compound 3.6 (0.141 g, 0.446 mmol) was
dissolved in 2.5 M HCl and heated to reflux for 2 h, upon which
time reaction was cooled to rt and acid was removed by vacuum.
The resulting crude mixture was co-evaporated with toluene to remove residual acid and
then subsequently dissolved in pyridine (5 mL). To the stirring mixture was added acetic
anhydride (0.157 mL, 1.67 mmol) and catalytic 4-dimethylaminopyridine (10 mol %).
The reaction was stirred 16 h at which time the mixture was concentrated by vacuum and
purified by column chromatography (40% acetone in hexanes) to yield the product as a
clear oil (0.091 g, 58%).
1
H NMR (400 MHz, CDCl 3) α-anomer: δ (ppm) 6.16 (d, J = 3.7
Hz, 1H), 5.52 (d, J = 9.2 Hz, 1H), 5.26-5.08 (m, 2H), 4.50-4.44 (m, 1H), 3.94-3.89 (m,
1H), 2.51-2.47 (m, 2H), 2.19 (s, 3H), 2.06 (s, 3H), 2.05 (s, 3H), 1.93 (s, 3H).
13
C NMR
(125 MHz, CDCl
3): δ (ppm) 171.72, 169.90, 169.17, 168.63, 90.52, 78.33, 75.70, 72.59,
70.90, 70.64, 69.59, 51.07, 29.66, 23.02, 21.91, 20.70. ESI/APCI-HRMS calculated for
C
16H 21NO 8Na [M+Na]
+
:378.1159, found 378.1554.
O
AcO
AcO
OAc
NHAc
209
Synthesis of known small molecules. Known compounds, alkyne- and azide-rhodamine
(alk-rho and az-rho),(Charron et al., 2009) UDP-GalNAz,(Hang et al., 2003)
Thiamet-G,(Yuzwa et al., 2008) Ac
45SGlcNAc,(Gloster et al., 2011) Ac 4GlcNAlk,(Zaro
et al., 2011b) Ac
4ManNAlk,(Hsu et al., 2007) and Ac 36AzGlcNAc(Chuh et al., 2014)
were synthesized according to literature procedures.
Cell culture. Cos-7, HEK293, and HeLa cells were cultured in high glucose DMEM
media (CellGro) enriched with 10% fetal bovine serum (FBS, Atlanta Biologicals).
MCF-7 cells were cultured in high glucose DMEM media (CellGro) enriched with 10%
fetal bovine serum (FBS, Atlanta Biologicals) and 1% insulin (Thermo Fischer Scientific,
#41400-045). H1299 cells were cultured in RPMI 1640 (CellGro) medium enriched with
10% FBS. NIH3T3 and MEF cells were cultured in DMEM media (CellGrow) enriched
with 10% fetal calf serum (FCS, Thermo). SH-SY5Y cells were cultured in 50:50
DMEM:F12K enriched with 10% FBS (Atlanta Biologicals). All cell lines were
maintained in a humidified incubator at 37 ℃ and 5.0% CO
2. All cells were purchased
from ATCC. All cells tested negative for mycoplasma infection before being used in
experiments. Although, HEK cells are on the ICLAC database of misidentified cell-lines,
they are a commonly used mammalian cell-line for biological experiments and therefore
were included in our MCR labeling panel.
Western blotting. Proteins were separated by SDS-PAGE before being transferred to
PVDF membrane (Bio-Rad) using standard Western blotting procedures unless otherwise
210
indicated. All Western blots besides anti-O-GlcNAc and caspase blots (see below) were
blocked in TBST (0.05% Tween-20, 150 mM NaCl, 10mM Tris pH 8.0) containing 5%
non-fat milk for 1 h at rt. They were then incubated with the appropriate primary
antibody in blocking buffer overnight at 4 ℃. Anti-O-GlcNAc blots were blocked in
TBST containing 5% bovine serum albumin (BSA) for 1 h at rt and incubated overnight
at 4 ℃. Caspase blots were blocked in TBST (0.1% Tween-20, 150 mM NaCl, 20 mM
Tris, pH 8.0) containing 5% nonfat milk overnight at 4 ℃, then incubated with the
appropriate primary antibody in blocking buffer for 24 h at 4 ℃. The anti-actin (Sigma,
#A2066), anti-β-Actin (Sigma, #A5441), anti-O-GlcNAc (RL2; Sigma, #MA1-072),
anti-caspase-8 (Cell Signaling 1C12, #9746; AdipoGen C15, #AG-20B-0057),
anti-caspase-9 (Cell Signaling, #9802), anti-caspase-3 (Cell Signaling, #9662),
anti-cleaved-caspase-3 (Cell Signaling, #9664), anti-OGT (Cell Signaling #24083),
anti-HA tag (C29F4) (Cell Signaling, #3724), anti-DYKDDDDK (Cell Signaling,
#2386), anti-Nup62 (MAb414; Covance, #MMS-120R), anti-pyruvate kinase (Abcam,
#ab6191) were used at 1:1,000 dilution. The blots were then washed three times in the
appropriate TBST for 10 min and incubated with the horseradish peroxidase
(HRP)-conjugated secondary antibody for 1 h in the appropriate blocking buffer at RT.
HRP-conjugated anti-mouse (Jackson ImmunoResearch, #715-035-150) and anti-rabbit
(Jackson ImmunoResearch, #711-035-152) were used at 1:10,000 dilutions. After being
washed three more times with TBST for 10 min, the blots were developed using ECL
reagents (Bio-Rad) and the ChemiDoc XRS+ molecular imager (BioRad). Western blots
were stripped when indicated with Thermo Scientific Restore Western Blot Buffer
211
(#21059). Specifically, following visualization, blots were washed 3X for 10 min each
with 1X PBS before being stripped for 30 mins and washed again 3X 10 mins each with
1X PBS. Blots were then imaged again using ECL as indicated above to ensure
successful stripping before being re-blocked. Streptavidin-HRP blots were blocked in
TBST (0.05% Tween-20, 150 mM NaCl, 10mM Tris pH 8.0) containing 5% bovine
serum albumin (BSA) for 1 h at rt before being washed 3X for 10 min with TBST. Blots
were then incubated with streptavidin-HRP at 1:10,000 (Jackson ImmunoResearch,
#016-030-084) in TBST overnight at 4 ℃. Blots were again washed 3X for ten min
before being developed using ECL reagents and imaged as described above.
Metabolic chemical reporter labeling and in-gel fluorescence. To cells at 80-85%
confluency, media containing Ac
4GlcNAc (control) or metabolic chemical reporter
(1,000X stock in DMSO), was added as indicated. The cells were then collected by
trypsinolysis and pelleted by centrifugation (4 min, 2,000 x g, 4 ℃), followed by
washing 2X with 1X PBS (1 mL). Cell pellets were then resuspended in 100 μL of 1%
NP-40 lysis buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4]
with Complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Roche) and
incubated on ice for 20 min. Cell debris was pelleted by centrifugation (10 min, 10,000 x
g, 4 ℃). The supernatant (soluble cell lysate) was collected and the protein concentration
was determined by BCA assay (Pierce, ThermoScientific). To each sample (diluted to 1
mg mL
-1
final protein concentration), newly made CuAAC cocktail was added [azido- or
alkyne-rhodamine tag (100 μM, 10 mM stock solution in DMSO);
212
tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (1 mM, 50 mM freshly prepared
stock solution in water); tris[(1-benzyl-1-H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (100
μM, 10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared
stock solution in water)]. The reaction was gently vortexed and allowed to sit at room
temperature for 1 h in the dark. Upon completion, 4X volume of ice cold methanol was
added to the reaction, and proteins were precipitated at -20 ℃ for a minimum of 2 h. The
reactions were then centrifuged (10 min, 10,000 x g, 4 ℃) to collect proteins. The
supernatant was removed, the pellet was allowed to air dry for 5-10 min, and then
dissolved in 4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) with
sonication in a bath sonicator to ensure complete dissolution. Equal volume of 2X
loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4% β-mercaptoethanol) was
then added and the samples were boiled for 5 min at 98 ℃ before being analyzed by
SDS-PAGE separation (4-20% Tris-Glycine Gel, Bio-Rad). Following SDS-PAGE
separation, gels were scanned on a Typhoon 9400 Variable Mode Imager (GE Healthcare)
using a 532 nm for excitation and 30 nm bandpass filter centered at 610 nm for detection.
FoxO1 labeling. NIH3T3 cells stably expressing FLAG-tagged FoxO1 were treated with
200 μM Ac
4GlcNAlk, Ac 36AlkGlcNAc (1,000x stock in DMSO) or Ac 4GlcNAc (as a
control) and allowed to incubate overnight. After 16 h, cells were washed with PBS,
trypsinized and pelleted. Cell pellets were resuspended in 100 μL of 1% NP-40 lysis
buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with Complete
Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Thermo Scientific) for 20 min and
213
then centrifuged at 4 ℃ for 10 min at 10,000 x g. The supernatant was collected and the
protein concentration was determined by BCA assay (Pierce, ThermoScientific). Total
cell lysate (1.5 mg) was diluted as necessary to a final volume of 1 mL with 1% NP-40
buffer with Complete Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Thermo
Scientific). EZview Red ANTI-FLAG M2 affinity beads (30 μL, Sigma), pre-washed with
cold NP-40 buffer 2X followed by cold PBS 2X, were added to each sample. The
samples were placed on a rotator for 2 h at 4 ℃. Beads were collected by centrifugation
at 2,000 x g for 2 min at 4 ℃, and the supernatant was carefully removed. Beads were
then washed with cold PBS by rotating for 5 mins before centrifuging 2 mins at 2,000 x
g. The final PBS wash was carefully removed, and the beads were suspended in 30 μL
4% SDS buffer and boiled for 5 min at 97 ℃. The appropriate amount of click chemistry
cocktail was added, and the reaction was allowed to proceed for 1 h after which time 30
μL of 2x loading buffer was added. Samples were boiled for 5 minutes at 97 ℃. Protein
samples (40 μg) were then loaded per lane for SDS-PAGE separation (Any Kd Criterion
Gel, Bio-Rad) and imaged by in-gel fluorescence scanning.
GlyCAM-IgG labeling. NIH3T3 cells stably expressing GlyCAM-IgG in 6-well dishes at
80-85% confluency were treated in DMEM with 10% FCS and 200 μM Ac
4GlcNAlk,
Ac
36AlkGlcNAc (1,000x stock in DMSO) or Ac 4GlcNAc (as a control) for 24 hours. The
media from each sample was collected by centrifugation at 3,000 x g for 10 min at 4 ˚C
to remove cell debris. The supernatant (1mL) was incubated with 50 μL of recombinant
protein G sepharose beads (Invitrogen) in 100 mM TEA pH 8 overnight. Beads were
214
collected by centrifugation at 2,000 x g for 2 min at 4 ℃. Beads were washed 3X with 1
mL 100 mM TEA pH 8. GlyCAM-Ig was eluted by addition of 50 μL 4% SDS buffer
(4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) and boiling for 5 min at 97 ˚C. Protein
concentration was determined by BCA assay (ThermoScientific). Final SDS
concentration was diluted to 0.5% by addition of 50 mM TEA pH 7.4. The appropriate
amount of click chemistry cocktail was added and the reaction was allowed to proceed
for 1 h after which time 4X loading buffer (200 mM Tris HCl, 4% SDS, 40% glycerol,
0.4% bromophenol blue, 1.4% β-mercaptoethanol, pH 6.8) was added. Samples were
boiled for 5 min at 97 ˚C and 50 μg were loaded for SDS-PAGE separation (Any Kd
Criterion Gel, Bio-Rad).
Flow cytometry of cell-surface labeling with picolyl azide. NIH3T3 cells grown in 100
mm plates at 80-85% confluency were treated with 200 μM Ac
4GlcNAc, Ac 4GlcNAlk,
Ac
4ManNAlk or Ac 36AlkGlcNAc in triplicate for 16 hours at which time media was
removed and cells were gently washed with PBS before being detached from the plate
with cold 1 mM EDTA in PBS. Cells were collected by centrifugation (5 min, 500 x g at
4 ℃) and were washed three times with PBS (5 min, 500 x g at 4 ℃). Cells were then
resuspended in 600 μL PBS. To each sample, newly made CuAAC cocktail was added
[picolyl azide (50 μM, 10 mM stock solution in DMSO, Click Chemistry Tools);
tris(3-hydroxypropyltriazolylmethyl)amine (THPTA, Click Chemistry Tools) (200 μM ,
10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared stock
solution in water, Sigma Aldrich); and sodium ascorbate (2.5 mM, 100 mM freshly
215
prepared stock solution in water, Alfa Aesar)]. The cells were gently vortexed and
allowed to react for 5 mins at rt at which time cells were pelleted by centrifugation and
washed 3X with PBS (5 min, 500 x g at 4 ℃). Cells were then resuspended in 600 μL of
PBS containing fluorescein isothiocynate (FITC) conjugated avidin (Sigma, 5 μg/mL)
and incubated on ice for 1 h in the dark. Cells were then pelleted and washed three more
times (5 min, 500 x g at 4 ℃) before being resuspended in 1 mL of PBS. A total of
10,000 cells [dead cells were excluded by treatment with propdium iodide (2.5 μg/mL in
water, 15 mins)] were analyzed on a MACSQuant Analyzer
®
10 using the 488 nm argon
laser.
Cell Proliferation Assay. NIH3T3 or HeLa cells were plated 24 h prior to treatment at
5,000 cells/well in a 96-well plate (Costar, black, clear bottom). Cells were then treated
with either DMSO, or 200 μM Ac
36AzGlcNAc, Ac 36AlkGlcNAc or Ac 4GlcNAlk
(1,000X stocks in DMSO) for 16 h. Before addition of assay, media was exchanged for
fresh media. Twenty μL of reagent was added per well and let incubate for 1 h at 37 ℃,
according to manufacturer’s protocol (Celltiter 96 Aqueous, Non-Radioactive
Cell-Proliferation Assay (MTS), Promega). The plate was then read at 490 nm after
shaking with medium intensity for 5 sec using a Biotek Synergy H4 Hybrid Multi-Mode
Microplate reader.
Apoptosis (Caspase-3/-7) Assay. NIH3T3 or HeLa cells were plated 24 h prior to
treatment at 5,000 cells/well in a 96-well plate (Costar, white, clear bottom). Cells were
216
then treated with either DMSO, or 200 μM Ac36AzGlcNAc, Ac 36AlkGlcNAc or
Ac
4GlcNAlk (1,000X stocks in DMSO) for 16 h at which time the plate was allowed to
equilibrate to rt. To each well containing 100 μL of media was added 100 μL of reagent
according to manufacturer’s protocol (Caspase-Glo 3/7 Assay, Promega) and let incubate
for 30 min at rt. Luminescence was then measured following 2 min of high intensity
shaking using a Biotek Synergy H4 Hybrid Multi-Mode Microplate reader (Biotek, Gain
= 135, integration time = 5 sec, read height of 1 mm).
Metabolic chemical reporter labeling and proteomics. Immortalized mouse embryonic
fibroblasts (MEFs) were treated for 16 h with 200 μM concentrations of
Ac
36AlkGlcNAc, Ac4GlcNAlk, or Ac4GlcNAc (as a control). Cells were then collected
by trypsinolysis, followed by centrifugation at (4 min, 2,000 x g, 4 ℃), followed by
washing 2X with 1X PBS (1 mL). Cell pellets were resuspended in 200 μL H
2O, 60 μL
PMSF in H
2O (250 mM), and 500 μL 0.05% SDS buffer (0.05% SDS, 10 mM TEA pH
7.4, 150 mM NaCl) with Complete Mini protease inhibitor cocktail (Roche Biosciences).
To this was added 8 μL Benzonase (Sigma), and the cells were incubated on ice for 30
min. Then, 4% SDS buffer (2,000 μL) was added, and the cells were briefly sonicated in
a bath sonicator followed by centrifugation (10,000 x g for 10 min at 15 °C). Soluble
protein concentration was normalized by BCA assay (Pierce, ThermoScientific) to 1 mg
mL
−1
, and 10 mg of total protein was subjected to the appropriate amount of click
chemistry cocktail containing alkyne-PEG
3-biotin (5 mM, Click Chemistry Tools) for 1 h,
after which time 10 volumes of ice-cold MeOH were added. Precipitation proceeded 2
217
hours at -20 °C. Precipitated proteins were collected by centrifugation (30 min, 5,000 x g,
4 °C) and washed 3 times with 40 mL ice-cold MeOH, with resuspension of the pellet
each time. The pellet was then air-dried for 15 min. To capture the biotinylated proteins
by streptavidin beads, the air-dried protein pellet was resuspended in 2 mL of
resuspension buffer (6 M urea, 2 M thiourea, 10 mM HEPES pH 8.0) by bath sonication.
To cap cysteine residues, 100 μl of freshly-made TCEP (200 mM stock solution, Thermo)
was then added and the mixture incubated for 30 min, followed by 40 μl of freshly
prepared iodoacetamide (1 M stock solution, Sigma) and incubation for a further 30 min
in the dark. Steptavadin beads (250 μL of a 50% slurry per sample, Thermo) were washed
2X with 1 mL PBS and 1X with 1 mL resuspension buffer and resuspended in
resuspension buffer (200 μL). Each sample was combined with streptavadin beads and
incubated on a rotator for 2 h. These mixtures were then transferred to Mini Bio-Spin
®

columns (Bio-Rad) and placed on a vacuum manifold. Captured proteins were then
washed with agitation 5X with resuspension buffer (10 mL), 5X PBS (10 mL), 5X with
1% SDS in PBS (10 mL), 30X with PBS (1 mL per wash, vacuum applied between each
wash), and 5X 2M urea in PBS (1 mL per wash, vacuum applied between each wash).
Beads were then resuspended in 2 M urea in PBS (1 mL), transferred to screw-top tubes,
and pelleted by centrifugation (2000 x g for 2 min). At this time, 800 μL of the
supernatant was removed, leaving a volume of 200 μL. To this bead-mixture was added 2
μL of CaCl
2 (200 mM stock, 1 mM final concentration) and 2 μL of 1 mg mL
−1
sequence
grade trypsin (Promega) and incubated at 37 °C for 18 hours. The resulting mixtures of
tryptic peptides and beads were transferred to Mini Bio-Spin
®
columns (Bio-Rad) and the
218
eluent was collected by centrifugation (1,000 x g for 2 min). Any remaining peptides
were eluted by addition of 100 μL of 2 M urea in PBS followed by centrifugation as
immediately above. The tryptic peptides were then applied to C18 spin columns (Pierce)
according to manufacturer's instructions, eluted with 70% acetonitrile in H
2O, and
concentrated to dryness on a speedvac. Peptides were then desalted on a trap column
following separation on a 12cm/75um reversed phase C18 column (Nikkyo Technos Co.,
Ltd. Japan). A 3 hour gradient increasing from 10% B to% 45% B in 3 hours (A: 0.1%
Formic Acid, B: Acetonitrile/0.1% Formic Acid) was delivered at 150 nL min
−1
. The
liquid chromatography setup (Dionex, Boston, MA, USA) was connected to an Orbitrap
XL (Thermo, San Jose, CA, USA) operated in top-5-mode. Acquired tandem MS spectra
(CID) were extracted using ProteomeDiscoverer v. 1.3 (Thermo, Bremen, Germany) and
queried against the human Uniprot protein database using MASCOT 2.3.02
(Matrixscience, London, UK). Peptides fulfilling a Percolator calculated 1% false
discovery rate threshold were reported. All LC-MS/MS analysis were carried out at the
Proteomics Resource Center at The Rockefeller University, New York, NY , USA.
FLAG- and WGA-enrichment of Caspases. NIH3T3 cells stably expressing either
FLAG-tagged Caspase-3 or -8 were grown to 80% confluency before being treated with
20 μM Thiamet-G for 16 hours. Cells were then harvested by trypsinization and washed
2X with PBS (2 min, 2,000 x g, 4 ℃ ). Cell pellets were resuspended in 100 μl of 1%
NP-40 lysis buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4]
with Complete Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Thermo Scientific)
219
for 20 min and then centrifuged at 4 ℃ for 10 min at 10,000 x g. The supernatant was
collected and the protein concentration was determined by BCA assay (Pierce,
ThermoScientific). Total cell lysate (4 mg) was diluted to a final concentration of 1
mg/mL with 1% NP-40 buffer with Complete Mini, EDTA-free Protease Inhibitor
Cocktail Tablets (Thermo Scientific). Diluted lysate was then pre-cleared with Protein-G
beads (30 μL, Pierce) pre-washed 3X with 1 mL of NP-40 buffer, for 1h at 4 ℃ with full
rotation. Beads were then pelleted by centrifugation (2 min, 2,000 x g, 4 ℃) and cleared
supernatant was transferred to tubes containing EZview Red ANTI-FLAG M2 affinity
beads (30 μL, Sigma), pre-washed with cold NP-40 buffer 2X followed by cold PBS 2X.
The samples were placed on a rotator for 2 h at 4 ℃. Beads were collected by
centrifugation at 2,000 x g for 2 min at 4 ℃, and the supernatant was carefully removed.
Beads were subsequently washed 3X by addition of 1 mL cold PBS with rotation for 5
mins before centrifuging 2 mins at 2,000 x g. Protein was eluted by addition of 30 μL of
1% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA, pH 7.4) and boiling 5 min at 98
℃. Supernatant was let cool to room temperature before being transferred to a new tube
in which SDS concentration was diluted to 0.5% by addition of 1% NP-40 buffer lysis
buffer. Resulting diluted FLAG enriched lysate was then transferred to new tubes
containing pre-washed (PBS 1X, cold NP-40 2X) sWGA agarose resin (60 μL, Vector
Labs) and was incubated with rotation for 16 h at 4 ℃. Following enrichment, resin was
washed 3X with PBS before proteins were eluted with 4% SDS buffer and boiling for 5
min at 98 ℃. Samples were then analyzed by SDS-PAGE (Any Kd, Criterion Gel,
Bio-Rad) followed by analysis by Western blotting.
220
FLAG-IP of Caspases. NIH3T3 cells stably expressing either FLAG-tagged Caspase-3
or -8 were grown to 80% confluency before being treated with 200 μM Ac
4GlcNAc or
Ac
36AlkGlcNAc for 16 hours. Cells were then harvested by trypsinization and washed
2X with PBS (2 min, 2,000 x g, 4 ℃ ). Cell pellets were resuspended in 100 μl of 1%
NP-40 lysis buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4]
with Complete Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Thermo Scientific)
for 20 min and then centrifuged at 4 ℃ for 10 min at 10,000 x g. The supernatant was
collected and the protein concentration was determined by BCA assay (Pierce,
ThermoScientific). Total cell lysate (2 mg) was diluted to a final concentration of 0.5
mg/mL with 1% NP-40 buffer with Complete Mini, EDTA-free Protease Inhibitor
Cocktail Tablets (Thermo Scientific). Diluted lysate was then pre-cleared with Protein-G
beads (30 μL, Pierce) pre-washed 3X with 1 mL of NP-40 buffer, for 1h at 4 ℃ with full
rotation. Beads were then pelleted by centrifugation (2 min, 2,000 x g, 4 ℃) and cleared
supernatant was transferred to tubes containing EZview Red ANTI-FLAG M2 affinity
beads (30 μL, Sigma), pre-washed with cold NP-40 buffer 2X followed by cold PBS 2X.
The samples were placed on a rotator for 2 h at 4 ℃. Beads were collected by
centrifugation at 2,000 x g for 2 min at 4 ℃, and the supernatant was carefully removed.
Beads were subsequently washed 3X by addition of 1 mL cold PBS with rotation for 5
mins before centrifuging 2 mins at 2,000 x g. Protein was eluted by addition of 30 μL of
4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA, pH 7.4) and boiling 5 min at 98
℃. Click chemistry with alkyne-biotin was performed in the presence of the
221
FLAG-beads assuming volume of 60 μL. Click chemistry reaction was quenched by
addition of 60 μL of 2X loading buffer and boil 5 min at 98 ℃. Samples were then
analyzed by SDS-PAGE (Any Kd, Criterion Gel, Bio-Rad) followed by analysis by
streptavidin or Western blotting.
Expression and purification of GalT Y289L. pET23a GalT Y289L plasmid was provided
by P. Qasba, National Cancer Institute, and was subsequently transformed into BL21 E.
coli (Novagen). A 1 L culture containing ampicillin (100 μg mL
-1
) was inoculated with 10
mL of 50 mL starter culture grown overnight at 37 ℃. The 1 L culture was grown at 37
℃ until an O.D. (A
600) of 0.60 was obtained, at which time expression was induced with
isopropyl β-D-1-thiogalactopyranoside (ITPG) (1 mM final concentration, 1,000X stock
in water) for 4 h at 37 ℃. Cells were harvested by centrifugation (10 min, 4,000 x g, 4
℃) and resuspended in 10 mL suspension buffer (25% sucrose w/v in 1X PBS). Cells
were lysed using sonication (30 s pulse, 30 s rest, 12 min at 4 ℃). The resulting lysate
was diluted to 80 mL in cold suspension buffer and inclusion bodies were harvested by
centrifugation (30 min, 15,000 x g, 4 ℃) and washed 8-10X or until inclusion bodies turn
white in color, by resuspension in 80 mL suspension buffer followed by centrifugation.
Inclusion bodies were then washed 1X with cold wash buffer (10 mM phosphate, pH 7.0)
and harvested by centrifugation. Inclusion bodies were then resuspended in 14 mL cold
H
2O and poured over solid guanidine HCl and Na 2SO 3 (resulting in 5M GuHCl and 300
mM Na
2SO3 final concentration), vortexed vigorously, and the volume was adjusted to 25
mL with cold H
2O. Freshly made NTSB solution (50 mM DTNB, 1 M Na 2SO 3 in water
222
pH 8.0) was then added with vigorous vortexting to sulfenate free thiols. Completion of
the reaction is indicated by a color change from dark orange to pale yellow. Protein was
then precipitated with cold H
2O (250 mL) and centrifuged immediately (30 min, 9,800 x
g, 4 ℃). Protein pellet was washed 3X by resuspension in cold H
2O and centrifugation.
Protein pellet was finally resuspended in 14 mL cold H
2O and poured over a solid
guanidine HCl and vortexted and then diluted to 25 mL with H
2O (resulting in 5M final
concentration). The protein solution was diluted to a final concentration of 1 mg mL
-1
with a 5M GuHCl solution (O.D. (A 275) ~2.0). This solution was diluted 10-fold into cold
refolding buffer (5 mM EDTA, 4 mM cysteamine, 2 mM cystamine, 100 mM Tris Base,
pH 8.0) with gentle stirring. Protein was allowed to refold for 48 h at 4 ℃ without
agitation. Refolding protein was dialyzed 2X into cold H
2O for 24 h and concentrated
using centrifugal filters (30 KDa cutoff, Amicon Ultra, Millipore) to 3 mL. Buffer was
exchanged 1X with 10 mL reaction buffer (10 mM Tris Base, pH 8.0) and final protein
concentration was determined (O.D. (A
275) ~1.5). Purified protein was stored at 4 ℃.
Chemoenzymatic labeling. Cells were collected by trypsinization and washed 2X with
PBS (2 min, 2,000 x g, 4 ℃ ). Cells were subsequently lysed by adding 13 μL H 2O, 25
μL 0.05% SDS buffer (0.05% SDS, 5 mM MgCl
2, 10 mM TEA, pH 7.4) and 1 μL
Benzonase (Sigma Ultra Pure) and incubated on ice for 30 min. Then, 100 μL of 4% SDS
buffer was added (4% SDS, 150 mM NaCl, 50 mM TEA, pH 7.4), and the resuspended
cell lysate was sonicated briefly in a bath sonicator and cell debris was pelleted (10 min,
10,000 x g, 15 ℃). Protein concentration was determined by BCA Assay (Pierce,
223
ThermoScientific), and the lysate was diluted to 1 μg μL
-1
in 1% SDS chemoenzymatic
buffer (1% SDS, 20 mM HEPES, pH 7.9). Proteins were precipitated by adding 3X
volume of methanol, 0.75X volume chloroform and 2X volume H
2O followed by
vortexing and centrifugation (5 min, 13,000 x g, rt). The aqueous phase was discarded
without disturbing the interface layer before adding 2.5X volume of methanol, vortexing
and pelleting protein (5 min, 13,000 x g, rt). The resulting protein pellet was allowed to
air dry for no longer than 20 mins before being resuspended in 1% SDS chemoenzymatic
buffer (1% SDS, 20 mM HEPES, pH 7.9). Protein concentration was normalized using
BCA Assay (Pierce, ThermoScientific) and diluted to 2.5 μg μL
-1
in 1% SDS
chemoenzymatic buffer. To start the chemoenzymatic transfer reaction, these reagents
were added in the following order (for 100 μg of total protein): 49 μL of H
2O, 80 μL of
labeling buffer (2.5X; 5% NP40, 125 mM NaCl, 50 mM HEPES, pH 7.9), 55 μL of
MnCl
2 (100 mM in H 2O) and 50 μL of UDP-GalNAz (0.5 mM in 10 mM HEPES, pH
7.9). Mix by pipetting. Finally, 7.5 μL of purified GalT Y289L (in 10 mM Tris pH 8.0)
was added and reaction mixture was incubated for 16 h at 4 ℃ without agitation.
Unreacted UDP-GalNAz was then removed by methanol-chloroform precipitation as
described previously. Air dried protein pellets were resuspended in 1% SDS CuAAC
buffer (1% SDS, 150 mM NaCl, 50 mM TEA, pH 7.4) and subjected to CuAAC as
detailed below.
Enrichment and selective elution of O-GlcNAcylated proteins. Cell lysates (~ 1 mg at 1
mg mL
-1
) that had been labeled either chemoenzymatically or with a metabolic chemical
224
reporter were subjected to CuAAC conditions with freshly made cocktail [azide- or
alkyne-azo-biotin (100 μM, 5 mM stock solution in DMSO) (Click Chemistry Tools);
tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (1 mM, 50 mM freshly prepared
stock solution in water); tris[(1-benzyl-1-H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (100
μM, 10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared
stock solution in water)]. After 1 h, the proteins were precipitated by addition of 4
volumes of ice-cold methanol and incubation at -20 °C for 2 h. Precipitated proteins were
collected by centrifugation (30 min, 5,000 x g, 4 °C) and washed 3X with ice-cold
methanol, with resuspension of pellet each time. The pellet was then air dried for 15 min
and then resuspended in 800 μL of resuspension buffer (6 M urea, 2 M thiourea, 10 mM
HEPES pH 8.0) by bath sonication. Samples were then transferred to 2 mL dolphin-nosed
tubes containing streptavidin beads (25 μL of a 50% slurry per sample, Thermo) were
washed 2 X with 1 mL PBS and 1 X with 1 mL resuspension buffer and resuspended in
resuspension buffer (200 μL). Samples were then incubated on a rotator for 2 h. Beads
were washed 2X with resuspension buffer, 2X in PBS (1 ml) and 2X with 1% SDS in
PBS buffer. Beads were then incubated in 25 μL of sodium dithionite solution (1% SDS,
25 mM sodium dithionite) for 30 min at room temperature to elute captured proteins. The
beads were centrifuged for 2 min at 2,000 x g and the eluent collected. The elution step
was repeated, and the eluents combined. Proteins were then precipitated in ice cold
methanol (1 mL) overnight in -20 °C. Protein was collected by centrifugation (10 min,
10,000 x g, 4 °C), and the pellet was allowed to air dry for 5 min, and then 30 μL 4%
SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) was added to each sample.
225
The mixture was sonicated in a bath sonicator to ensure complete dissolution, and 30 μL
of 2x loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4% β-mercaptoethanol)
was then added. Samples were then boiled for 5 mins at 98 °C before being separated by
SDS-PAGE (4-20% Tris-Glycine Gel, Bio-Rad).
Caspase-8/OGT Co-IP . H1299 cells at 90% confluency were washed with ice-cold 1X
PBS before being removed from the plate by scraping. Cells were then pelleted by
centrifugation ( 3 min, 500 x g, 4 ℃ ). Cells were then lysed by resuspension in 500 μL
of lysis buffer (10X Lysis buffer, Cell Signaling) with 1mM phenylmethanesulfonyl
fluoride (PMSF, 200 mM stock in isopropyl alcohol) followed by sonication (5 sec on, 5
sec rest, 3X). Protein concentration was determined using BCA Assay (Pierce,
ThermoScientific), and 200 μg of lysate was diluted to 1 μg μL
-1
in lysis buffer
containing PMSF. Lysate was then pre-cleared with 30 μL of magnetic protein-G beads
(Pierce, #88847) for 1 h at 4 ℃ with end-over-end rotation. Inputs were generated at 2 μg
μL
-1
using 4X loading buffer (8% SDS, 40% glycerol, 0.4% bromophenol blue, 2.8%
B-mercaptoethanol, 200 mM Tris-HCl pH 6.8) before boiling for 5 min at 98 ℃.
Magnetic beads were captured using a magnetic rack and pre-cleared lysate was
transferred to a new, pre-cooled tube. Primary antibody (Caspase-8 (1C12) 1:50 dilution,
Cell Signaling, #9746 or OGT 1:50 dilution, Cell Signaling #24083) was then added to
the pre-cleared lysate. The appropriate isotype control was used at the same protein
concentration as the primary antibody (Mouse isotype control, Cell Signaling #5415 or
Rabbit isotype control, Cell Signaling #3900). Thirty μL of magnetic protein-G beads
226
were then added and let incubate over night with end-over-end rotation at 4 ℃. Protein-G
beads were then washed 5X with 500 μL of lysis buffer containing 1 mM PMSF. Beads
were the pelleted gently be centrifugation (3 min, 2,000 x g, 4 ℃) before 3X loading
buffer containing DTT (Cell Signaling, #7722) was added and samples were boiled for 5
min at 98 ℃. After removal of magnetic beads, samples were analyzed by SDS-PAGE
(4-20% Tris-Glycine Gel, Bio-Rad) and analyzed by Western blotting.
Endogenous caspase-8 chemoenzymatic/Biotin IP following induction of apoptosis.
HeLa cells were plated at 2 x 10
6
cells in a 150 mm dish 48 h prior to activation.
Apoptosis was then induced by treating with 10 ng mL
-1
TNFα and 1 μg mL
-1

cycloheximide (CHX) for 6 h. Cells in the media were combined with cells that were
collected by trypsinization before they were pelleted by centrifugation (6 min, 2,000 x g,
4 ℃) and washed with PBS (1 mL) three times. Cells were subsequently lysed by adding
26 μL H
2O, 50 μL 0.05% SDS buffer (0.05% SDS, 5 mM MgCl 2, 10 mM TEA, pH 7.4)
containing Z-V AD-FMK (Enzo Life Sciences) and 1 μL Benzonase (Sigma Ultra Pure)
and incubated on ice for 30 min. Then, 200 μL of 4% SDS buffer was added (4% SDS,
150 mM NaCl, 50 mM TEA, pH 7.4), and the resuspended cell lysate was sonicated
briefly in a bath sonicator and cell debris was pelleted (10 min, 10,000 x g, 15 ℃).
Precipitated protein was subjected to chemoenzymatic transfer followed by click
chemistry with alkyne-azo-biotin before being subjected to streptavidin enrichment,
SDS-PAGE and Western blotting as previously described.
227
Caspase activation kinetics. HeLa cells were treated at 70-75% confluency with either
100 μM 5SGlcNAc, 5 μM Thiamet-G (1,000X stocks in DMSO), or DMSO vehicle for
16 h. Apoptosis was then induced by treating with 10 ng mL
-1
TNFα and 1 μg mL
-1

cycloheximide (CHX) for either 0, 1, 2, 3, 4, or 6 h. Cells in the media were combined
with cells that were collected by trypsinization. The cells were then pelleted by
centrifugation (6 min, 2,000 x g, 4 ℃) and washed with PBS (1 mL) three times. Cell
pellets were then resuspended in 50 μl of 1% NP-40 lysis buffer [1% NP-40, 150 mM
NaCl, 50 mM triethanolamine (TEA) pH 7.4] with Complete Mini protease inhibitor
cocktail (Roche Biosciences) and Z-V AD-FMK (Enzo Life Sciences) for 15 minutes and
then centrifuged (10 min, 10,000 x g, 4 ℃). The supernatant (soluble cell lysate) was
collected and the protein concentration was determined by BCA assay (Pierce,
ThermoScientific). Samples were prepared at a protein concentration of 2 mg mL
-1
with
4X loading buffer (8% SDS, 40% glycerol, 0.4% bromophenol blue, 2.8%
B-mercaptoethanol, 200 mM Tris-HCl pH 6.8) and boiled at 5 min at 98 ℃. Forty μg of
protein was then loaded per lane for SDS-PAGE separation (Criterion™ TGX 4-20%,
Bio-Rad).
Caspase wild-type vs. Mutant chemoenzymatic/Biotin IP . H1299 cells at 90%
confluency were co-transfected with pCDNA Caspase-8 catalytically dead, wild-type
(HA-tagged, 20 μg) and either (AAA)-mutant (FLAG-tagged, 20 μg),
(S386,386A)-mutant (FLAG-tagged, 20 μg), or (S375A)-mutant (FLAG-tagged, 20 μg)
DNA using Liptofectamine 2000 (Invitrogen), according to manufacturer’s protocol.
228
Cells were harvested 48 hours post-transfection and subsequently lysed using SDS-lysis
before being subjected to chemoenzymatic transfer according to protocols described
above. Following chemoenzymatic transfer, protein was precipitated by MeOH/CHCl
3
and resuspended in 1% SDS buffer. CuAAC was preformed as described previously with
alkyne-azo-biotin. Following CuAAC, enrichment was conducted according to protocols
previously described. Following the final elution step, both eluents were combine and
proteins were then precipitated in ice cold methanol (1 mL) overnight in -20 °C. Protein
was collected by centrifugation (10 min, 10,000 x g, 4 °C), and the pellet was allowed to
air dry for 5 min, at which time 20 μL 4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM
TEA pH 7.4) was added to each sample. The mixture was sonicated in a bath sonicator to
ensure complete dissolution, and 20 μL of 2X loading buffer (20% glycerol, 0.2%
bromophenol blue, 1.4% β-mercaptoethanol) was then added. Samples were boiled for 5
min at 98 °C before being subjected to SDS-PAGE and Western blotting as described
above. Plasmids available upon request.
Peptide synthesis. All solid-phase peptide syntheses were conducted manually on
unprotected Rink amide ChemMatrix
®
resin, (PCAS BioMatrix) with an estimated
loading of 0.6 mmol g
-1
. Protected O-GlcNAcylated serine was prepared as previously
described (Marotta et al., 2012; Mitchell et al., 2001). Commercially available N-Fmoc
and side-chain protected amino acids (10 eq, Advanced ChemTech) were activated for 20
min with HBTU (10 eq, Novabiochem) and DIEA (20 eq, Sigma) and then coupled to the
resin for 1 h, bubbling with N
2 to mix. Reaction completion was checked using the Kaiser
229
test. Briefly, a small amount of resin was incubated with equal volumes of 5% w/v
ninhydrin in EtOH, 80% w/v phenol in EtOH, and 20 μM KCN in pyridine and heated to
99 °C for 5 min in a sealed tube. If necessary, a second coupling was conducted with 10
eq amino acid, 10 eq HOBt (Novabiochem) and 15 eq DCC (Sigma) for 2 h, with N
2
mixing. After successful coupling, the terminal Fmoc group was removed with 20% v/v
piperidine in DMF for 5 min with N
2 mixing, and then for an addition 15 min with fresh
20% piperidine in DMF. When peptides were completed, the final Fmoc group was
removed as described above and the N-terminal amine was acetylated with 5 eq each of
pyridine and acetic anhydride in DMF. Peptides were then cleaved from the resin by
incubating in cleavage cocktail (95:2.5:2.5 TFA/H2O/Triisopropylsilane) for 3.5 h at
room temperature. The peptides were then diluted ~1/10 in cold diethyl ether and
precipitated overnight (-80°C). The resulting suspensions were centrifuged (30 min,
5,000 x g, 4 °C) and the pellets were resuspended in fresh Et
2O and centrifuged again (30
min, 5,000 x g, 4 °C). The pellets were then resuspended in H
2O, flash frozen, and
lyophillized. At this point, glycopeptides 2 and 4 were deprotected with hydrazine
hydrate (80% v/v in MeOH) for 2 h, followed by concentration under vacuum. Crude
lyophillized material was purified by RP-HPLC (0-50% B gradient over 60 min; A =
0.1% trifluoroacetic acid in H
2O; B = 0.1% trifluoroacetic acid, 10% H 2O, 90%
acetonitrile) over a C18 semi-preparative column (Vydac). Purified peptides were then
characterized by ESI-MS (Agilent 6100 series Quadrupole LC-MS): 1, calculated = 720.3
Da (M + H
+
), observed = 720.2 Da (M + H
+
); 2, calculated = 923.4 Da (M + H
+
),
observed = 923.3 Da (M + H
+
); 3, calculated = 835.4 Da (M + H
+
), observed = 835.2 Da
230
(M + H
+
); 4, calculated = 1241.5 Da (M + H
+
), observed = 1241.2 Da (M + H
+
); 5,
calculated = 704.3 Da (M + H
+
), observed = 704.2 Da (M + H
+
); 6, calculated = 819.3 Da
(M + H
+
), observed = 819.2 Da (M + H
+
); 7, calculated = 803.4 Da (M + H
+
), observed =
803.4 Da (M + H
+
); 8, calculated = 1033.5 Da (M + H
+
), observed = 1033.2 Da (M + H
+
).
In vitro caspase-8 cleavage assay. Peptides 1-8 (1000X stock in DMSO) were added to 4
units of recombinant caspase-8 (Enzo Life Sciences) in 100 μl of general caspase activity
buffer (20 mM HEPES, 100 mM NaCl, 0.1% CHAPS, 10% sucrose, 1 mM EDTA, 10
mM DTT, pH 7.2) to give a final concentration of 2 mM. The mixtures were incubated
overnight at 37 ℃ and subjected to HPLC analysis by RP-HPLC. Purified peptides were
then characterized by ESI-MS (Agilent 6200 series Quadrupole LC-MS): 1 uncleaved,
calculated = 720.3 Da (M + H
+
), observed = 720.2 Da (M + H
+
); 1 cleaved, calculated =
505.2 Da (M + H
+
), observed = 505.3 Da (M + H
+
); 2 uncleaved, calculated = 923.4 Da
(M + H
+
), observed = 923.2 Da (M + H
+
); 2 cleaved, calculated = 505.2 Da (M + H
+
),
observed = 505.2 Da (M + H
+
); 3 uncleaved, calculated = 835.4 Da (M + H
+
), observed =
835.2 Da (M + H
+
); 3 cleaved, calculated = 549.2 Da (M + H
+
), observed = 549.2 Da (M
+ H
+
); 4 uncleaved, calculated = 1241.5 Da (M + H
+
), observed = 1241.5 Da (M - H
+
); 4
cleaved, calculated = 549.2 Da (M + H
+
), observed = 549.2 Da (M + H
+
); 5 uncleaved,
calculated = 704.3 Da (M + H
+
), observed = 704.2 (M + H
+
); 5 cleaved, calculated =
505.2 Da (M + H
+
), observed = 505.2 Da (M + H
+
); 6 uncleaved, calculated = 819.3 Da
(M + H
+
), observed = 819.2 Da (M + H
+
); 6 cleaved, calculated = 505.2 Da (M + H
+
),
observed = 505.2 Da (M + H
+
) 7 uncleaved, calculated = 803.4 Da (M + H
+
), observed =
231
803.4 (M + H
+
); 7 uncleaved, calculated = 803.4 Da (M + H
+
), observed = 803.3 Da (M +
H
+
); 7 cleaved, calculated = 548.2 Da (M + H
+
), observed = 548.2 Da (M + H
+
); 8
uncleaved, calculated = 1033.5 Da (M + H
+
), observed = 1033.2 Da (M + H
+
); 8 cleaved,
calculated = 549.2 Da (M + H
+
), observed = 549.2 Da (M + H
+
)
Calcium Phosphate transfection. One day before transfection, 5 x 10
5
target cells were
plated. One hour before transfection, media was exchanged for fresh media. To an
Eppendorf tube was added water (398 μL, autoclaved), 2M CaCl
2 62 μL (filter sterilized),
and 40 μL DNA (500 ng/μL, 20 μg total, phenol/chloroform extracted, EtOH
precipitated). Solution was mixed by pipetting, then DNA solution is added dropwise to a
falcon tube containing 500 μL 2X HBS (500 mM HEPES, 1.5 mM Na
2HPO 4, 280 mM
NaCl, 10 mM KCl, 12 mM dextrose, pH 7.05, filter sterilized) while pumping bubbles.
The resulting solution was added dropwise to target cells while swirling slowly. A fine
black precipitate was observed after 30 min and the media was opaque in color. Seven
hours after transfection, media was exchanged for fresh media.
Caspase-8 knockout with CRISPR. Expression vector pSpCas9(BB)-2A-Puro (PX459)
(Addgene no. 48139) was digested with BbsI, and a pair of annealed oligonucleotides
corresponding the first exon of human caspase-8 (CACCGCTCTTCCGAATTAATAGAC
and AAACGTCTATTAATTCGGAAGAGC (Kranz and Boutros, 2014), was ligated into
the guide RNA to generate SpCas9-Casp8, which was confirmed by sequencing (Laragen,
Culver City, CA). HeLa cells were then transiently transfected with SpCas9-Casp8 using
232
calcium phosphate transfection methods as described above. Forty-eight hours
post-transfection, 1 ug mL
-1
puromycin was added to the media for the next 72 h.
Caspase-8 knockout cells were then further selected for by inducing apoptosis with 10 ng
mL
-1
TNF-alpha and 1 ug ml
-1
cycloheximide for 24 h. Following clonal selection using
cloning cylinders, Western blot analysis showed five out of five selected colonies had no
detectable caspase-8. Colony number one was selected for sequencing and further
experiments. A 2 kb fragment of genomic caspase-8 DNA was amplified using PCR with
primers CTTTGTGACATGGTACCTGGC and GTGTCCGATAATAGGAGATGTT. And
additional round of PCR was then used to introduce attB recombination sites enabling
Gateway cloning (ThermoFisher) into donor vector pDONR201 according to the
manufacturer’s protocol. E. Coli (DH5α, ThermoFisher) were then transformed with
these plasmids and grown on antibiotic-containing agar plates. Twenty colonies were then
selected for sequencing (Laragen, Culver City, CA).
Caspase-8 Overexpression Co-IP . HeLa Caspase-8 CRISPR cells at 90% confluency
were co-transfected with pCDNA Caspase-8 catalytically dead, wild-type (HA-tagged, 20
μg) and wild-type (FLAG-tagged, 20 μg) DNA using Liptofectamine 2000 (Invitrogen),
according to manufacturer’s protocol). Forty eight hours post transfection, cells were
harvested by trypsinization and then lysed by resuspension in 150 μL of lysis buffer with
1 mM phenylmethanesulfonyl fluoride (PMSF, 200 mM stock in isopropyl alcohol) (10X
Lysis buffer, Cell Signaling) followed by sonication (5 sec on, 5 sec rest, 3X). Protein
concentration was determined using BCA Assay (Pierce, ThermoScientific), and 200 μg
233
of lysate was diluted to 1 μg μL
-1
in lysis buffer. Lysate was then pre-cleared with 30 μL
of magnetic protein-G beads (Cell Signaling, #8740) for 1 h at 4 ℃ with end-over-end
rotation. Inputs were generated at 2 μg μL
-1
using 4X loading buffer (8% SDS, 40%
glycerol, 0.4% bromophenol blue, 2.8% B-mercaptoethanol, 200 mM Tris-HCl pH 6.8)
before boiling for 5 min at 98 ℃. Magnetic beads were captured using a magnetic rack
and pre-cleared lysate was transferred to a new, pre-cooled tube. Either FLAG primary
antibody (1:50 dilution, Cell Signaling, #2368) or HA primary antibody (1:50 dilution,
Cell Signaling, #3724) was added to the pre-cleared lysate. The appropriate isotype
control was used at the same protein concentration as the primary antibody (Rabbit
isotype control, Cell Signaling, #3900). Thirty μL of magnetic protein-G beads were then
added and let incubate over night with end-over-end rotation at 4 ℃. Protein-G beads
were then washed 5X with 500 μL of lysis buffer containing 1 mM PMSF. Beads were
the pelleted gently be centrifugation (3 min, 2,000 x g, 4 ℃) before 3X loading buffer
(Cell Signaling, #7722) was added and samples were boiled for 5 min at 98 ℃. After
removal of magnetic beads, samples were analyzed by SDS-PAGE (4-20% Tris-Glycine
Gel, Bio-Rad) and analyzed by Western blotting.
Caspase-8 Overexpression Co-IP with inhibitor treatment. HeLa Caspsae-8 CRISPR
cells at 90% confluency were co-transfected with pCDNA Caspase-8 catalytically dead,
wild-type (HA-tagged, 10 μg) and wild-type (FLAG-tagged, 10 μg) DNA using
Liptofectamine 2000 (Invitrogen), according to manufacturer’s protocol). Seven hours
following transfection cells were replated at 7.5 x 10
5
per 10 cm dish for treatment. The
234
following day, cells were treated with either 5SGlcNAc for 16 h (200 μM, 1,000X stock
in DMSO), Thiamet-G for 20 h (20 μM, 1,000X stock in DMSO) or DMSO vehicle.
Following treatment, cells were harvested and IP was conducted according to the protocol
described above.
Caspase wild-type vs. mutant auto-activation. Caspase-8 knockout HeLa cells were
transiently transfected using Lipofectamine 2000 according to manufacturer’s protocol
with 19 μg of a pRetroX-Tet-on empty vector (Clontech) and 1 μg of a pcDNA3 vector
(ThermoFisher) containing either caspase-8, caspase-8(AAA), or caspase-8(WWW).
Media was exchanged after 6 h of transfection, and this time was defined as 0 h after
transfection. After the indicated lengths of time, cells in the media were combined with
cells that were collected by trypsinization. The cells were then pelleted by centrifugation
(6 min, 2,000 x g, 4 ℃) and washed with PBS (1 mL) three times. Cell pellets were then
resuspended in 50 μL of 1% NP-40 lysis buffer [1% NP-40, 150 mM NaCl, 50 mM
triethanolamine (TEA) pH 7.4] with Complete Mini protease inhibitor cocktail (Roche
Biosciences) and Z-V AD-FMK (Enzo Life Sciences) for 15 min and then centrifuged (10
min, 10,000 x g, 4 ℃). The supernatant (soluble cell lysate) was collected and the protein
concentration was determined by BCA assay (Pierce, ThermoScientific). Samples were
prepared at a protein concentration of 3 mg mL
-1
with 4X loading buffer (8% SDS, 40%
glycerol, 0.4% bromophenol blue, 2.8% B-mercaptoethanol, 200 mM Tris-HCl pH 6.8)
and boiled at 5 min at 98 ℃. Forty μg of protein was then loaded per lane for SDS-PAGE
separation (Criterion™ TGX 4-20%, Bio-Rad).
235
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243
Chapter 4. N-Propargyloxycarbamate Monosaccharides as Metabolic
Chemical Reporters of Carbohydrate Salvage Pathways and Protein
Glycosylation
‡
Introduction
An ever expanding repertoire of bioorthogonal reactions has enabled the specific labeling
of reporter molecules in a range of biological contexts (Prescher and Bertozzi, 2005;
Sletten and Bertozzi, 2011). In many applications, this two-step detection strategy relies
upon the metabolic delivery and subsequent enzymatic installation of a chemical reporter.
Emblematic of this technology, metabolic chemical reporters of glycosylation have been
in use for over a decade and contain the founding member of azide-containing reporters
an analogue of N-acetyl-mannosamine (ManNAc) termed N-azidoacetyl- mannosamine
(ManNAz), which was developed for the visualization of sialic acid-containing
carbohydrates upon reaction with an immuno-tag using the Staudinger ligation(Saxon,
2000). Inspired by this result, a variety of other metabolic chemical reporters have been
developed to target sialic acid modification,(Chang et al., 2009; Hsu et al., 2007;
Luchansky et al., 2004) mucin O-linked glycosylation(Hang et al., 2003; Zaro et al.,
2011a), fucosylation(Hsu et al., 2007; Rabuka et al., 2006), and intracellular O-GlcNAc
modification (O-GlcNAcylation)(V ocadlo et al., 2003; Zaro et al., 2011b). These
‡
Leslie A. Bateman and Balyn W. Zaro (University of Southern California) contributed to the work
presented in this chapter.
244
chemical reporters take advantage of carbohydrate scavenging pathways that convert
them into the corresponding nucleotide sugar-donors for utilization by
glycosyltransferases. Until recently, these metabolic chemical reporters were thought to
function largely in a one-input one-output paradigm, where treatment with one chemical
reporter would read-out on one type of glycoconjugate (e.g. ManNAz treatment results in
sialic acid labeling). However, cells are armed with metabolic pathways that can
enzymatically interconvert monosaccharides and uridine diphosphate (UDP) sugar
donors(Yarema and Bertozzi, 2001). For example, N-acetyl-glucosamine (GlcNAc) can
be reversibly converted to both N-acetyl-galactosamine (GalNAc)(Thoden, 2001) and
N-acetyl-mannosamine (ManNAc){Hinderlich:1997ti}. Therefore, it was not entirely
surprising when multiple reports demonstrated that the azide-containing chemical
reporter, N-azidoacetyl-galactosamine (GalNAz), can be transformed to
N-azidoacetyl-glucosamine (GlcNAz), resulting in the labeling of a combination of
mucin O-linked, O-GlcNAcylation, and some N-linked glycans(Banerjee et al., 2010;
Boyce et al., 2011; Zaro et al., 2011b). While these different types of glycosylation can be
separated using biochemical methods, a more ideal metabolic chemical reporter would
specifically read-out on only one type of glycosylation. Because each of the carbohydrate
scavenging pathways and glycosyltransferases is likely to display unique tolerance to
chemically-modified monosaccharides, it should be possible to create chemical reporters
that discriminate between glycosylation pathways. In support of this, we previously
reported on alkyne-containing chemical reporters, N-butynyl-glucosamine (GlcNAlk) and
N-butynyl-galactosamine (GalNAlk), which are not interconverted, enabling the more
245
selective visualization and identification of O-GlcNAc modified proteins using the
Cu(I)-catalyzed azide–alkyne cycloaddition (CuAAC) (Zaro et al., 2011b).
Herein, we continue to examine the chemical tolerance of mammalian glycosylation
pathways through the synthesis and characterization of the N-propargyloxycarbamate
(Poc) containing chemical reporters GlcPoc(Sridhar and Chandrasekaran, 2002), GalPoc,
and ManPoc(Yu et al., 2005) (Figure 4-1A). Interestingly, we find that each chemical
reporter displays unique labeling efficiencies and that qualitatively GlcPoc and GalPoc
are incorporated into the same proteins while ManPoc labels a different pattern. Finally,
we show that all three chemical reporters allow for the selective enrichment of a known
O-GlcNAc modified protein NEDD4, suggesting that they all can be metabolically
converted to enter the O-GlcNAcylation pathway.
246
Results
Fluorescence labeling of proteins by Poc analogs
Figure 4-1. N-Propargylcarbamate-containing metabolic chemical reporters incorporated
onto proteins. (A) Metabolic chemical reporters Ac 4GlcPoc:
N-propargylcarbamate-1,3,4,6-tetra-O-acetyl-glucosamine, Ac 4GalPoc:
N-propargylcarbamate-1,3,4,6-tetra-O-acetyl-galactosamine, Ac 4ManPoc:
N-propargylcarbamate-1,3,4,6-tetra-O-acetyl-mannosamine. (B) NIH3T3 cells were treated with
each chemical reporter (200 μM) for 16 hours before reaction with azido-rhodamine under
CuAAC conditions and analysis by in-gel fluorescence. Coomassie blue shows protein loading.
To test the ability of Poc-modified monosaccharides to transit carbohydrate salvage
pathways and serve as substrates for glycosyltransferases, the hydrochloride salts of
glucosamine, galactosamine, and mannosamine were reacted with
propargyl-chloroformate. The remaining free hydroxyl groups were subsequently
acetylated to give Ac
4GlcPoc, Ac 4GalPoc, and Ac 4ManPoc (Figure 4-1A). NIH3T3 cells
were metabolically labeled at 200 μM for 16 hours. The cells were washed, lysed, and the
soluble protein fraction was reacted with a previously reported azide-containing
247
rhodamine fluorescent dye (Az-Rho) using CuAAC(Zaro et al., 2011b). In-gel fluorescent
scanning showed labeling of a variety of proteins with all three chemical reporters
(Figure 4-1B). ManPoc labels cells at the highest level followed by GlcPoc and, finally,
GalPoc. Interestingly, GlcPoc and GalPoc qualitatively label the same subset of proteins,
while ManPoc enables visualization of a non-overlapping population, suggesting that the
Poc group is tolerated to different degrees by all the monosaccharide salvage pathways
but discriminated against at the level of the metabolic interconversion and/or
glycosyltransferases. To test the concentration dependence of the chemical reporters,
NIH3T3 cells were treated with various concentrations of each molecule for 16 hours,
followed by lysis, CuAAC with Az-Rho, and analysis by in-gel fluorescent scanning
(Figure 4-2A). All three reporters labeled proteins at concentrations as low as 50 μM.
248
Figure 4-2. Characterization of N-propargyloxycarbamate (Poc) bearing metabolic
chemical reporters. A) NIH3T3 cells were treated with the indicated concentrations of
Ac4GlcPoc, Ac 4GalPoc, or Ac 4ManPoc for 16 hours, followed by analysis by in-gel fluorescence
scanning. B) NIH3T3 cells were treated with 150 μM Ac4GlcPoc, Ac 4GalPoc, or Ac 4ManPoc for
the indicated times before analysis by in-gel fluorescence scanning. C) NIH3T3 cells were treated
with 150 μM Ac4GlcPoc, Ac 4GalPoc, or Ac 4ManPoc and chased with 150 μM Ac 4GlcNAc,
Ac4GalNAc, or Ac 4ManNAc, respectively, followed by in-gel fluorescence scanning. Coomassie
blue staining demonstrates protein loading.
249
A key feature of all metabolic chemical reporters is their ability to label a
newly-generated protein fraction in a pulse experiment; however, this requires that they
are efficiently incorporated on short time-scales. To qualitatively determine the rate of
chemical reporter labeling, NIH3T3 cells were treated with Ac
4GlcPoc (150 μM),
Ac
4GalPoc (200 μM), or Ac 4ManPoc (150 μM) for different amounts of time.
Fluorescent modification using CuAAC and in-gel fluorescence demonstrated
incorporation of all three chemical reporters in as little as 4 h (Figure 4-2B), consistent
with the labeling times of other reporters including GlcNAz and GlcNAlk(Zaro et al.,
2011b). Additionally, metabolic chemical reporters can be used to determine the stability
of a modification or protein in a pulse-chase format by measuring the decrease in
fluorescence signal over time. Accordingly, NIH3T3 cells were treated with Ac
4GlcPoc
(150 μM), Ac
4GalPoc (200 μM), or Ac 4ManPoc (150 μM) for 8, 16, and 16 hours,
respectively. At these times, the cells were washed with PBS and media containing the
corresponding unmodified sugar (Ac
4GlcNAc, Ac 4GalNAc, or Ac4ManNAc) at 150 mM
was added. Cells were treated for 0–72 hours before analysis by in-gel fluorescence as
above (Figure 4-2C). Interestingly, GlcPoc and GalPoc displayed similar signal decay
rates, again suggesting that they label the same protein substrates. In contrast, loss of
ManPoc signal was somewhat slower, supporting its incorporation into a different type of
glycosylation.
250
GlcPoc and ManPoc label cell surface proteins
We further analyzed the efficiency of the Poc metabolic chemical reporters using
flow-cytometry. Accordingly, NIH3T3 cells were treated with each chemical reporter at
150 mM concentration for 8 hours. At this time, the cells were fixed with
paraformaldehyde (PFA) and permeabilized with Triton X-100 before CuAAC in the
presence of Az-Rho. Consistent with our fluorescence scanning, ManPoc showed the
highest level of signal, followed by GlcPoc and finally GalPoc (Fig. 4-3). Again this
demonstrates that the Poc group is differentially tolerated by the carbohydrate salvage
pathways and/or glycosyltransferases.
Figure 4-3. Flow cytometry analysis of metabolic chemical reporter incorporation. NIH3T3
cells were treated with each chemical reporter (150 mM) for 16 hours. Cells were then fixed with
3.7% PFA and permeabilized with 0.1% Triton X-100 before reaction with azido-rhodamine
under CuAAC conditions and analysis by flow cytometry. Error bars represent S.E.M. of three
experiments.
Finally, to examine the metabolic interconversion of the Poc-bearing chemical reporters
we examined their ability to read out on O-GlcNAc modifications. Accordingly, NIH3T3
251
cells were treated with Ac 4GlcPoc (150 μM), Ac 4GalPoc (200 μM), or Ac 4ManPoc (150
μM) for 8 h. The corresponding cell-lysates were then reacted under CuAAC conditions
with a known azide-containing biotin affinity probe (azido-azo-biotin)(Zaro et al.,
2011b). Labeled proteins were enriched with streptavidin beads, washed, and eluted with
sodium dithionite. Western blot analysis of these enriched proteomes was then performed
using an antibody against a known O-GlcNAc substrate NEDD4 (Figure 4-4)(Zaro et al.,
2011b). Interestingly, NEDD4 was enriched by all three chemical reporters, with GlcPoc
and ManPoc displaying approximately the same level of enrichment followed by GalPoc,
demonstrating that both ManPoc and GalPoc can be metabolically converted to GlcPoc
and enter the O-GlcNAc modification pathway.
Figure 4-4. Incorporation of metabolic chemical reporters into the O-GlcNAcylation
pathway. NIH3T3 cells were treated with each chemical reporter (150 mM) for 16 hours.
Labelled proteins were reacted with azido-azo-biotin using CuAAC before streptavidin
enrichment and analysis of O-GlcNAc modification by anti-NEDD4 Western blotting.
Discussion and Conclusion
The continually expanding range of metabolic chemical reporters of glycosylation has
enabled the visualization and identification of many types of glycans and underlying
protein substrates. We and others have demonstrated that many chemical reporters can be
metabolically transformed and therefore label multiple glycosylation pathways. However,
252
given the complete chemical control of these small molecules, it might be possible to
structurally bias their metabolic fates to generate more specific reporters. To further test
this possibility, we have developed the N-propargyloxycarbonyl (Poc) bearing metabolic
chemical reporters GlcPoc, GalPoc, and ManPoc. Comparison of these reporters revealed
that they are all incorporated into large numbers of cellular proteins, which can be readily
visualized using both in-gel fluorescence scanning and flow cytometry. The pattern of
GlcPoc and GalPoc labeled proteins are approximately identical, suggesting that they are
incorporated into the same type of glycosylation. In contrast to the azide-bearing
chemical reporter GalNAz when compared to GlcNAz(Banerjee et al., 2010; Hubbard et
al., 2011), GalPoc is utilized at a much lower efficiency than GlcPoc, consistent with our
previous analysis of the structurally similar alkyne-containing derivative GalNAlk(Zaro
et al., 2011b). However, based on the enrichment of NEDD4, GalPoc can be
interconverted to GlcPoc, which does not occur with GalNAlk and other chemical
reporters with larger substituents at the N-acyl position(Yu et al., 2012; Zaro et al.,
2011b). This demonstrates that even small alterations to the structure of these chemical
reporters can influence their metabolism and final destination. Likewise, ManPoc can be
converted to GlcPoc and subsequently participate in O-GlcNAc modification of NEDD4.
Interestingly, the level of NEDD4 enrichment is similar in both GlcPoc and ManPoc
treated cells. Together with the much higher global levels of ManPoc incorporation and
its distinct labeling pattern, we believe that only a minority of ManPoc enters the
O-GlcNAcylation pathway, while the majority is transformed to sialic acid, supporting
previous in vitro experiments(Sridhar and Chandrasekaran, 2002).
253
Materials and Methods
All reagents used for chemical synthesis were purchased from Sigma-Aldrich unless
otherwise specified and used without further purification. All anhydrous reactions were
performed under argon atmosphere. Analytical thin-layer chromatography (TLC) was
conducted on EMD Silica Gel 60 F
254 plates with detection by potassium permanganate
(KMnO
4), anisaldehyde or UV . For flash chromatography, 60 Å silica gel (EMD) was
utilized.
1
H spectra were obtained at 600 MHz or 500 MHz on a Varian VNMRS-600 or
AMX-500. Chemical shifts are recorded in ppm (δ) relative to CHCl
3 (7.26 ppm) for
spectra acquired in CDCl
3 or methanol.
13
C spectra were obtained at 150 or 125 MHz on
the same instruments.
Chemical Synthesis.
Known compounds, N-(9-(2-(4-(6-azidohexanoyl)piperazine-1-
carbonyl)phenyl)-6-(diethylamino)-3H-xanthen-3-ylidene)-N-ethylethanaminium
(az-rho; (Charron et al., 2009)), and azido-azo-biotin ((Yang et al., 2010)) were
synthesized according to literature procedures as described as described in Chapter 2.
Compound 4.1 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-glucosamine
(Ac
4GlcPoc). Glucosamine HCl (1 g, 4.6 mmol), and sodium
bicarbonate (0.7 g, 7.88 mmol, Mallinckrodt) were dissolved in
H
2O (10 mL). To the stirring solution, propargyl chloroformate
O
AcO
NH
OAc
O
O
OAc
AcO
254
(679 μL, 6.96 mmol) was added dropwise. The solution was allowed to stir for 16 h at
room temperature. The reaction mixture was concentrated, washed with methanol (10
mL) and filtered. Resulting filtrate was concentrated and dissolved in pyridine (10 mL)
and stirred. Acetic anhydride (1.8 mL, 19.14 mmol) was then added and allowed to stir
for 16 h at room temperature. Purification by silica gel column chromatography (45%
EtOAc in Hexanes) afforded the product (853.2 mg, 52% yield) as a white solid.
1
H
NMR (600 MHz, CDCl
3) δ 5.70 (d, J = 8.3 Hz, 1H), 5.18 (t, J = 9.8 Hz, 1H), 5.11 (t, J =
9.6 Hz, 1H), 4.96 (d, J = 8.1 Hz, 1H), 4.66 (s, 1H), 4.28 (dd, J = 12.5, 4.4 Hz, 1H), 4.11
(dd, J = 12.4, 1.7 Hz, 1H), 4.06 (t, J = 6.7 Hz, 1H), 3.93 (dd, J = 19.0, 9.4 Hz, 1H), 3.81
(dd, J = 9.9, 4.6, 2.2 Hz, 1H), 2.45 (bs, 1H), 2.13 (s, 3H), 2.09 (s, 3H), 2.05 (s, 3H), 2.03
(s, 3H).
13
C NMR (150 MHz, CDCl3) δ 170.78, 169.48, 155.00, 92.60, 77.96, 73.00,
72.36, 68.01, 64.51, 61.76, 55.16, 53.07, 21.02, 20.87, 20.78, 20.73.
Compound 4.2 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-galactosamine
(Ac
4GalPoc). Galactosamine HCl (100 mg, 0.46 mmol,
Carbosynth), and sodium bicarbonate (133 mg, 1.58 mmol,
Mallinckrodt) were dissolved in H
2O (2.3 mL) and 1,4-dioxane (4
mL). To the stirring solution, propargyl chloroformate (68 μL, 0.7
mmol) was added dropwise. The solution was allowed to stir for 16 h at room
temperature. The reaction mixture was concentrated, washed with methanol (10 mL) and
filtered. Resulting filtrate was concentrated and purified by silica gel column
chromatography (20:80, methanol, methylene chloride). Resulting product was dissolved
O
AcO
NH
OAc
O
O
OAc
AcO
255
in pyridine (704 μL, 8.7 mmol) and stirred. Acetic anhydride (328 μL, 3.48 mmol) was
then added and allowed to stir for 16 h at room temperature. Reaction mixture was
concentrated and pyridine was removed. The crude was then resuspended in CH
2Cl 2 and
extracted with 1 M HCl, saturated sodium bicarbonate, water and brine. Purification by
silica gel column chromatography (45% EtOAc in Hexanes) afforded the product (118
mg, 79% yield) as a white solid.
1
H NMR (500 MHz, CDCl 3) δ 6.22 (d, 1H), 5.41 (d, J =
2.6 Hz, 1H), 5.17 (dd, J = 11.6, 2.9 Hz, 1H), 4.88 (d, J = 9.7 Hz, 1H), 4.73 – 4.58 (m,
2H), 4.41 (td, J = 11.4, 3.5 Hz, 1H), 4.22 (dd, J = 13.5, 6.7 Hz, 1H), 4.10 – 4.02 (m, 2H),
2.48 (t, J = 2.0 Hz, 1H), 2.16 (s, 6H), 2.01 (s, 6H).
13
C NMR (125 MHz, CDCl 3) δ
170.93, 170.47, 170.27, 169.96, 168.93, 155.01, 91.45, 75.16, 68.67, 68.09, 66.83, 61.35,
53.14, 48.92, 21.03, 20.81, 20.76, 20.74.
Compound 4.3 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-manosamine
(Ac
4ManPoc). Mannosamine HCl (100 mg, 0.46 mmol.
Carbosynth), and sodium bicarbonate (133 mg, 1.58 mmol,
Mallinckrodt) were dissolved in H
2O (2.3 mL) and 1,4-dioxane
(4 mL). To the stirring solution, propargyl chloroformate (68 μL, 0.7 mmol) was added
dropwise. The solution was allowed to stir for 16 h at room temperature. The reaction
mixture was concentrated and washed with methanol (10 mL) and filtered. Resulting
filtrate was concentrated and purified by silica gel column chromatography (15% MeOH
in CH
2Cl 2). Resulting product was dissolved in pyridine (671 μL, 8.3 mmol) and stirred.
Acetic anhydride (314 μL, 3.3 mmol) and DMAP (0.4 mg, 0.003 mmol) were then added
O
AcO
AcO
OAc
AcO
HN O
O
256
and allowed to stir for 16 h at room temperature. Reaction mixture was concentrated and
pyridine was removed. The cruse was then resuspended in CH
2Cl 2 and extracted with 1
M HCl, saturated sodium bicarbonate, water and brine. Purification by silica gel column
chromatography (50% EtOAc in Hexanes) afforded the product (123.5 mg, 86% yield) as
a white solid.
1
H NMR (500 MHz, CDCl 3) δ 6.07 (d, J = 1.8 Hz, 1H), 5.41 (d, J = 9.5 Hz,
1H), 5.29 (dd, J = 10.2, 4.3 Hz, 1H), 5.20 (d, J = 10.2 Hz, 1H), 4.68 (dt, J = 4.9, 2.5 Hz,
2H), 4.24 (dd, J = 12.7, 5.0 Hz, 1H), 4.11 – 3.96 (m, 3H), 2.49 (d, J = 2.2 Hz, 1H), 2.16
(d, J = 0.6 Hz, 3H), 2.08 (s, 4H), 2.04 (s, 4H), 2.01 (d, J = 3.7 Hz, 3H).
13
C NMR (125
MHz, CDCl
3) δ 170.78, 170.19, 169.71, 168.20, 155.18, 91.82, 75.22, 73.46, 70.28,
69.12, 65.39, 62.12, 53.14, 51.39, 20.95, 20.83, 20.72.
Cell culture. NIH3T3 cells were cultured in high glucose DMEM media (Cellgro) with
10% fetal calf serum (FCS, Cellgro) and were maintained in a humidified incubator at 37
°C and 5.0% CO
2.
Metabolic labeling. To cells at 80-85% confluency, high glucose media containing Poc
analog (1,000 x stock in DMSO), or DMSO vehicle was added as indicated. For chase
experiments, media was supplemented with 150 μM GlcNAc, GalNAc or ManNAc.
Preparation of NP-40-soluble lysates. The cells were collected by scraping and pelleted
by centrifugation at at 4 °C for 2 min at 2,000 x g, followed by washing with PBS (1 mL)
two times. Cell pellets were then resuspended and lysed in 75 μl of 1% NP-40 lysis
257
buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with Complete
Mini protease inhibitor cocktail (Roche Biosciences) for 15 min and followed by
centrifugation at 4 °C for 10 min at 10,000 x g. The resulting supernatant (soluble cell
lysate) was collected and separated to determine protein concentration via BCA assay
(Pierce, ThermoScientific).
Cu(I)-catalyzed [3 + 2] azide-alkyne cycloaddition (CuAAC). Soluble cell lysate (200
μg) was diluted with cold 1% NP-40 lysis buffer to a concentration of 1 μg/μL. Newly
made click chemistry cocktail (12 μL) was added to each sample [azido-rhodamine tag
(100 μM, 10 mM stock solution in DMSO); tris(2-carboxyethyl)phosphine hydrochloride
(TCEP) (1 mM, 50 mM freshly prepared stock solution in water); tris[(1-benzyl-1-H-
1,2,3-triazol-4-yl)methyl]amine (TBTA) (100 μM, 10 mM stock solution in DMSO); Cu-
SO
4•5H 2O (1 mM, 50 mM freshly prepared stock solution in water) for a total reaction
volume of 200 μL. The reaction was gently vortexed and allowed to sit at room
temperature for 1 h. Upon completion, 1 mL of ice cold methanol was added to the
reaction, and proteins were precipitated at -20 °C for 2 h. The reactions were then
centrifuged at 4 °C for 10 min at 10,000 x g. The supernatant was removed, the pellet was
allowed to air dry for 5 min, and then 50 μL 4% SDS buffer (4% SDS, 150 mM NaCl, 50
mM TEA pH 7.4) was added to each sample. The mixture was sonicated in a bath
sonicator to ensure complete dissolution, and 50 μL of 2x loading buffer (20% glycerol,
0.2% bromophenol blue, 1.4% β-mercaptoethanol) was then added. The samples were
boiled for 5 min at 98 °C, and 40 μg of protein was then loaded per lane for SDS- PAGE
258
separation (Any kD Criterion Gel, Bio-Rad).
In-gel Fluorescence Scanning. The gel was scanned on a Molecular Imager FX
(Bio-Rad) using a 580 nm laser for excitation and a 620 nm bandpass filter for detection.
Biotin Enrichment. NIH3T3 cell pellets labeled with GlcPoc, GalPoc, ManPoc (150 μM)
or DMSO were resuspended in 13 μL H 2O, and 25 μL 0.05% SDS buffer (0.05% SDS, 10
mM TEA pH 7.4, 150 mM MgCl
2) with Complete Mini protease inhibitor cocktail
(Roche Biosciences). To this was added 1 μL Benzonase (Sigma), and the cells were
incubated on ice for 30 min. At this time, 4% SDS buffer (100 μL) was added, and the
cells were briefly sonicated in a bath sonicator and collected by centrifugation at 20,000 x
g for 10 min at 15 °C. Protein concentration was normalized by BCA assay (Pierce,
ThermoScientific) to 1 mg/mL (1 mg total cell lysate). The appropriate amount of click
chemistry cocktail was added and the reaction was allowed to proceed for 1 h, after
which time 10 volumes of ice-cold methanol were added. Precipitation proceeded 2 hours
at -20 °C. Precipitated proteins were centrifuged at 5,200 x g for 30 min at 0 °C and
washed 3x with 10 mL ice-cold MeOH, with resuspension of the pellet each time. The
pellet was then air-dried for 1 h. To capture the biotinylated proteins by streptavidin
beads, the air-dried protein pellet was resuspended in 400 μL of resuspension buffer (6 M
urea, 2 M thiourea, 10 mM HEPES pH 8.0) by bath sonication. Samples were then
transferred to 2 mL dolphin-nosed tubes containing streptavidin beads (25 μL) that were
pre-washed (2x with PBS (1 ml) and 1x with resuspension buffer (2,000 x g, 2 min)).
259
Samples were then incubated on a rotator for 2 h. Beads were washed 2x with
resuspension buffer (1 mL), 2x in PBS (1 mL) and 2x with 1% SDS in PBS buffer (1 mL)
and collected by centrifugation (2,000 x g, 2 min). Beads were then incubated in 25 μL of
sodium dithionite solution (1% SDS, 25 mM sodium dithionite) for 30 min at room
temperature to elute captured proteins. The beads were centrifuged for 2 min at 2,000 x g
and the eluent collected. The elution step was repeated and the eluents combined. Protein
was precipitated in ice cold methanol (1 mL) overnight in -20 °C. Protein was collected
by centrifugation (10 min, 10,000 x g, 4 °C), and the pellet was allowed to air dry for 5
min, and then 30 μL 4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) was
added to each sample. The mixture was sonicated in a bath sonicator to ensure complete
dissolution, and 30 μL of 2x loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4%
β-mercaptoethanol) was then added. The samples were boiled for 5 min at 98 °C, and
samples were loaded into a gel for SDS-PAGE separation (Any kD Criterion Gel,
Bio-Rad).
Western Blotting. Proteins were separated by SDS-PAGE before being transferred to
PVDF membrane (Bio-Rad) using standard Western blotting procedures. Briefly, all
Western blots were blocked in TBST (0.1% Tween-20, 150 mM NaCl, 10mM Tris pH
8.0) containing 5% non-fat milk for 1 h at rt. They were then incubated with the
appropriate primary antibody in blocking buffer overnight at 4 °C. The anti-NEDD4
WW2 antibody (Millipore) was used at a 1:10,000 dilution. The blots were then washed
three times in TBST and incubated with the horseradish peroxidase(HRP)-conjugated
260
secondary antibody for 1 h in blocking buffer at RT. HRP-conjugated anti-mouse and
anti-human antibodies (Jackson ImmunoResearch) were used at 1:10,000 dilutions. After
being washed three more times with TBST, the blots were developed using ECL reagents
(Bio-Rad) and the ChemiDoc XRS+ molecular imager (Bio-Rad).
Flow Cytometry. NIH3T3 cells were treated with GlcPoc, GalPoc, ManPoc (150 μM) or
DMSO for 16 h. Cells were collected and washed 2X with cold PBS and fixed with 3.7%
PFA in PBS for 10 min. Cells were then washed 1x with 2% FCS in PBS and
permeabilized (0.1% Triton X-100 in PBS for 10 min at room temperature). Cell were
washed with PBS and blocked for 10 min with 2% FCS in PBS. Cells were resuspended
in 100 μL PBS that contained 100 μM Az-Rho, 1 mM TCEP, 100 μM TBTA and 1 mM
CuSO
4•5H 2O. Samples were incubated in the dark for 1 h and washed 5x with 1%
Tween-20 and 0.5 mM EDTA in PBS and 1x with 2% FCS in PBS. Flow cytometry
analysis was then performed on a Beckman Coulter LSR II at USC Flow Cytometry
Core.
261
Chapter 4 References
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Chapter 5. A Chemical Reporter for Visualizing Metabolic Cross-Talk
between Carbohydrate Metabolism and Protein Modification
§
Introduction
An increasing number of posttranslational modifications (PTMs) have been discovered
that can have dramatic effects on the function (i.e., activity, localization, stability, etc.) of
substrate proteins. To catalog and investigate these important modifications, a variety of
chemical approaches have been developed to visualize and identify PTMs in cell lysates,
living cells and in vivo (Agard and Bertozzi, 2009). One of the most successful chemical
technologies involves the biosynthetic incorporation of synthetic analogs of endogenous
PTMs onto proteins in living cells or animals (Grammel and Hang, 2013). Typically,
these metabolic chemical reporters (MCRs) contain unique chemical-functionalities that
can undergo bioorthogonal reactions to install visualization or affinity tags. Until recently,
research using MCRs has primarily focused on the end-point of their biosynthetic
incorporation, namely the specific PTM of interest. However, because MCRs must be
metabolically transformed, typically into high-energy donor substrates [e.g.,
uridine-diphosphate (UDP) monosaccharides or acetyl-CoA], they provide a direct
opportunity to chemically track cellular metabolism. For example, we and others
demonstrated that after the azide-containing MCR N-azidoacetyl glucosamine (GlcNAz)
is metabolized into UDP-GlcNAz, it can be enzymatically converted to UDP
§
Balyn W. Zaro (University of Southern California) contributed to the work presented in this chapter.
266
N-azidoacetyl galactosamine (UDP-GalNAz), resulting in the incorporation into at least
three classes of glycoproteins (Boyce et al., 2011; Zaro et al., 2011b). While this
“metabolic crosstalk” is less than ideal for the analysis of a single type of glycosylation, it
raises the possibility that MCRs could be used to isolate, analyze and potentially discover
different branching biosynthetic-pathways from common metabolic intermediates (Figure
5-1). For example, one recently-discovered branching pathway involves metabolism from
the N-acetyl glucosamine (GlcNAc) salvage pathway (Boehmelt et al., 2000) to protein
acetylation (Shahbazian and Grunstein, 2007). Specifically, Varki and co-workers
demonstrated that the previously uncharacterized enzyme
amidohydrolase-domain-containing 2 (AMDHD2) converts GlcNAc-6-phosphate into
glucosamine-6-phosphate and acetate {Bergfeld:2012co}. This acetate might then be
activated on CoA and subsequently used for protein acetylation. While it had been
previously demonstrated that acetyl-CoA was required for de novo synthesis of
UDP-GlcNAc from glucose through the hexosamine biosynthetic pathway (Love and
Hanover, 2005; V ocadlo et al., 2003), these data reveal that under certain nutrient or
metabolic conditions, cells may utilize scavenged GlcNAc for not only for the
biosynthesis of glycans but also for other posttranslational modifications.
267
Figure 5-1. Using metabolic chemical reporters (MCRs) to detect cellular metabolism. A)
Salvaged N-acetyl glucosamine (GlcNAc) can enter a linear biosynthetic pathway that yields
UDP-GlcNAc that can be directly incorporated onto glycoproteins. Additionally, GlcNAc
metabolic intermediates can enter branching pathways to generate acetate and other
monosaccharides. B) MCRs have the potential to isolate branching metabolic pathways, like the
transformation of GlcNAc into acetate and subsequent acetylation of proteins.
Here, we report the development of a MCR that isolates the metabolism of GlcNAc into
posttranslational modifications that are not glycosylation. This MCR, termed
1-deoxy-GlcNAlk, builds upon our published chemical reporter for glycosylation,
N-pentynyl glucosamine (GlcNAlk) (Zaro et al., 2011b), but structurally lacks the
1-hydroxyl group that is absolutely required for biosynthesis into the corresponding
UDP-monosaccharide and subsequent incorporation into glycans. Treatment of a variety
of cells with 1-deoxy-GlcNAlk, followed by copper-catalyzed azide-alkyne cycloaddition
(CuAAC) with a fluorescent tag, resulted in differential labeling that is detectable in a
majority of cell-lines. Notably, the intensity of this signal was inhibited by the addition of
268
the acetyl-transferase inhibitor curcumin and competition with sodium acetate, suggesting
that some of the protein labeling is a result of lysine acetylation. Furthermore, proteomic
analysis using 1-deoxy-GlcNAlk identified 60 known acetylated proteins. Finally,
labeling of the acetylated-proteins histones H1.1 and H2B was confirmed using in-gel
fluorescence scanning. These data demonstrate that 1-deoxy-GlcNAlk is a MCR of
protein modification and more importantly suggest that MCRs can be used to characterize
and potentially discover branching metabolic-pathways in living cells.
Results
Fluorescent detection of 1-DeoxyGlcNAlk labeling
To create a MCR capable of isolating the cellular metabolism of GlcNAc into protein
modifications that are not glycosylation, we synthesized a structural analog of our
previously-published glycoprotein MCR, GlcNAlk (Scheme 5-1) (Zaro et al., 2011b).
This analog, 1-deoxy-GlcNAlk, lacks the 1-hydroxyl group and therefore cannot enter
any glycosylation metabolic-pathways. Additionally, we generated the per-acetylated
derivative, 1-deoxy-Ac
3GlcNAlk, as the acetates allow for passive diffusion of the MCR
into living cells where they are subsequently removed by esterases (Saxon, 2000).
NIH3T3 cells were treated with either 1-deoxy-Ac
3GlcNAlk (200 μM),
1-deoxy-GlcNAlk (10 mM) or Ac
4GlcNAlk (200 μM) as a positive control. After 16
hours, the corresponding cell-lysates were subjected to the bioorthogonal reaction
copper-catalyzed azide-alkyne cycloaddition (CuAAC) with a fluorescent tag,
azido-rhodamine (az-rho). In-gel fluorescence scanning revealed that both versions of the
269
1-deoxy MCR were robustly incorporated onto proteins (Figure 5-2A), albeit at a lower
level than the highly-efficient GlcNAlk. The per-acetylated MCR, 1-deoxy-Ac
3GlcNAlk,
was incorporated more efficiently than 1-deoxy-GlcNAlk, consistent with other MCRs
(Saxon, 2000), and was therefore used in all our subsequent experiments.
Scheme 5-1. Synthesis of Ac 3-1-DeoxyGlcNAc, Ac 3-1-DeoxyGlcNAlk, and 1-DeoxyGlcNAlk.
(a) Acetyl Chloride 16 h, RT, 38% yield; (b) Tributyltin Hydride, AIBN, Toluene, 1.5 h, Reflux,
98% yield; (c) 2.5 M HCl, 2 h, Reflux, 75% yield; (d) i. DCC, 4-Pentynoic Acid, TEA, cat.
DMAP, DMF, 16 h, RT ii. Acetic Anhydride, Pyridine, 16 h, RT, 33% yield; (e) NaOMe, MeOH,
pH 9, 1 h, RT, 84% yield.
We and others have previously demonstrated that protein labeling by certain MCRs can
be competed by the availability of specific nutrients in cell culture. For example, MCRs
that largely read out on the intracellular glycosylation O-GlcNAc modification can be
competed by increasing glucose concentrations (Zaro et al., 2011b), and increasing the
amount of serum can inhibit the incorporation of radio-labelled glucosamine
270
{Liepkalns:1983tg}. To investigate the sensitivity of 1-deoxy-GlcNAlk to different
cell-culture conditions, NIH3T3 cells were treated with 1-deoxy-Ac
3GlcNAlk (200 μM)
in the presence of low or high glucose concentrations (1.0 vs. 4.5 g/mL) or three different
amounts of serum (0, 2 or 10% v/v). In-gel fluorescence scanning, following lysis and
CuAAC with az-rho, demonstrated that 1-deoxy-GlcNAlk labeling is largely insensitive
to these different culture conditions (Figure 5-3A).
271
Figure 5-2. Characterization of proteins that are labeled by the MCR 1-deoxy-GlcNAlk. A)
NIH3T3 cells were treated with the indicated MCRs for 16 hours before the corresponding lysates
were subjected to CuAAC with az-rho and analyzed by in-gel fluorescence scanning. B) NIH3T3
cells were treated with 1-deoxy-Ac3GlcNAlk with or without sodium acetate for 6 hours before
CuAAC az-rho and in-gel fluorescence scanning. C) NIH3T3 cells were pretreated with the
protein acetyltransferase inhibitor curcumin for 30 min before addition of 1-deoxy-Ac3GlcNAlk
for an additional 5.5 hours. Labeled proteins were then visualized using in-gel fluorescence
scanning following CuAAC with az-rho. Coomassie blue staining shows equal loading.
272
1-deoxy-GlcNAlk is metabolized and incorporated onto acetylated proteins
To determine if any 1-deoxy-GlcNAlk labeling could be attributable to protein
acetylation, we used sodium acetate and the p300-specific acetyltransferase inhibitor
curcumin {Das:2009dn}. NIH3T3 cells were treated with or without sodium acetate (10
mM) and 1-deoxy-Ac
3GlcNAlk (200 μM) for 6 hours. In-gel fluorescence showed that
sodium acetate was able to compete 1-deoxy-GlcNAlk labeling (Figure 5-2B). To
investigate whether any observed protein acetylation by 1-deoxy-GlcNAlk is enzymatic
in nature, NIH3T3 cells were pretreated with curcumin (60 μM) for 30 minutes prior to
treatment with 1-deoxy-Ac
3GlcNAlk (200 μM) for 5.5 hours. Cell lysates were then
subjected to CuAAC with az-rho and analyzed by in-gel fluorescence scanning (Figure
5-2C). Curcumin-treated cells also showed reduced 1-deoxy-GlcNAlk labeling compared
to controls. Notably, in both of these experiments, the effect on the labeling of different
proteins was not uniform. For example, labeled proteins in the region of histones and
other small proteins (~15 kDa) are more sensitive to both competition by sodium acetate
and curcumin treatment. Together, these data suggest that pentynoic acid is likely
removed from 1-deoxy-GlcNAlk, where it is know to be enzymatically incorporated into
protein acetylation (Yang et al., 2010) and potentially other protein modifications. We
next directly compared 1-deoxy-Ac
3GlcNAlk to the known acetylation reporter sodium
pentynoate (Yang et al., 2010). Specifically, NIH3T3 cells were treated with
1-deoxy-Ac
3GlcNAlk (200 μM) or sodium pentynoate (200 or 5000 μM) for 8 hours.
Visualization of the labeled proteins by in-gel fluorescence showed that sodium
pentynoate is a more efficient MCR, even at equal concentrations (Figure 5-3B). Notably,
273
the pattern of proteins that are labeled by 1-deoxy-GlcNAlk and pentynoate are also
different. Together, these data suggest that while at least some of the proteins that become
modified by 1-deoxy-Ac
3GlcNAlk treatment are acetylated, its metabolism and/or
distribution into different types of posttranslational modification (e.g., acetylation vs.
long-chain fatty acylation) are different than sodium pentynoate.
Figure 5-3. Characterization of 1-deoxy-GlcNAlk labeling. A) 1-deoxy-GlcNAlk labeling is
not affected by glucose or serum levels in media. NIH-3T3 cells were treated with
1-deoxy-Ac3GlcNAlk (200 μM) for 16 h under high- or low-glucose conditions, 4.5 g/L or 1 g/L
respectively, supplemented with 10%, 2% or 0% fetal calf serum. Lysate from treated cells was
subjected to CuAAC and in-gel fluorescence scanning. B) Comparison of 1-deoxy-Ac3GlcNAlk
and sodium 4-pentynoate labeling. MEFs were treated with or without 1-deoxy-Ac3GlcNAlk (200
μM), sodium 4-pentynoate (200 μM or 5 mM) or DMSO vehicle for 8 h. Lysate from treated cells
was subjected to CuAAC following by SDS-PAGE separation and in-gel fluorescence scanning.
To further characterize this MCR, NIH3T3 cells were treated with various concentrations
of 1-deoxy-Ac
3GlcNAlk for 16 hours prior to lysis and CuAAC with az-rho. In-gel
fluorescence scanning showed that proteins are dose-dependently labelled by 50-200 μM
1-deoxy-Ac
3GlcNAlk treatment (Figure 5-4A). To determine the kinetics and dynamics
274
of 1-deoxy-GlcNAlk labeling, we next performed pulse and pulse-chase experiments. We
first treated NIH3T3 cells with 1-deoxy-Ac
3GlcNAlk (200 μM) for different lengths of
time. After lysis and CuAAC with az-rho, in-gel fluorescence scanning revealed protein
labeling in as little as 2 hours, with similar kinetics to other direct MCRs of protein
acetylation (Figure 7-4B) (Yang et al., 2010). NIH3T3 cells were then treated with
1-deoxy-Ac
3GlcNAlk (200 μM) for 16 hours, after which time the growth medium was
replaced with fresh media containing 1-deoxy-N-acetyl glucosamine
(1-deoxy-Ac
3GlcNAc, 200 μM). In-gel fluorescence scanning after CuAAC revealed a
time-dependent loss of signal (Figure 5-4C). To ascertain the generality of
1-deoxy-GlcNAlk as a MCR, a small panel of cell-lines were treated with
1-deoxy-Ac
3GlcNAlk (200 μM) for 16 hours before lysis and reaction with az-rho using
CuAAC. In-gel fluorescence scanning showed labeling of proteins in each of the
cell-lines tested (Figure 5-5). To determine if treatment of cells with the MCR resulted in
any toxicity, NIH3T3 cells were treated with either 1-deoxy-Ac
3GlcNAc (200 μM),
1-deoxy-Ac
3GlcNAlk (200 μM) or DMSO vehicle. After 24 or 48 hours of treatment, the
viability of the cells was measured using a commercially available MTS assay (Figure
5-6). No toxicity was observed with 1-deoxy-Ac
3GlcNAlk treatment, despite some
toxicity with the control compound 1-deoxy-Ac
3GlcNAc.
275
Figure 5-4. Dose-dependence and dynamics of 1-deoxy-GlcNAlk protein labeling. A)
NIH3T3 cells were treated with the indicated concentrations of 1-deoxy-Ac3GlcNAlk for 16
hours before CuAAC with az-rho and analysis by in-gel fluorescence scanning. B) NIH3T3 cells
were treated with 1-deoxy-Ac3GlcNAlk (200 μM) for the indicated lengths of time. Cell lysates
were reacted with az-rho and visualized with in-gel fluorescence scanning. C) NIH-3T3 cells
were treated with 1-deoxy-Ac3GlcNAlk (200 μM) for 16 h after which time media was
exchanged with fresh media containing 1-deoxy-Ac3GlcNAc (200 μM) for the indicated lengths
of time. Time dependent loss of protein labeling was visualized using in-gel fluorescence
scanning. Coomassie blue staining shows equal loading.
276
Figure 5-5. Generality of 1-deoxy-GlcNAlk labeling. The indicated cell lines were treated with
200 μM 1-deoxy-Ac3GlcNAlk for 16 hours before modified proteins were subjected to CuAAC
with az-rho and in-gel fluorescent scanning.
Figure 5-6. Toxicity of 1-deoxy-Ac 3GlcNAlk. Cell viability following treatment for 24-48 h
with 1-deoxy-Ac3GlcNAc, 1-deoxy-Ac 3GlcNAlk or DMSO vehicle was measured using a
commercially available MTS assay (Promega). Quantitation is from three individual experiments
normalized to DMSO treated cells; error bars indicate ±s.e.m.
277
Identification of 1-deoxy-GlcNAlk labelled proteins
Finally, we performed a large-scale mass spectroscopy experiment to identify proteins
labeled by 1-deoxy-GlcNAlk and directly compare them to those modified by our
published MCR GlcNAlk. NIH3T3 cells were treated in triplicate with 1-deoxy-GlcNAlk
(200 μM), GlcNAlk (200 μM) or GlcNAc (200 μM) as a negative control for 16 hours.
Treated cells were pelleted and lysed with a denaturing buffer (4% SDS). Protein
concentration was normalized, and 10 mg of protein was subjected to CuAAC with an
azide-functionalized biotin affinity-tag. The biotinylated samples were enriched with
streptavidin beads, washed extensively and subjected to on-bead trypsin digestion, and
the recovered peptides were subjected to LC-MS/MS analysis. Proteins were identified
using Proteome Discover and Mascot and curated using the following criteria to identify
“hits”: (1) Proteins must be identified in all 3 runs (at least 1 spectral count per run) with
a sum of at least 4 spectral-counts overall; (2) The sum of the spectral counts must be
4-fold greater in the 1-deoxy-GlcNAlk or GlcNAlk samples than the GlcNAc-treated
samples; (3) The number of spectra counts in the MCR-treated sample compared to the
control must be statistically significant (p-value < 0.05, t test). Following these
requirements, we identified 99 proteins modified by 1-deoxy-GlcNAlk (Figure 5-7A) and
433 proteins modified by GlcNAlk (Figure 5-8). Of the 1-deoxy-GlcNAlk labeled
proteins, 60 have been previously identified as acetylated proteins, including those
annotated in Figure 5-7A, supporting this MCR as a reporter of acetylation. Forty-six
proteins identified using 1-deoxy-GlcNAlk were also present in the GlcNAlk treated
278
samples (Figure 5-7B), suggesting that these proteins are either simultaneously modified
by O-GlcNAc glycosylation or that GlcNAlk can also be metabolized through an “off-
target” pathway. Notably, 16 of these overlapping proteins are known acetylated proteins,
suggesting that both MCRs may be metabolized into the protein acetylation pathway.
The proteins identified using 1-deoxy-GlcNAlk also contained 39 previously
uncharacterized substrates, suggesting that this MCR can be used to find new
modification (e.g., acetylation) events. To confirm 1-deoxy-GlcNAlk labeling of known
acetylated proteins, Histones H1.1 and H2B.(7) Histones were enriched from NIH-3T3
cells treated with 1-deoxy-Ac3
GlcNAlk (200 μM) or 1-deoxy-Ac
3
GlcNAc (200 μM) as
negative control for 16 h using acid precipitation (Shechter et al., 2007). Purified
histones were then subjected to CuAAC with az-rho and in-gel fluorescence scanning
confirmed labeling of Histones 1.1 and H2B (Figure 5-7C).
Figure 5-7. Identification of posttranslationally modified proteins using 1-deoxy-GlcNAlk.
(A) NIH3T3 cells were treated with 1-deoxy-Ac3GlcNAlk, Ac 4GlcNAlk or Ac 4GlcNAc (all at
200 μM concentration) for 16 hours. At this time, the corresponding cell-lysates were subjected to
CuAAC with azide-biotin, enrichment with streptavidin-coated beads, and on-bead trypsinolysis.
Proteins identified by LC-MS/MS are graphically presented as total number of positive minus
total number of control spectral counts. Three known acetylated proteins are annotated in black.
(B) Overlap between proteins identified using 1-deoxy-Ac3GlcNAlk and Ac 4GlcNAlk. (C)
Enriched histones from NIH3T3 cells labeled with 1-deoxyAc3GlcNAlk or DMSO vehicle were
subjected to CuAAC with az-rho. In-gel fluorescence reveals modification of Histones 1.1 and
2B.
279
Discussion
Bioorthogonal chemistries have enabled the creation of MCRs for the visualization and
enrichment of a wide array of PTMs (Grammel and Hang, 2013) including glycosylation
(V ocadlo et al., 2003; Zaro et al., 2011a; 2011b), lipidation (Hang and Linder, 2011),
methylation (Wang et al., 2013b), and different forms of acetylation (Bao et al., 2013;
Yang et al., 2010). Because MCRs must be metabolized by living cells, they provide
unique opportunities to simultaneously interrogate a certain PTM and the upstream
metabolic and biosynthetic pathways. We have demonstrated that alterations in the
chemical structure of a MCR can impact its acceptance into different glycosylation
pathways (Zaro et al., 2011b). Building upon those results, we synthesized and
characterized a MCR (1-deoxy-GlcNAlk) that reports on the metabolic crosstalk between
the GlcNAc salvage pathway and non-glycosylation modifications on proteins. Using a
fluorescent azide-tag, in combination with CuAAC, we demonstrated that
1-deoxy-GlcNAlk treatment results in labeling of a range of proteins in different cell
lines. The labeling intensities in these cell-lines varies dramatically, raising the possibility
that MCRs could be used to classify metabolic flux in different cells. Co-treatment with
different nutrient sources and an inhibitor of protein acetyltransferases showed that
1-deoxy-GlcNAlk labeling is competed by exogenous acetate and acetyltransferase
inhibition. This demonstrates that at least some 1-deoxy-GlcNAlk enters the protein
acetylation pathway. This is further supported by our proteomic identification of 60
previously identified, acetylated proteins, which account for ~60% of the total proteins
identified. The most likely pathway responsible for these observations is the one
280
identified by Varki and co-workers mentioned above {Bergfeld:2012co}. In this case,
1-deoxy-GlcNAlk would be phosphorylated and then deacetylated by the enzyme
AMDHD2 to generate pentynoic acid, although this remains to be experimentally
confirmed.
Figure 5-8. Identification of proteins labelled by GlcNAlk. NIH3T3 cells were treated with
Ac4GlcNAlk or Ac 4GlcNAc (both at 200 μM concentration) for 16 hours. At this time, the
corresponding cell-lysates were subjected to CuAAC with azide-biotin, enrichment with
streptavidin-coated beads, and on-bead trypsinolysis. Proteins identified by LC-MS/MS are
graphically presented as total number of positive minus total number of control spectral counts.
Three known O-GlcNAcylated proteins are annotated in black.
However, not all of the labeled proteins were equally susceptible to competition by
sodium acetate or inhibition of the p300 acetyltransferase (Figures 5-2B and Figure
5-2C). In the case of sodium acetate competition, the intensity of all of the labeled
proteins is reduced; however, the proteins at ~15 kDa molecular weight display a more
dramatic effect. This difference could be attributable to acetylation dynamics. Rapidly
cycling acetylation marks, like those on the core histones that are found around 15 kDa
281
{Waterborg:2002jx, KatanKhaykovich:2002eo}, could be more sensitive to competition
by excess sodium acetate. In contrast, any long-lived pentynyl-modification events could
persist throughout the experiment. Likewise, treatment with curcumin resulted in
dramatic reduction of the labeling of proteins at low molecular weights but less-so for
other proteins. Since curcumin is a specific inhibitor of the p300 acetyltransferase
{Das:2009dn}, the proteins that show no change in labeling intensity might be modified
by other acetyltransferases. We next directly compared 1-deoxy-GlcNAlk with pentynoic
acid. At equal concentrations, 1-deoxy-GlcNAlk is significantly less efficient at labeling
proteins, and pentynoate-labeling can be performed at higher concentrations to maximize
incorporation (Figure 5-3B). Interestingly, 1-deoxy-GlcNAlk and pentynoate treatment
resulted in the visualization of different patterns of proteins. This demonstrates that
1-deoxy-GlcNAlk is not a simple replacement of a known MCR of protein acetylation
(Yang et al., 2010). The differences between the two MCRs could simply arise from
changes in their metabolism. For example, if the two MCRs are metabolized at different
rates, a different subset of proteins could be modified after the same length of labeling. It
is also possible that the two MCRs are incorporated into different types of
posttranslational modifications. For example, short-chain fatty acid reporter could be
metabolized into the corresponding lipid-reporter (e.g., palmitoylation) (Charron et al.,
2009). It is also possible that either pentynoate or 1-deoxy-GlcNAlk is metabolized into
an unknown, non-acetylation pathway that contributes to some of the signal, or results in
non-enzymatic modification of proteins (Bateman et al., 2013b; Moellering and Cravatt,
2013).
282
Finally, to compare 1-deoxy-GlcNAlk to a glycoprotein MCR, we performed a
proteomics experiment using 1-deoxy-GlcNAlk and GlcNAlk. Enrichment with
1-deoxy-GlcNAlk resulted in the identification of 99 proteins. Treatment with GlcNAlk
resulted in the identification of a large number of O-GlcNAc modified proteins and 64
proteins that overlapped with the 1-deoxy-GlcNAlk sample. Notably, 16 of these proteins
were also previously identified as being acetylated. This raises the likely possibility that
any glycoprotein MCR bearing its chemical functionality at the N-acetyl position will
read-out on some acetylated proteins. Therefore, care should be taken to confirm the
glycosylation of candidate proteins identified using these reporters.
Conclusion
In summary, our competition, inhibition and proteomics experiments support the
conclusion that a large fraction of 1-deoxy-GlcNAlk is metabolized into the protein
acetylation pathway. We cannot definitively rule out the incorporation of our MCR into
other types of protein modifications, but believe that our data demonstrates the unique
utility of chemical synthesis to develop new MCRs that can be used to visualize cellular
metabolism in addition to their traditional roles as probes of posttranslational
modifications. Given the resurgent importance of cellular metabolism in human disease
(e.g., diabetes and cancer), we believe that these tools can provide important and exact
information on the transformation of metabolites to PTMs where they can directly effect
protein function.
283
Materials and Methods
All reagents used for chemical synthesis were purchased from Sigma-Aldrich, Alfa Aesar
or EMD Millipore unless otherwise specified and used without further purification. All
anhydrous reactions were performed under argon or nitrogen atmosphere. Analytical
thin-layer chromatography (TLC) was conducted on EMD Silica Gel 60 F
254
plates with
detection by ceric ammonium molybdate (CAM), anisaldehyde or UV . For flash
chromatography, 60 Å silica gel (EMD) was utilized. Electrospray (ESI) and
Atmospheric Pressure Chemical Ionization (APCI) was preformed on an Agilent LCTOF
(2006) by University of California Riverside Mass Spectrometry Facility.
1
H spectra were
obtained at 400, 500, or 600 MHz on a Varian spectrometers Mercury 400, VNMRS-500,
or -600. Chemical shifts are recorded in ppm (δ) relative to solvent.
13
C spectra were
obtained at 100, 125 or 150 MHz on the same instruments.
Chemical Synthesis.
Known compounds Ac
4GlcNAlk (Zaro et al., 2011b), azido-rhodamine (Charron et al.,
2009) and sodium 4-pentynoate (Yang et al., 2010) were synthesized according to
literature procedures.
284
Compound 5.1 3,4,6-Tri-O-Acetyl-1-Chloro-1-Deoxy-N-Acetylglucosamine
{Macmillan:2002te}. Commercially-available N-acetylglucosamine
(10.00 g, 45.21 mmol) under Argon atmosphere was stirred vigorously
in Acetyl Chloride (17.68 mL, 248.63 mmol) for 16 h at room
temperature. Upon completion, CH
2Cl 2 was added and the resulting
mixture was extracted with ice-water (1x), saturated NaHCO
3 (1x), H 2O (1x), and Brine
(1x). The organic layer was dried over Na
2SO 4, filtered and concentrated. The crude
mixture was then purified over silica-gel chromatography (33% CH
2Cl 2 in EtOAc) to
yield the purified product (6.33 g, 38% yield).
1
H NMR (500 MHz, CDCl 3) δ 6.18 (d, J =
3.7 Hz, 1H), 5.83 (d, J = 8.7 Hz, 1H), 5.35(t, J = 9.3 Hz, 1H), 5.23 (t, J = 9.5 Hz, 1H),
4.53 (ddd, J = 10.7, 8.7, 3.7 Hz, 1H), 4.32 – 4.23 (m, 2H), 4.13 (d, J = 10.4 Hz, 1H), 2.10
(d, J = 0.7 Hz, 3H), 2.05 (t, J = 1.0 Hz, 6H), 1.98 (d, J = 0.6 Hz, 3H).
Compound 5.2 3,4,6-Tri-O-Acetyl-1-Deoxy-N-acetylglucosamine (1-deoxy-Ac
3GlcNAc)
(Schäfer et al., 1998). 5.1(6.30 g, 17.22 mmol) was dissolved in toluene
(100 mL) and purged with Argon for 30 min. At this time, tributyltin
hydride (5.56 mL, 20.66 mmol) and azobisisobutyronitrile (560 mg, 3.44
mmol) were added and the reaction stirred at reflux (~110 ℃) for 1.5 h.
Upon completion, the reaction was concentrated and purified by silica gel
chromatography (50% CH2Cl2 in EtOAc for 2 column volumes, 25% CH2Cl2 for 1
volume, 100% EtOAc for 1 volume, 10% MeOH in EtOAc for 2 volumes) to afford the
product (5.58 g, 98% yield).
O
AcO
OAc
AcO
Cl
NH
O
O
AcO
OAc
AcO
NH
O
285
Compound 5.3 1-Deoxyglucosamine hydrochloride (Schäfer et al., 1998). 5.2 (2.50 g,
7.55 mmol) was dissolved in 2.5 M HCl (20 mL) and refluxed for 2
h. Upon completion the mixture was concentrated. The resulting oil
was dissolved in minimal EtOH and Et
2O was added under stirring
until the product precipitated and was filtered and washed with isopropanol to yield the
product as an off-white powder (1.14 g, 75% yield).
1
H NMR (500 MHz, D 2O) δ 3.99 –
3.80 (m, 3H), 3.72 (dd, J = 12.3, 5.8 Hz, 1H), 3.53 (dd, J = 10.1, 8.7 Hz, 1H), 3.43 (dd, J
= 9.8, 8.7 Hz, 1H), 3.39-3.35 (m, 1H), 3.27 (t, J = 11.2 Hz, 1H).
Compound 5.4 3,4,6-Tri-O-Acetyl-1-Deoxy-N-4-pentynylglucosamine
(1-deoxy-Ac
3GlcNAlk). 5.3 (250 mg, 1.25 mmol) was concentrated
from toulene 3x and resuspended in DMF under Argon. To the
starting material was added TEA (183 μL, 1.31 mmol) and the
reaction proceeded for 10 min. 4-Pentynoic acid (147 mg, 1.50
mmol) followed by N,N’-Dicyclohexylcarbodiimide (309 mg, 1.50 mmol) were then
added followed by a catalytic amount of DMAP (~5 mg). The starting material slowly
went in to solution and the reaction proceeded for 16 h. Upon completion, the reaction
mixture was concentrated under reduced pressure and purified by column
chromatography (10% MeOH in CH
2Cl 2). The purified product was then resuspended in
pyridine (5 mL) and acetic anhydride (1.5 mL) and allowed to stir for 16 h. Upon
completion the reaction was concentrated, resuspended in CH
2Cl 2, washed with 1M HCl
O
HO
OH
HO
NH
3
+
Cl
-
O
AcO
NH
O
OAc
AcO
286
(1x), saturated NaHCO 3 (1x), H 2O (1x), and Brine (1x). The organic layer was dried over
Na
2SO 4, filtered and concentrated. The crude mixture was then purified over silica-gel
chromatography 50-60% EtOAc in Hexanes to afford crude (154 mg, 33% yield over 2
steps).
1
H NMR (500 MHz, CDCl 3) δ 5.78 (d, J = 7.2 Hz, 1H), 5.07 (t, J = 9.6 Hz, 1H),
4.97 (t, J = 9.7 Hz, 1H), 4.26 – 4.16 (m, 3H), 4.13 (dd, J = 12.3, 2.4 Hz, 1H), 3.55 (ddd, J
= 9.8, 5.0, 2.4 Hz, 1H), 3.20 (t, J = 12.5 Hz, 1H), 2.48 (td, J = 7.1, 2.7 Hz, 2H), 2.34 (t, J
= 7.1 Hz, 2H), 2.09 (s, 2H), 2.06 (s, 2H), 2.04 (s, 3H), 1.98 (t, J = 2.7 Hz, 1H).
13
C NMR
(126 MHz, CDCl
3) δ 172.29, 171.24, 170.94, 169.52, 82.73, 76.84, 74.33, 69.66, 68.44,
68.33, 62.56, 50.80, 35.49, 21.05, 20.98, 20.84, 14.95. ESI-MS calculated for
C
17H 23NNaO 8 [M+Na]
+
392.13, found 392.00.
Compound 5.5 1-Deoxy-N-4-pentynylglucosamine
(1-deoxy-GlcNAlk). 5.4 (154 mg, 0.417 mmol) was dissolved in
MeOH (10 mL). Freshly prepared NaOMe (Na in MeOH) was added
dropwise to reach pH 9-10. The reaction was monitored TLC and
determined finished in 1 h. The reaction was quenched with dilute acetic acid in MeOH
until pH 7 reached. The mixture was then concentrated and column chromatograpy (10%
MeOH in CH
2Cl 2) yielded the pure product (85 mg, 84% yield).
1
H NMR (500 MHz,
CD
3OD) δ 3.91 (dd, J = 10.9, 5.2 Hz, 1H), 3.87 – 3.77 (m, 2H), 3.62 (dd, J = 11.9, 6.0
Hz, 1H), 3.38 (dd, J = 10.1, 8.6 Hz, 1H), 3.29 – 3.24 (m, 2H), 3.20 – 3.14 (m, 1H), 2.51 –
2.35 (m, 4H), 2.25 (t, J = 2.5 Hz, 1H).
13
C NMR (125 MHz, CD 3OD) δ 173.05, 82.12,
O
HO
NH
O
OH
HO
287
81.02, 75.49, 71.04, 68.87, 67.69, 61.73, 51.64, 34.67, 14.36. ESI-MS calculated for
C
11H 18NO 5 [M]
+
244.12, found 244.20.
Cell Culture. COS-7, HEK293, HeLa and MCF7 cells were cultured in high-glucose
DMEM media (Corning) enriched with 10% fetal bovine serum (HyClone,
ThermoScientific). NIH3T3 and MEF cells were cultured in high-glucose DMEM media
(Corning) enriched with 10% fetal calf serum (HyClone, ThermoScientific). H1299 cells
were cultured in RPMI media enriched with 10% fetal bovine serum (HyClone,
ThermoScientific). All cell lines were maintained in a humidified incubator at 37 °C and
5.0% CO
2.
Metabolic Labeling. To cells at 80-85% confluency, media containing
1-deoxy-Ac 3GlcNAlk, Ac 4GlcNAlk (1,000 x stock in DMSO) or 1-deoxy-GlcNAlk
(dissolved directly in media) or DMSO vehicle was added as indicated. For chase
experiments, media was replaced with media supplemented with 1-deoxy-Ac
3GlcNAc
(200 μM, 1,000 x stock in DMSO).
Preparation of Nonidet P-40 (NP-40)-Soluble Lysates. The cells were then collected by
trypsinization and pelleted by centrifugation at for 4 min at 500 x g, followed by washing
2x with PBS (1 mL). Cell pellets were then resuspended in 100 μl of 1% NP-40 lysis
buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with
Complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Roche Biosciences) for
288
20 min and then centrifuged for 10 min at 10,000 x g at 4 °C. The supernatant (soluble
cell lysate) was collected and the protein concentration was determined by BCA assay
(Pierce, ThermoScientific).
Preparation of 4% SDS-soluble lysates. Cells were collected by trypsinization and
pelleted by centrifugation for 4 min at 500 x g, followed by washing 2x with PBS (1 mL).
Cell pellets were then resuspended in 75 μL 0.05% SDS buffer (0.05% SDS, 10 mM TEA
pH 7.4, 150 mM MgCl
2) with Complete Mini protease inhibitor cocktail (Roche
Biosciences). To this was added 1 μL Benzonase (Sigma), and the cells were incubated
on ice for 30 min. Then, 200 μL 4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA
pH 7.4) was added, and the cells were briefly sonicated in a bath sonicator followed by
centrifugation (20,000 x g for 10 min at 15 °C). Soluble protein concentration was
normalized by BCA assay (Pierce, ThermoScientific) to 1 mg/mL.
Cu(I)-Catalyzed [3 + 2] Azide-Alkyne Cycloaddition. Cell lysate (200 μg) was diluted
with cold 1% NP-40 lysis buffer to obtain a desired concentration of 1 μg/μL. Newly
made click chemistry cocktail (12 μL) was added to each sample [azido- or
alkynyl-rhodamine tag (100 μM, 10 mM stock solution in DMSO);
tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (1 mM, 50 mM freshly prepared
stock solution in water); tris[(1-benzyl-1-H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (100
μM, 10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared
stock solution in water) for a total reaction volume of 200 μL. The reaction was gently
289
vortexed and allowed to sit at room temperature for 1 h. Upon completion, 1 mL of ice
cold methanol was added to the reaction, and it was placed at -20 °C for 2 h to precipitate
proteins. The reactions were then centrifuged at 10,000 x g for 10 min at 4 . The
supernatant was removed, the pellet was allowed to air dry for 15 min, and then 50 μL
4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) was added to each
sample. The mixture was sonicated in a bath sonicator to ensure complete dissolution,
and 50 μL of 2x SDS-free loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4%
β-mercaptoethanol, pH 6.8) was then added. The samples were boiled for 5 min at 97 °C,
and 40 μg of protein was then loaded per lane for SDS-PAGE separation (Any Kd,
Criterion Gel, Bio-Rad).
In-Gel Fluorescence Scanning. Following SDS-PAGE separation, gels were scanned on
a Typhoon 9400 Variable Mode Imager (GE Healthcare) using a 532 nm for excitation
and 30 nm bandpass filter centered at 610 nm for detection.
Metabolic labelling comparison of Sodium Pentynoate and 1-deoxy-Ac
3GlcNAlk.
MEFs were treated with 0.2 mM, 5 mM or DMSO vehicle for 6 h. Upon treatment
completion, cells were washed, trypsinized and pelleted. Cell pellets were washed 3x
with PBS, lysed with 1% NP-40 buffer and soluble lysate subjected CuAAC with
azido-rhodamine as described in the methods section above.
290
Competition of 1-deoxy-Ac 3GlcNAlk labelling with NaOAc. NIH-3T3 cells were treated
with or without NaOAc (10 mM) and 1-deoxy-Ac3GlcNAlk (200 μM) for 6 h. Cells were
then lysed by 4% SDS buffer with benzonase as outlined in the methods section above .
Inhibition of histone acetyl transferases by curcumin. NIH-3T3 cells were treated with
or without curcumin (60 μM) for 30 min prior to treatment with 1-deoxy-Ac
3GlcNAlk
(200 μM) for 5.5 h. Cells were then lysed by 4% SDS buffer with benzonase as outlined
in the methods section above.
MTS Assay. MEFs (10
4
) plated 4 h prior in a 96-well plate were treated for 24 or 48 h
with either 1-deoxy-Ac3GlcNAlk (200 µM) or DMSO vehicle in triplicate. Upon
treatment completion, cell viability was determined by MTS assay according to literature
procedure (CellTiter 96 AQeous Non-Radioactive Cell Proliferation Assay, Promega,
Madison, WI) with the following minor change. After sufficient color development, a
reaction-quenching formazan-solubilization solution (100 µL) was added to each well
(50% DMF, 20% SDS in H
2O). Absorbance at 490 nm was read using a BioTek Synergy
H4 Multi-Mode Microplate reader.
Biotin enrichment and On-bead trypsinolysis. NIH-3T3 cell pellets labeled with
1-deoxy-Ac
3GlcNAlk, Ac 4GlcNAlk or 1-deoxy-Ac 3GlcNAc for 16 hours were
resuspended in 200 μL H2O, 60 μL PMSF in H2O (250 mM), and 500 μL 0.05% SDS
buffer (0.05% SDS, 10 mM TEA pH 7.4, 150 mM MgCl
2) with Complete Mini protease
291
inhibitor cocktail (Roche Biosciences). To this was added 8 μL Benzonase (Sigma), and
the cells were incubated on ice for 30 min. Then, 2000 μL 4% SDS buffer (4% SDS, 150
mM NaCl, 50 mM TEA pH 7.4) was added, and the cells were briefly sonicated in a bath
sonicator followed by centrifugation (20,000 x g for 10 min at 15 °C). Soluble protein
concentration was normalized by BCA assay (Pierce, ThermoScientific) to 1 mg/mL, and
10 mg of total protein was subjected to the appropriate amount of click chemistry cocktail
containing azido-PEG
3-biotin (5 mM, Click Chemistry Tools) for 1 h, after which time 10
volumes of ice-cold MeOH were added. Precipitation proceeded 2 hours at -20 °C.
Precipitated proteins were centrifuged at 5,200 x g for 30 min at 0 °C and washed 3 times
with 40 mL ice-cold MeOH, with resuspension of the pellet each time. The pellet was
then air-dried for 1 h. To capture the biotinylated proteins by streptavidin beads, the
air-dried protein pellet was resuspended in 2 mL of resuspension buffer (6 M urea, 2 M
thiourea, 10 mM HEPES pH 8.0) by bath sonication. To cap cysteine residues, 100 μl of
freshly-made TCEP (200 mM stock solution, Thermo) was then added and the mixture
incubated for 30 min, followed by 40 μl of freshly prepared iodoacetamide (1 M stock
solution, Sigma) and incubation for a further 30 min in the dark. Steptavadin beads (250
μL of a 50% slurry per sample, Thermo) were washed 2x with 1 mL PBS and 1x with 1
mL resuspension buffer and resuspended in resuspension buffer (200 μL). Each sample
was combined with streptavidin beads and incubated on a rotator for 2 h. These mixtures
were then transferred to Mini Bio-Spin® columns (Bio-Rad) and placed on a vacuum
manifold. Captured proteins were then washed with agitation 5x with resuspension buffer
(10 mL), 5x PBS (10 mL), 5x with 1% SDS in PBS (10 mL), 30x with PBS (1 mL per
292
wash, vacuum applied between each wash), and 5x 2M urea in PBS (1 mL per wash,
vacuum applied between each wash). Beads were then resuspended in 2 M urea in PBS (1
mL), transferred to screw-top tubes, and pelleted by centrifugation (2000 x g for 2 min).
At this time, 800 μL of the supernatant was removed, leaving a volume of 200 μL. To this
bead-mixture was added 2 μL of CaCl
2 (200 mM stock, 1 mM final concentration) and 2
μL of 1 mg/mL sequence grade trypsin (Promega) and incubated at 37 °C for 18 hours.
The resulting mixtures of tryptic peptides and beads were transferred to Mini Bio-Spin®
columns (Bio-Rad) and the eluent was collected by centrifugation (1,000 x g for 2 min).
Any remaining peptides were eluted by addition of 100 μL of 2 M urea in PBS followed
by centrifugation as immediately above. The tryptic peptides were then applied to C
18

spin columns (Pierce) according to manufacturer's instructions, eluted with 70%
acetonitrile in H
2O, and concentrated to dryness on a speedvac.
LC-MS Analysis. Peptides were desalted on a trap column following separation on a
12cm/75μm reversed phase C18 column (Nikkyo Technos Co., Ltd. Japan). A 3 hour
gradient increasing from 10% B to% 45% B in 3 hours (A: 0.1% Formic Acid, B:
Acetonitrile/0.1% Formic Acid) was delivered at 150 nL/min. The liquid chromatography
setup (Dionex, Boston, MA, USA) was connected to an Orbitrap XL (Thermo, San Jose,
CA, USA) operated in top-5- mode. Acquired tandem MS spectra (CID) were extracted
using ProteomeDiscoverer v. 1.3 (Thermo, Bremen, Germany) and queried against the
human Uniprot protein database using MASCOT 2.3.02 (Matrixscience, London, UK).
Peptides fulfilling a Percolator calculated 1% false discovery rate threshold were
293
reported. All LC-MS/MS analysis were carried out at the Proteomics Resource Center at
The Rockefeller University, New York, NY , USA.
Acid extraction of histones. NIH-3T3 cells were treated with 1-deoxy-Ac 3GlcNAlk (200
μM) or 1-deoxy-Ac
3GlcNAc (200 μM) for 16 hr. Cells were collected by trypsinization
and pelleted by centrifugation at 4 ℃ for 2 min at 2,000 x g, followed by washing with
PBS (1 mL) two times. Cell pellets were then resuspended in ice-cold hypotonic lysis
buffer [10 mM triethanolamine (TEA), 1 mM KCl, 1.5 mM MgCl
2, 1 mM PMSF, pH 7.4
with Complete Mini protease inhibitor cocktail (Roche Biosciences)]. The resuspended
cells were homogenized by Dounce homogenizer and lysed in 3 cycles of freeze-thaw.
Intact nuclei were pelleted at 4 ℃ for 10 min at 10,000 x g and washed 2x with ice-cold
hypotonic lysis buffer. The nuclear pellet was resuspended in 0.4 N H
2SO 4 and agitated
overnight on a rotator at 4 ℃. Nuclear debris was pelleted at 4 ℃ for 10 min at 16,000 x
g and the supernatant containing histones was collected and precipitated in ice-cold
MeOH in -80 ℃ overnight. Precipitated histones were collected at 4 ℃ for 10 min at
16,000 x g and washed 2x with ice-cold MeOH. The resulting protein pellet was air dried
and resuspended in water. Concentration was determined by BCA Assay and normalized
with 1% NP-40 lysis buffer [1% NP-40, 150 mM NaCl, 50 mM triethanolamine (TEA)
pH 7.4] with Complete Mini protease inhibitor cocktail] for CuAAC.
294
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298
Chapter 6. The small molecule 2-Azido-2-deoxy-glucose is a metabolic
chemical reporter of O-GlcNAc modifications in mammalian cells,
revealing an unexpected promiscuity of O-GlcNAc transferase.
**
Introduction
Metabolic chemical reporters (MCRs) of glycans are typically monosaccharide analogs
that contain bioorthogonal functional groups, such as an alkyne or azide.(Chuh and Pratt,
2015b; Chuh et al., 2016; Grammel and Hang, 2013) Once living cells are treated with
one of these analogs, the MCRs can be metabolically transformed into at least one, if not
more, high-energy sugar donors that can be subsequently utilized by glycosyltransferases
resulting in the incorporation of MCRs into glycans. An ever growing number of
bioorthogonal reactions can then be employed for the selective installation of tags for the
visualization, characterization, and identification of glycoconjugates, most notably
glycoproteins.(Patterson et al., 2014b) In the past five years, we and others have
demonstrated that the majority of glycoprotein MCRs are not specific for one type of
glycosylation.(Bateman et al., 2013a; Boyce et al., 2011; Chuh et al., 2014; Zaro et al.,
2011b) For example, the original MCR for intracellular O-GlcNAc modification,
N-azidoacetyl-glucosamine (GlcNAz),(V ocadlo et al., 2003) labels both O-GlcNAcylated
proteins and cell-surface glycans (i.e., N-linked and mucin O-linked).(Chuh et al., 2014;
**
Balyn W. Zaro, Anna R. Batt, and Marisol X. Navarro(University of Southern California) contributed to
the work presented in this chapter.
299
Zaro et al., 2011b) Additionally, we have shown that MCRs that have their bioorthogonal
functionality at the N-acetyl position, which makes up the vast majority of glycan
reporters, have the potential to be de-N-acetylated, resulting in labeling of the protein
acetylation pathway.(Zaro et al., 2014) These issues result in a lack of specificity that can
limit the information that can be gained in glycoprotein visualization experiments or the
number of a specific type of glycoprotein that can be identified using enrichment and
proteomics.
We recently demonstrated that changes to the MCR structure resulted in the first selective
reporter of O-GlcNAcylation, 6-azido-6-deoxy-N-acetyl-glucosamine
(6AzGlcNAc).(Chuh et al., 2014) O-GlcNAcylation is the addition of the
monosaccharide N-acetyl-glucosamine to serine and threonine side-chains of proteins
located in the cytosol, nucleus, and mitochondria.(Holt and Hart, 1986; Torres and Hart,
1984; Zachara and Hart, 2002) O-GlcNAc transferase (OGT) uses the uridine
diphosphate sugar donor UDP-GlcNAc to add O-GlcNAc to protein substrates, and these
modifications can be subsequently removed by O-GlcNAcase (OGA), rendering
O-GlcNAcylation dynamic.(V ocadlo, 2012) Genetic knockout of either of these enzymes
is lethal in mammals and insects,(O'Donnell et al., 2004; Shafi et al., 2000; Sinclair et al.,
2009; Yang et al., 2012) and the modification plays key roles in human disease, including
cancer and neurodegeneration.(Ma and V osseller, 2013; Yuzwa and V ocadlo, 2014) Here,
we demonstrate that 2-azido-2-deoxy-glucose (2AzGlc) is also a selective MCR for
O-GlcNAc modifications. Treatment of cells with the per-acetylated reporter Ac
42AzGlc
300
(Figure 6-1A) resulted in the labeling of a variety of proteins by 2AzGlc in mammalian
cells but no detectable incorporation of 2AzGlc into cell-surface glycoproteins.
Additionally, bioorthogonal enrichment of proteins that were labeled with Ac
42AzGlc
followed by proteomics identified many known O-GlcNAcylated proteins. In contrast to
6AzGlcNAc, 2AzGlc appears to not be removed from substrate proteins by OGA,
consistent with the enzymatic mechanism that requires anchimeric assistance of the
N-acetyl group.(Macauley, 2005) Additionally, treatment of cells with Ac
42AzGlc at a
high concentration (200 μM) was toxic to cells after overnight exposure. Interestingly,
this toxicity is inherent to the structure of the small-molecule, as deprotection of the
6-O-acetate to yield Ac
32AzGlc removed all cell-death but retained similar levels of
protein labeling. Importantly, this observation has been made on other MCRs by the
Yarema lab, which they attribute to changing the balance between the productive
biosynthetic transformation of reporters to their corresponding donor sugars versus toxic
side-reactions.{Aich:2008fe}
Results
Given our interesting previous results that structural changes in the chemical structure of
MCRs can change their selectivity, we set out to explore 2AzGlc, as it lacks the
acetamido-functionality that is common to the vast majority of monosaccharide positions
found in glycoproteins and that has been a consistent feature of all previous MCRs.
Therefore, we rationalized that it may have interesting properties concerning its
metabolism and glycoprotein selectivity. Towards this goal, we prepared the
301
per-O-acetylated compound, Ac 42AzGlc, in two steps from glucosamine. These acetates
enable diffusion of the reporter through cellular membranes where they are removed by
lipases and/or hydrolases. NIH3T3 and H1299 cells were then treated with increasing
concentrations of Ac
42AzGlc for 6 h, and the corresponding cell lysates were subjected to
the copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC) with an alkyne-rhodamine
tag. In-gel fluorescence scanning showed a range of labeled proteins (Figure 6-1B). In
both cell lines, we observed a large increase in labeling when the concentration of
Ac
42AzGlc was increased from 150 to 200 μM. Notably, this higher concentration also
resulted in toxicity when the cells were treated for longer periods of time (i.e., 16 h). This
observation is consistent with previous experiments by the Bertozzi lab, who commented
on the toxicity of Ac
42AzGlc but did not characterize its labeling.(Saxon et al., 2002) To
explore this toxicity further, the same cell lines were treated with a range of Ac
42AzGlc
concentrations for 16 h and their NAD(P)H-dependent reductase activity was measured
using a commercially available MTS assay (Figure 6-1C). Interestingly, an increase in
activity was observed for lower concentrations of Ac
42AzGlc, indicating additional
metabolic flux into reductive equivalents. However, the cells displayed a dramatic loss of
activity between 150 to 200 μM Ac
42AzGlc, consistent with our morphological
observations.
302
Figure 6-1. Evaluation of per-acetylated-2-azido-glucose (Ac 42AzGlc) as a metabolic
chemical reporter (MCR). (A) Treatment of living cells with MCRs, like Ac42AzGlc, results in
modification of proteins that can then be visualized using bioorthogonal reactions. (B)
Ac42AzGlc-treatment results in labeling of a wide range of proteins. NIH3T3 or H1299 cells
were treated with the indicated concentrations of Ac42AzGlc for 6 h, followed by CuAAC with
alkyne-rhodamine and visualization of labeled proteins by in-gel fluorescence scanning. (C) High
concentrations of Ac42AzGlc are toxic to mammalian cells. NIH3T3 or H1299 cells were treated
with the indicated concentrations of Ac42AzGlc for 16 h and their viability was then measured
using an MTS assay. Quantitated data is the average of three separate biological experiments, and
error bars represent ±s.e.m. from the mean of biological replicates (n = 3).
303
Despite this toxicity, we next set out to characterize the selectivity of Ac 42AzGlc as an
MCR. To determine if 2AzGlc is incorporated into cell-surface glycans, H1299 cells were
treated with Ac
42AzGlc (200 μM) for 6 h, which is before the onset of obvious cellular
toxicity. The intact cells were then subjected to a strain-promoted azide-alkyne
cycloaddition (SPAAC) with DBCO-biotin, which can only label extracellular azides,
followed by incubation with FITC-avidin and analysis by flow-cytometry (Figure 6-2A).
As controls, we also simultaneously treated cells with either Ac
4GalNAz (200 μM),
which does label cell-surface glycoproteins,(Chuh et al., 2014; Hang et al., 2003) or
Ac
36AzGlcNAc (200 μM), which does not.(Chuh et al., 2014) We observed essentially
no fluorescence signal from cells treated with either Ac
42AzGlc or Ac 36AzGlcNAc,
despite strong signal from the Ac
4GalNAz-treated cells, indicating that Ac 42AzGlc does
not label cell-surface glycans. We next used Ac
42AzGlc in an unbiased proteomics
experiment in NIH3T3 cells, as we have utilized these cells in the past for the
identification of labeled proteins with other MCRs. Specifically, NIH3T3 cells were
treated in triplicate with Ac
42AzGlc or Ac 4GlcNAc as a control (both at 200 μM) for 6 h,
again before any signs of toxicity. The cells were then lysed under denaturing conditions
(4% SDS) and the labeled proteins underwent CuAAC with an alkyne biotin-tag. The
labeled proteomes were then reduced and alkylated with iodoacetamide, followed by
incubation with streptavidin-conjugated beads. After extensive washing, the enriched
proteins were subjected to on-bead trypsinolysis, and the resulting peptides were
analyzed using LC-MS/MS and identified using Proteome Discoverer and Mascot.
Spectral counting was used to quantify the proteins enriched in either sample. Proteins
304
were considered to have been labeled by 2AzGlc if they met certain criteria. First, the
proteins must have been identified by at least 1 peptide in each of the three
Ac
42AzGlc-treated samples. Second, the sum of the spectral counts from the
Ac
42AzGlc-treated samples must be three-times greater than the sum of the spectral
counts for the same protein in the Ac
4GlcNAc-treated sample. Finally, the spectral counts
in the MCR-treated samples must be statistically significantly different from the control
samples (P < 0.05, t-test). With these requirements, we identified 361 proteins (Figure
6-2B). Importantly, in this list were 265 proteins that had been previously identified in
other proteomics experiments as being potentially O-GlcNAcylated. We then annotated
these proteins based on their subcellular localization, since O-GlcNAcylated proteins will
be intracellular while cell-surface glycoproteins will be extra-cellular/lumenal or have a
transmembrane domain. Using the Uniprot database, we found that Ac
42AzGlc labeled
essentially exclusively intracellular or transmembrane proteins and only resulted in the
enrichment of one exclusively extracellular protein, EGF-containing fibulin-like
extracellular matrix protein, that could not bear O-GlcNAcylation. Next, we used
retroviral transformation to generate NIH3T3 cells that stably overexpress an HA-tagged
OGT or were transduced with an empty plasmid as a negative control. These cells were
treated with Ac
42AzGlc (200 μM) for 6 h, followed by CuAAC with alkyne-rhodamine
and analysis using in-gel fluorescence (Figure 6-2C). Notably, we observed significantly
more labeling in the OGT overexpressing cells, further indicating that 2AzGlc is an MCR
for O-GlcNAcylation. To explore the generality of 2AzGlc labeling, we next treated a
variety of mammalian cell-lines with Ac
42AzGlc (200 μM) for 6 h. In-gel fluorescence
305
scanning, after CuAAC with alkyne-rhodamine, showed strong labeling in all of the
cell-lines tested (Figure 6-3), indicating that 2AzGlc is a general MCR for the
visualization and identification of O-GlcNAcylated proteins.
Figure 6-2. Ac 42AzGlc is an O-GlcNAc metabolic chemical reporter. (A) Cell surface
glycoproteins are not labeled after treatment with Ac42AzGlc. H1299 cells were treated with the
indicated metabolic chemical reporters (200 μM) for 6 h at which time the cells were collected
and subjected to copper-free click chemistry with DBCO-biotin (Click Chemistry Tools). After
incubation with FITC-avidin, surface glycoprotein labeling was analyzed by flow cytometry. (B)
Unbiased identification of O-GlcNAcylated proteins using 2AzGlc. NIH3T3 cells were treated in
triplicate with Ac42AzGlc (200 μM) or Ac 4GlcNAc (200 μM) vehicle for 6 h, followed by
CuAAC with alkyne-biotin, enrichment with streptavidin-coated beads, and on-bead
trypsinolysis. Proteins that were identified using LC-MS/MS are represented as the total number
of positive spectral counts minus the total number of control spectral counts. Representative
known O-GlcNAcylated proteins are shown. (C) O-GlcNAc transferase (OGT) overexpression
increases 2AzGlc labeling. NIH3T3 cells that stably express either HA-tagged OGT or an empty
plasmid were treated with Ac42AzGlc (200 μM) or DMSO vehicle for 6 h. At this time, the cell
lysates were subjected to CuAAC with alkyne-rhodamine and labeled proteins were visualized
using in-gel fluorescence.
We next explored the reversibility of 2AzGlc labeling. As stated in the introduction, the
enzymatic mechanism of OGA involves anchimeric assistance of the N-acetyl group to
generate an oxazoline intermediate. A wealth of carbohydrate chemistry has demonstrated
that a 2-azido group is incapable of participating in this fashion,{Kerns:2012fy} raising
306
the likely possibility that once 2AzGlc is installed by OGT it cannot be removed by
OGA. To directly test this hypothesis, we performed a pulse-chase experiment. Briefly,
HeLa cells were treated with Ac
42AzGlc (200 μM) for 6 h. At this time the media was
exchanged for fresh media containing Ac
4GlcNAc (200 μM) and either the OGA inhibitor
Thiamet-G (10 μM)(Yuzwa et al., 2008) or DMSO vehicle. After an additional 12 h, the
corresponding cell-lysates were subjected to CuAAC with alkyne-rhodamine and analysis
by in-gel fluorescence (Figure 6-4A). In contrast to 6AzGlcNAc labeling that we
previously demonstrated is maintained by Thiamet-G treatment,(Chuh et al., 2014) we
observed no difference in the reduction of 2AzGlc labeling, indicating that it is not a
substrate for OGA. Interestingly, we did, however, observe a relatively rapid loss in
2AzGlc labeling over 12 h, suggesting that the labeled proteins were being turned over by
another mechanism. Proteasomal degradation is one of the major pathways by which
proteins are targeted for degradation. To determine if 2AzGlc-labeled proteins are being
destroyed by this pathway, we performed another pulse-chase experiment. In this case,
the pulse was identical to the one above, but in the chase, we added either the proteasome
inhibitor MG-132 (10 μM) or DMSO control. In-gel fluorescence scanning following
CuAAC with alkyne-rhodamine only showed a slight stabilization of the fluorescent
signal (Figure 6-4B), indicating that the majority of the 2AzGlc-dependent labeling is
being removed by an unknown mechanism.
307
Figure 6-3. Ac 42AzGlc is a general chemical reporter in a variety of mammalian cell lines.
(A) The indicated cell lines were treated with Ac42AzGlc (200 μM) or DMSO vehicle for 6 h.
Total cell lysates were then collected and subjected to CuAAC with alkyne-rhodamine, and
labeled proteins were visualized using in-gel fluorescence scanning.
Figure 6-4. Characterization of the dynamics of 2AzGlc labeling. (A) Protein labeling by
2AzGlc is not enzymatically removed by O-GlcNAcase (OGA). HeLa cells were treated with 200
μM Ac42AzGlc for 6 h. At this time, the media was exchanged for fresh media containing 200
μM Ac4GlcNAc and either the OGA inhibitor Thiamet-G (10 μM) or DMSO. Cells were
collected at the indicated times, subjected to CuAAC with alkyne-rhodamine and analyzed by
in-gel fluorescence scanning. (B) Labeling by 2AzGlc is not notably stabilized by proteasome
inhibition. HeLa cells were treated with 200 μM Ac42AzGlc for 6 h. At this time, the media was
exchanged for fresh media containing 200 μM Ac4GlcNAc and either the proteasome inhibitor
MG132 (10 μM) or DMSO. Cells were collected at the indicated times, subjected to CuAAC with
alkyne-rhodamine and analyzed by in-gel fluorescence scanning.
308
Finally, to explore the potential mechanism underlying the toxicity of
Ac
42AzGlc-treatment, we looked to the elegant experiments performed by the Yarema
lab who were exploring the off-target effects of monosaccharides modified by short-chain
fatty acids.{Aich:2008fe} Although the exact biological mechanism behind the toxicity
of certain acetylated monosaccharides are not entirely known, the authors demonstrated
that the toxicity of these molecules could be eliminated by revealing the 6-hydroxyl
functionality, which increases the flux of the carbohydrate into cellular biosynthetic
pathways. To test whether this observation could explain the toxicity of Ac
42AzGlc, we
synthesized the 6-OH analog (Ac
32AzGlc, Figure 6-5A and Scheme 6-1). First, we
examined whether aqueous conditions would result in migration of the 4-O-acetate to the
6-hydroxyl group, as can be observed under certain reaction conditions. To accomplish
this, we dissolved Ac
32AzGlc in a 5:1 mixture of D 2O and deuterated DMSO and
incubated the mixture for 16 h at room temperature. After this length of time the solution
was diluted to 1:1 D
2O and deuterated DMSO and analyzed by proton NMR. Comparison
of this spectra to a freshly prepared sample in the same solvent composition showed
essentially no differences, demonstrating that any acetate migration is minimal and below
the detection limit of NMR. Next, treatment with either NIH3T3 or H1299 cells with
Ac
32AzGlc resulted in robust labeling of proteins, as visualized by in-gel fluorescence
(Figure 6-5B). However, unlike Ac
32AzGlc, the 6-OH analog showed no cellular toxicity
at all of the concentrations tested (Figure 6-5C). These results support an inherent toxicity
mechanism that is largely unrelated to protein labeling by 2AzGlc as an O-GlcNAc
reporter.
309
Scheme 6-1. Synthesis of Ac 32AzGlc. (a) Imidazole, TBSCl, DMF 16 h, rt. (b) Acetic anhydride,
pyridine, 16 h, rt. (c) Acetic acid, 1M TBAF, THF 0 ℃ to rt over 16 h.
Figure 6-5. The acetylation pattern and labeling by 2AzGlc is responsible for cellular
toxicity. (A) Proteins can also be labeled by Ac32AzGlc, which bears a free hydroxyl at the 6-OH
position. (B) Ac32AzGlc-treatment results in labeling of a wide range of proteins. NIH3T3 or
H1299 cells were treated with the indicated concentrations of Ac32AzGlc for 16 h, followed by
CuAAC with alkyne-rhodamine and visualization of labeled proteins by in-gel fluorescence
scanning. (C) No concentrations of Ac32AzGlc are toxic to mammalian cells. NIH3T3 or H1299
cells were treated with the indicated concentrations of Ac32AzGlc for 16 h and their viability was
then measured using an MTS assay. Quantitated data is the average of four separate biological
experiments, and error bars represent ±s.e.m. from the mean of biological replicates (n = 4).
310
Discussion
The development of new MCRs for glycans has continued to grow the chemical toolbox
for the investigation of glycobiology. In particular, these reporters have been key for the
visualization and identification of glycoproteins. We have been focused on understanding
the selectivity of different MCRs, including some that would not be predicted to label
proteins based on the previously established biosynthetic and salvage pathways.
Continuing that theme here, we performed the cellular characterization of 2AzGlc as a
chemical reporter of glycosylation. Using bioorthogonal chemistry, we show that 2AzGlc
labels a range of proteins in total cell-lysate (Figure 6-1B and Figure 6-3), but does not
label cell-surface glycans at a detectable level (Figure 6-2A), suggesting that it might be
selective for O-GlcNAc modifications. This selectivity was further supported by an
unbiased proteomics experiment that used 2AzGlc to enrich a total of 361 proteins,
73.4% of which had been previously identified in other O-GlcNAcylation proteomics
experiments and only 1 exclusively extracellular or lumenal glycoprotein (Figure 6-2C).
The ability for 2AzGlc to be transferred by OGT from the corresponding UDP-2AzGlc
sugar donor in cells is somewhat surprising, given the ternary crystal structures of OGT
in complex with its peptide and donor-sugar substrates, as well as the structure with the
corresponding products.(Lazarus et al., 2012) These structures show that the 2-acetamide
group of GlcNAc undergoes significant movement during the enzymatic reaction and that
the amide nitrogen contributes a hydrogen-bond to the phosphate of the UDP
leaving-group. Additionally, the same authors show that in vitro OGT will accept
UDP-GalNAc but not UDP-Glc or UDP-2-keto-Glc.(Lazarus et al., 2012) However,
311
another structural and biochemical study found that OGT accepts UDP-GlcNAcF 3
(2-N-trifluoroacetamide), which they interpreted as evidence that the electronics of the
2-acetamide are not key for catalysis.(Schimpl et al., 2012) Given these data, the 2-azido
group might either be functionally benign or could participate in weak hydrogen bonds,
given its unique electronics.{Tchertanov:1999gq} We also confirmed a statement from
the Bertozzi lab that 2AzGlc is toxic to mammalian cells (Figure 6-1B), although they did
not show any supporting data.(Saxon et al., 2002) Here, we demonstrate that this toxicity
can be eliminated by deprotection of the 6-position to reveal the hydroxyl group, which
the Yarema lab has shown increased flux of other monosaccharide analogs into the
corresponding biosynthetic pathways. Given the mechanism of OGA, we also explored
the reversibility of the 2AzGlc modifications. We used a pulse-chase experiment to
demonstrate that 2AzGlc is not removed from proteins by OGA (Figure 6-4A). As stated
above, this is not necessarily surprising, given the requirement for anchimeric assistance
in the enzymatic mechanism.(Macauley, 2005) Interestingly, although loss of 2AzGlc
labeling is not OGA-dependent, the rate of loss is similar to other MCRs that are removed
by OGA, such as 6AzGlcNAc,(Chuh et al., 2014) suggesting to us that the
2AzGlc-labeled proteins might be degraded. However, a pulse-chase experiment with the
proteasomal inhibitor MG-132 only showed a modest stabilization of the fluorescent
signal (Figure 6-4B), indicating that another pathway exists to remove these
modifications. Further experiments will be needed to directly test this hypothesis. Our
proteomics data show that the vast majority of identified proteins have been previously
characterized as being potentially O-GlcNAcylated by a variety of chemical and
312
biological techniques. This indicates that the specificity for protein substrates between
2AzGlc and other MCRs is similar. However, we can not rule out that different proteins
are more readily modified by OGT with a reporter that lacks the 2-acetamido
functionality versus an MCR like 6AzGlcNAc. Therefore, we suggest that 6AzGlcNAc
be used for studies aimed at the biology of O-GlcNAcylation and that all
potentially-modified proteins be confirmed by either antibodies or the chemoenzymatic
modification strategy developed by the Hsieh-Wilson lab.(Clark et al., 2008) In summary,
2AzGlc is the second MCR that has been characterized to be selective for
O-GlcNAcylation. It also further highlights the promiscuity of OGT, which now includes
2AzGlc, as well as several N-acetyl-glucosamine monosaccharides,(Bateman et al.,
2013a; Boyce et al., 2011; Chuh et al., 2014; Li et al., 2016; V ocadlo et al., 2003; Zaro et
al., 2011b) and could have interesting implications for the modification of intracellular
proteins by natural, non N-acetyl-glucosamine monosaccharides that can be
biosynthetically transformed to the corresponding UDP-sugar donors.
Materials and Methods
Chemical Synthesis.
Known compounds Thiamet-G, Ac
4GalNAz, Ac 4GlcNAz, Ac 42AzGlucose and
azido-rhodamine were synthesized according to literature procedures(Charron et al.,
2009; Chuh et al., 2014; Saxon, 2000; Yuzwa et al., 2008). All reagents used for chemical
synthesis were purchased from Sigma-Aldrich, Alfa Aesar or EMD Millipore unless
otherwise specified and used without further purification. All anhydrous reactions were
313
performed under argon or nitrogen atmosphere. Analytical thin-layer chromatography
(TLC) was conducted on EMD Silica Gel 60 F
254 plates with detection by ceric
ammonium molybdate (CAM), anisaldehyde or UV . For flash chromatography, 60 Å
silica gel (EMD) was utilized.
1
H spectra were obtained at 400, 500, or 600 MHz on a
Varian spectrometers Mercury 400, VNMRS-500, or -600. Chemical shifts are recorded
in ppm (δ) relative to solvent.
13
C spectra were obtained at 100, 125 or 150 MHz on the
same instruments.
Synthesis of 1,3,4,6-Tetra-O-Acetyl-2-Azido-2-Deoxyglucose (Ac
42AzGlc)
Compound 6.1 2-Azido-2-Deoxyglucose: Commercially available
glucosamine hydrochloride was subjected to azide formation using a
diazo transfer reaction according to literature
procedure.(Goddard-Borger and Stick, 2007)
1
H NMR (500 MHz, Deuterium Oxide) of
β-anomer δ 4.71 (d, J = 8.1 Hz, 1H), 3.92 – 3.82 (m, 1H), 3.81 – 3.70 (m, 1H), 3.54 –
3.43 (m, 3H), 3.28 (dd, J = 9.6, 8.2 Hz, 1H). The product was used in the subsequent
reaction without further characterization.
Compound 6.2 1,3,4,6-Tetra-O-Acetyl-2-Azido-2-Deoxyglucose
(Ac
42AzGlc, 2): Compound 6.1 was resuspended in pyridine and
acetic anhydride. A catalytic amount of dimethylaminopyridine was
added and reaction stirred at room temperature for 16 h. Upon completion reaction was
concentrated, and the crude was resuspended in dichloromethane. The organic layer was
O
HO
N
3
OH
HO
OH
O
AcO
N
3
OAc
AcO
OAc
314
washed with equivolume 1 M HCl (1x), saturated Sodium Bicarbonate (x2), Water (x2)
and brine (1x). The washed organic layer was then concentrated and subjected to column
chromatography to yield the pure product. NMR characterization of this known
compound was consistent with previous reports.(Saxon et al., 2002)
1
H NMR (500 MHz,
CDCl
3) of β-anomer δ 5.55 (d, J = 8.5 Hz, 1H), 5.15 – 5.00 (m, 2H), 4.34 – 4.25 (m, 1H),
4.11 – 4.05 (m, 1H), 3.84 – 3.77 (m, 1H), 3.71 – 3.62 (m, 1H), 2.19 (s, 3H), 2.09 (s, 3H),
2.07 (d, 3H), 2.02 (s, 3H).
Synthesis of 1,3,4-Tri-O-Acetyl-2-Azido-2-Deoxyglucose (Ac
32AzGlc)
Compound 6.3 6-Tert-Butyldimethylsilyl-2-Azido-2-Deoxyglucose
(6TBS2AzGlc): Compound 6.1 (668 mg, 3.26 mmol) was dissolved
in dry DMF (8 mL) under nitrogen at room temperature. Imidazole
(667 mg, 9.8 mmol) and tert-butyldimethylsilyl chloride (613 mg, 4.07 mmol) were
added and the reaction stirred for 16 h. The reaction was diluted in ethyl acetate and
washed with sodium bicarbonate (2x), water (2x), and brine (1x). The organic layer was
dried with sodium sulfate, filtered, and concentrated by vacuum. The compound was
purified by column chromatography (25% ethyl acetate:hexanes) to afford the product as
a light brown syrup (327 mg, 31%).
1
H NMR (400 MHz, CDCl 3) of α-, β-anomer
mixture 5.18 (d, J = 3.4 Hz, 1H), 4.55 (d, J = 8.0 Hz, 1H), 4.01 (dd, J = 14.0, 6.0 Hz, 1H),
3.95 (t, J = 9.5 Hz, 1H) 3.86-3.65 (m, 6H), 3.53-3.46 (m, 2H), 3.37 (t, J = 9.5 Hz 1H),
3.31-3.25 (m, 1H), 3.20-3.14 (m, 2H), 0.81 (s, 18H), 0.00 (s, 12H).
13
C NMR (125 MHz,
O
HO
N
3
OTBS
HO
OH
315
CDCl3): δ (ppm) 95.89, 91.59, 76.91, 74.47, 72.45, 71.18, 70.08, 66.33, 63.9, 62.99,
25.77, -5.80. ESI-MS calculated for C
12H 26N 3O 5Si [M + H]: 320.1636, found: 319.5972.
Compound 6.4 1,3,4-Tri-O-Acetyl-6-Tert-Butyldimethylsilyl-2-
Azido-2-Deoxyglucose (Ac 36TBS2AzGlc): Compound 6.3 (150 mg,
0.47 mmol) was dissolved in pyridine (2 mL) and cooled to 0°C.
Acetic anhydride (200 uL, 2.11 mmol) was added and the reaction warmed to room
temperature over 16 h. The reaction was diluted in DCM and washed with sodium
bicarbonate (2x), water (2x), and brine (1x). The organic layer was dried with sodium
sulfate, filtered, and concentrated by vacuum before purification by column
chromatography (10% ethyl acetate:hexanes) to afford the product as a light brown syrup
(170 mg, 81%).
1
H NMR (400 MHz, CDCl 3) of α-, β-anomer mixture: δ (ppm) 6.28 (d, J
= 3.6 Hz, 1H), 5.52 (d, J = 8.6 Hz, 1H), 5.46-5.40 (m, 1H), 5.14 (dd, J = 10.2, 9.4 Hz,
1H), 5.08-5.04 (m, 2H), 3.91-3.85 (m, 1H), 3.76-3.70 (m, 1H), 3.69-3.58 (m, 6H), 2.16
(d, J = 1.8 Hz, 6H), 2.09 (d, J = 4.7 Hz, 6H), 2.03-1.99 (m, 6H), 0.86 (s, 18H),
0.04--0.03, (m, 12H).
13
C NMR (125 MHz, CDCl 3): δ (ppm) 169.7, 169.14, 168.38,
92.23, 89.83, 74.95, 73.01, 72.17, 71.08, 68.01, 62.34, 61.53, 60.19, 25.51, 20.52, -5.72.
ESI-MS calculated for C
18H 31N 3O 8SiNa [M + Na]
+
: 468.1773, found: 468.1766.
Compound 6.5 1,3,4-Tri-O-Acetyl-2-Azido-2-Deoxyglucose
(Ac 32AzGlc): Compound 6.4 (170 mg, 0.38 mmol) was dissolved in
THF (2 mL) and cooled to 0°C. Glacial acetic acid (445 ul) was
O
AcO
N
3
OTBS
AcO
OAc
O
AcO
N
3
OH
AcO
OAc
316
added followed by addition of 1.0 M TBAF in THF (7.8 mL) and the reaction was
allowed to warm to room temperature over 16 hours. The solution was diluted with ethyl
acetate, washed with sodium bicarb (2x), water (2x), and brine (1x), and dried with
sodium sulfate. The resulting mixture was filtered, concentrated by vacuum, and purified
by column chromatography (40% ethyl acetate:hexanes) to afford product as a yellow
syrup (107 mg, 85%).
1
H NMR (400 MHz, CDCl 3) of α-, β-anomer mixture: δ (ppm)
6.27 (d, J = 3.7 Hz, 1H), 5.54 (d, J = 8.5 Hz, 1H), 5.32 (dd, J = 10.5, 9.3 Hz, 1H),
4.95-4.90 (m, 1H), 4.56-4.47 (m, 2H), 4.21 (dd, J = 12.5, 1.7 Hz, 1H), 4.21 (dd, 12.5, 2.3
Hz, 1H), 3.92-3.87 (m, 1H), 3.65-3.48 (m, 6H), 2.20-2.18 (m, 12H), 2.12 (s, 6H).
13
C
NMR (125 MHz, CDCl
3): δ (ppm) 171.28, 170.95, 168.34, 92.35, 89.91, 74.86, 72.73,
72.03, 68.27, 62.16, 59.92, 20.53. ESI-MS calculated for C
12H 18N 3O 8 [M + H]
+
:
332.1088, found: 332.1085.
Cell culture. NIH3T3 cells stably expressing either HA-tagged O-GlcNAc Transferase
(OGT) or an empty vector were generated using Amphopack293 retroviral packaging cell
lines according to manufacturer procedure (Clontech).
NIH3T3 and MEF cells were cultured in high glucose DMEM media (HyClone,
ThermoScientific) enriched with 10% fetal calf serum (FCS, HyClone,
ThermoScientific). MDA-MB-231, MDA-MB-468, HeLa and MCF-7 were cultured in
high glucose DMEM (HyClone, ThermoScientific) enriched with 10% fetal bovine serum
(FBS HyClone, ThermoScientific). H1299 and HCT15 cells were cultured in RPMI 1640
medium (HyClone, ThermoScientific) enriched with 10% FBS. A549 cells were cultured
317
in F-12K medium (HyClone, ThermoScientific) enriched with 10% FBS. MCF10A and
SH-SY5Y were cultured in a 1:1 mixture of F-12 and DMEM media (HyClone,
ThermoScientific) enriched with 10% FBS.
All cell lines were maintained in a humidified incubator at 37 ℃ and 5.0% CO
2.
Metabolic labeling. To cells at 80-85% confluency, high- or low-glucose media
containing Ac 4GlcNAz, Ac 4GalNAz, Ac 42AzGlc (1,000 x stock in DMSO), or DMSO
vehicle was added as indicated. For chase experiments, low-glucose media was replaced
with high-glucose media supplemented with 200 µM Ac
4GlcNAc (Sigma) with or
without 20 µM Thiamet-G.
Preparation of NP-40-soluble lysates. The cells were collected by trypsinization and
pelleted by centrifugation at for 4 min at 2,000 x g, followed by washing 2x with PBS (1
mL). Cell pellets were then resuspended in 100 μL of 1% NP-40 lysis buffer [1% NP-40,
150 mM NaCl, 50 mM triethanolamine (TEA) pH 7.4] with Complete, Mini, EDTA-free
Protease Inhibitor Cocktail Tablets (Roche) for 20 min and then centrifuged for 10 min at
10,000 x g at 4 °C. The supernatant (soluble cell lysate) was collected and the protein
concentration was determined by BCA assay (Pierce, ThermoScientific).
Cu(I)-catalyzed [3 + 2] azide-alkyne cycloaddition (CuAAC). Cell lysate (200 μg) was
diluted with 1% NP-40 lysis buffer to obtain a desired concentration of 1 μg μL
−1
.
Newly-made click chemistry cocktail (12 μL) was added to each sample
318
[alkynyl-rhodamine tag (100 μM, 10 mM stock solution in DMSO);
tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (1 mM, 50 mM freshly prepared
stock solution in water); tris[(1-benzyl-1-H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (100
μM, 10 mM stock solution in DMSO); CuSO
4•5H 2O (1 mM, 50 mM freshly prepared
stock solution in water)] for a total reaction volume of 200 μL. The reaction was gently
vortexed and allowed to sit at room temperature for 1 h. Upon completion, 1 mL of ice
cold methanol was added to the reaction, and it was placed at -20 °C for 2 h to precipitate
proteins. The reactions were then centrifuged at 10,000 x g for 10 min at 4 °C. The
supernatant was removed, the pellet was allowed to air dry for 15 min, and then 50 μL
4% SDS buffer (4% SDS, 150 mM NaCl, 50 mM TEA pH 7.4) was added to each
sample. The mixture was sonicated in a bath sonicator to ensure complete dissolution,
and 50 μL of 2x SDS-free loading buffer (20% glycerol, 0.2% bromophenol blue, 1.4%
β-mercaptoethanol, pH 6.8) was then added. The samples were boiled for 5 min at 97 °C,
and 40 μg of protein was then loaded per lane for SDS-PAGE separation (Any Kd,
Criterion Gel, Bio-Rad).
In-gel Fluorescence Scanning. Following SDS-PAGE separation, gels were scanned on
a Typhoon 9400 Variable Mode Imager (GE Healthcare) using a 532 nm for excitation
and 30 nm bandpass filter centered at 610 nm for detection.
MTS Assay. NIH-3T3 and H1299 cells (1 x 10
4
cells) were plated per well in a 96-well,
white bottom dish 24 hours before treatment with 0-200 μM Ac
4GlcNAc or
319
Ac 32AzGlucose for 16 hours under low glucose media conditions (1 g/L). CellTiter 96
®

AQueous Non-Radioactive Cell Proliferation Assay (Promega, Madison, WI) was used
according to the provided protocol. Absorbance at 490 nm was read using a BioTek
Synergy H4 Multi-Mode Microplate reader.
Flow Cytometry of Cell-Surface Labeling with DBCO-Biotin. H1299 cells grown in
6-well plates at 80-85% confluency were treated with 200 μM Ac
4GlcNAc, Ac 4GlcNAz,
Ac
4GalNAz or Ac 32AzGlc in triplicate for 16 hours at which time media was removed
and cells were gently washed with PBS before being detached from the plate with 1 mM
EDTA in PBS. Cells were collected by centrifugation (5 min, 300 x g at 4 °C) and were
washed three times with PBS (5 min, 300 x g at 4 °C). Cells were then resuspended in
200 μL PBS containing DBCO-biotin (Click Chemistry Tools, 60 μM) for 1 h, after
which time they were washed three times with PBS (5 min, 300 x g at 4 °C) before being
resuspended in ice-cold PBS containing fluorescein isothiocynate (FITC) conjugated
avidin (Sigma, 5 μg/mL, 30 min at 4 °C). Cells were then washed three times in PBS (5
min, 300 x g at 4 °C) before being resuspended in 400 μL PBS for flow-cytometry
analysis. A total of 10,000 cells [dead cells were excluded by treatment with propidium
iodide (2.5 μg mL
−1
in water, 30 min)] were analyzed on a BD SORP LSRII Flow
Cytometer using the 488 nm argon laser.
Biotin Enrichment and On-bead Trypsinolysis. NIH3T3 or H1299 cell-pellets labeled
with Ac
36AzGlcNAc, Ac 32AzGlc or Ac 4GlcNAc for 16 hours were resuspended in 200
μL H
2O, 60 μL PMSF in H 2O (250 mM), and 500 μL 0.05% SDS buffer (0.05% SDS, 10
320
mM TEA pH 7.4, 150 mM NaCl) with Complete Mini protease inhibitor cocktail (Roche
Biosciences). To this was added 8 μL Benzonase (Sigma), and the cells were incubated
on ice for 30 min. Then, 4% SDS buffer (2000 μL) was added, and the cells were briefly
sonicated in a bath sonicator followed by centrifugation (20,000 x g for 10 min at 15 °C).
Soluble protein concentration was normalized by BCA assay (Pierce, ThermoScientific)
to 1 mg mL
−1
, and 10 mg of total protein was subjected to the appropriate amount of click
chemistry cocktail containing alkyne-PEG3-biotin (5 mM, Click Chemistry Tools) for 1
h, after which time 10 volumes of ice-cold MeOH were added. Precipitation proceeded 2
hours at -20 °C. Precipitated proteins were centrifuged at 5,200 x g for 30 min at 0 °C
and washed 3 times with 40 mL ice-cold MeOH, with resuspension of the pellet each
time. The pellet was then air-dried for 1 h. To capture the biotinylated proteins by
streptavidin beads, the air-dried protein pellet was resuspended in 2 mL of resuspension
buffer (6 M urea, 2 M thiourea, 10 mM HEPES pH 8.0) by bath sonication. To cap
cysteine residues, 100 μl of freshly-made TCEP (200 mM stock solution, Thermo) was
then added and the mixture incubated for 30 min, followed by 40 μl of freshly prepared
iodoacetamide (1 M stock solution, Sigma) and incubation for a further 30 min in the
dark. Steptavadin beads (250 μL of a 50% slurry per sample, Thermo) were washed 2 X
with 1 mL PBS and 1 X with 1 mL resuspension buffer and resuspended in resuspension
buffer (200 μL). Each sample was combined with streptavadin beads and incubated on a
rotator for 2 h. These mixtures were then transferred to Mini Bio-Spin
®
columns
(Bio-Rad) and placed on a vacuum manifold. Captured proteins were then washed with
agitation 5 X with resuspension buffer (10 mL), 5 X PBS (10 mL), 5x with 1% SDS in
321
PBS (10 mL), 30x with PBS (1 mL per wash, vacuum applied between each wash), and
5x 2M urea in PBS (1 mL per wash, vacuum applied between each wash). Beads were
then resuspended in 2 M urea in PBS (1 mL), transferred to screw-top tubes, and pelleted
by centrifugation (2000 x g for 2 min). At this time, 800 μL of the supernatant was
removed, leaving a volume of 200 μL. To this bead-mixture was added 2 μL o f CaCl
2
(200 mM stock, 1 mM final concentration) and 2 μL of 1 mg mL
−1
sequence grade
trypsin (Promega) and incubated at 37 °C for 18 hours. The resulting mixtures of tryptic
peptides and beads were transferred to Mini Bio-Spin
®
columns (Bio-Rad) and the eluent
was collected by centrifugation (1,000 x g for 2 min). Any remaining peptides were
eluted by addition of 100 μL of 2 M urea in PBS followed by centrifugation as
immediately above. The tryptic peptides were then applied to C18 spin columns (Pierce)
according to manufacturer's instructions, eluted with 70% acetonitrile in H
2O, and
concentrated to dryness on a speedvac.
LC-MS/MS Proteomic Analysis. Peptides were desalted on a trap column following
separation on a 12cm/75um reversed phase C18 column (Nikkyo Technos Co., Ltd.
Japan). A 3 hour gradient increasing from 10% B to% 45% B in 3 hours (A: 0.1% Formic
Acid, B: Acetonitrile/0.1% Formic Acid) was delivered at 150 nL min
−1
. The liquid
chromatography setup (Dionex, Boston, MA, USA) was connected to an Orbitrap XL
(Thermo, San Jose, CA, USA) operated in top-5-mode. Acquired tandem MS spectra
(CID) were extracted using ProteomeDiscoverer v. 1.3 (Thermo, Bremen, Germany) and
queried against the human Uniprot protein database using MASCOT 2.3.02
322
(Matrixscience, London, UK). Peptides fulfilling a Percolator calculated 1% false
discovery rate threshold were reported. All LC-MS/MS analysis were carried out at the
Proteomics Resource Center at The Rockefeller University, New York, NY , USA. Excel
files containing identified proteins will be made available upon request.
323
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Appendix A: NMR Spectra
374
Compound 2.1 1,3,4-Tri-O-acetyl-2-acetamido-6-azido-2,6-dideoxy-D-
glucopyranose (6AzGlcNAc).
O
AcO
NH
O
N
3
AcO
OAc
375
Compound 2.2 4-(prop-2-yn-1-yl)phenol
OH
376
Compound 2.3 (E)-methyl 4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)benzoate.
OH
N
N
O
O
377
Compound 2.3 (E)-methyl 4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)benzoate.
OH
N
N
O
O
378
Compound 2.4 (E)-4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)benzoic acid.
OH
N
N
OH
O
379
Compound 2.4 (E)-4-((2-hydroxy-5-(prop-2-yn-1-yl)phenyl)diazenyl)benzoic acid.
OH
N
N
OH
O
380
Compound 2.9 α-1-O-benzyl-N-acetyl-glucosamine
O
AcHN
OH
HO
HO
OBn
381
Compound 2.10 α-1-O-benzyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine
O
AcHN
OTs
HO
HO
OBn
382
Compound 2.11 3,4-di-O-acetyl-α-1-O-benzyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine
O
AcHN
OTs
AcO
AcO
OBn
383
O
AcHN
OTs
AcO
AcO
O
P
O
O
O
Compound 2.13 Diallyl(3,4-di-O-acetyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
384
O
AcHN
OTs
AcO
AcO
O
P
O
O
O
Compound 2.13 Diallyl(3,4-di-O-acetyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
385
O
AcHN
OTs
AcO
AcO
O
P
O
O
O
Compound 2.13 Diallyl(3,4-di-O-acetyl-6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
386
Compound 2.14 Diallyl(6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
O
AcHN
OTs
HO
HO
O
P
O
O
O
387
Compound 2.14 Diallyl(6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
O
AcHN
OTs
HO
HO
O
P
O
O
O
388
O
AcHN
OTs
HO
HO
O
P
O
O
O
Compound 2.14 Diallyl(6-O-p-methylbenzenesulfonate-N-acetyl-glucosamine)-α-1-phosphate
389
Compound 2.15 Diallyl(6-azido-6-deoxy-N-acetyl-glucosamine)-α-1-phosphate
O
AcHN
N
3
HO
HO
O
P
O
O
O
390
Compound 2.15 Diallyl(6-azido-6-deoxy-N-acetyl-glucosamine)-α-1-phosphate
O
AcHN
N
3
HO
HO
O
P
O
O
O
391
Compound 2.15 Diallyl(6-azido-6-deoxy-N-acetyl-glucosamine)-α-1-phosphate
O
AcHN
N
3
HO
HO
O
P
O
O
O
392
Compound 2.16 6-azido-6-deoxy-N-acetyl-glucosamine-1-phosphate
O
AcHN
N
3
HO
HO
OPO
3
-2
393
Compound 2.16 6-azido-6-deoxy-N-acetyl-glucosamine-1-phosphate
O
AcHN
N
3
HO
HO
OPO
3
-2
394
Compound 2.16 6-azido-6-deoxy-N-acetyl-glucosamine-1-phosphate
O
AcHN
N
3
HO
HO
OPO
3
-2
395
Compound 3.1 2-Acetamido-2-deoxy-3,4,5,6-di-O-isopropylidine-aldehydo-D-glucose dimethyl acetal
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
3.18
3.27
2.96
3.02
2.96
3.04
3.05
1.03
1.05
2.11
1.07
1.00
1.01
0.90
MeO OMe
NHAc
O
O
O
O
396
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
6.34
3.40
3.30
3.21
3.88
1.08
1.14
1.07
1.00
MeO OMe
NHAc
O
O
OH
OH
Compound 3.2 2-Acetamido-2-deoxy-3,4-O-isopropylidine-aldehydo-D-glucose dimethyl acetal
397
MeO OMe
NHAc
O
O
OH
OMs
Compound 3.3 2-Acetamido-6-O-mesyl-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
3.13
3.23
3.44
3.43
3.47
3.49
1.10
1.08
1.40
1.25
1.21
1.18
1.07
1.00
398
MeO OMe
NHAc
O
O
OH
OMs
Compound 3.3 2-Acetamido-6-O-mesyl-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200
f1 (ppm)
399
Compound 3.4 2-Acetamido-6-cyano-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
MeO OMe
NHAc
O
O
OH
C N
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
3.24
2.99
3.25
1.21
1.18
1.14
3.30
3.35
1.12
1.12
1.16
1.17
1.07
1.00
400
Compound 3.4 2-Acetamido-6-cyano-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
MeO OMe
NHAc
O
O
OH
C N
0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200
f1 (ppm)
401
O O
NHAc
O
O
OH
CHO
Compound 3.5 2-Acetamido-6-aldehydo-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0 10.5 11.0
f1 (ppm)
3.18
3.04
3.42
1.13
1.13
3.64
3.89
0.65
1.11
2.28
1.07
1.03
1.00
0.94
402
O O
NHAc
O
O
OH
CHO
Compound 3.5 2-Acetamido-6-aldehydo-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200
f1 (ppm)
403
O O
NHAc
O
O
OH
Compound 3.6 2-Acetamido-6-alkyne-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
6.71
4.40
1.20
1.18
3.57
3.63
1.18
1.06
1.20
2.19
1.00
404
O O
NHAc
O
O
OH
Compound 3.6 2-Acetamido-6-alkyne-2-deoxy-3,4-O-isopropylidene-aldehydo-D-glucose dimethyl acetal
0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200
f1 (ppm)
405
Compound 3.7 1,3,4-Tri-O-acetyl-2-deoxy-2-N-acetyl-6-deoxy-alkynyl-glucopyranose (Ac 36AlkGlcNAc)
O
AcO
AcO
OAc
NHAc
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0
f1 (ppm)
3.06
5.08
2.82
2.92
2.25
1.03
1.10
2.38
1.67
1.00
406
Compound 3.7 1,3,4-Tri-O-acetyl-2-deoxy-2-N-acetyl-6-deoxy-alkynyl-glucopyranose (Ac 36AlkGlcNAc)
O
AcO
AcO
OAc
NHAc
0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200
f1 (ppm)
407
Compound 4.1 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-glucosamine (Ac 4GlcPoc)
O
AcO
NH
OAc
O
O
OAc
AcO
408
Compound 4.1 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-glucosamine (Ac 4GlcPoc)
O
AcO
NH
OAc
O
O
OAc
AcO
409
O
AcO
NH
OAc
O
O
OAc
AcO
Compound 4.2 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-galactosamine (Ac 4GalPoc)
410
O
AcO
NH
OAc
O
O
OAc
AcO
Compound 4.2 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-galactosamine (Ac 4GalPoc)
411
Compound 4.3 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-manosamine (Ac4ManPoc)
O
AcO
AcO
OAc
AcO
HN O
O
412
Compound 4.3 N-propargyloxycarbamate-1,3,4,6-tetra-O-acetyl-manosamine (Ac4ManPoc)
O
AcO
AcO
OAc
AcO
HN O
O
413
Compound 5.1 3,4,6-Tri-O-Acetyl-1-Chloro-1-Deoxy-N-Acetylglucosamine
O
AcO
OAc
AcO
Cl
NH
O
414
Compound 5.3 1-Deoxyglucosamine hydrochloride
O
HO
OH
HO
NH
3
+
Cl
-
415
Compound 5.4 3,4,6-Tri-O-Acetyl-1-Deoxy-N-4-pentynylglucosamine (1-deoxy-Ac 3GlcNAlk)
O
AcO
NH
O
OAc
AcO
416
Compound 5.4 3,4,6-Tri-O-Acetyl-1-Deoxy-N-4-pentynylglucosamine (1-deoxy-Ac 3GlcNAlk)
O
AcO
NH
O
OAc
AcO
417
Compound 5.5 1-Deoxy-N-4-pentynylglucosamine (1-deoxy-GlcNAlk)
O
HO
NH
O
OH
HO
418
Compound 6.1 2-Azido-2-Deoxyglucose
419
Compound 6.2 1,3,4,6-Tetra-O-Acetyl-2-Azido-2-Deoxyglucose (Ac42AzGlc)
420
Compound 6.3 6-Tert-Butyldimethylsilyl-2-Azido-2-Deoxyglucose (6TBS2AzGlc)
O
HO
N
3
OTBS
HO
OH
421
Compound 6.3 6-Tert-Butyldimethylsilyl-2-Azido-2-Deoxyglucose (6TBS2AzGlc)
O
HO
N
3
OTBS
HO
OH
422
Compound 6.4 1,3,4-Tri-O-Acetyl-6-Tert-Butyldimethylsilyl-2-Azido-2-Deoxyglucose (Ac 36TBS2AzGlc)
O
AcO
N
3
OTBS
AcO
OAc
423
Compound 6.4 1,3,4-Tri-O-Acetyl-6-Tert-Butyldimethylsilyl-2-Azido-2-Deoxyglucose (Ac 36TBS2AzGlc)
O
AcO
N
3
OTBS
AcO
OAc
424
Compound 6.5 1,3,4-Tri-O-Acetyl-2-Azido-2-Deoxyglucose (Ac32AzGlc)
O
AcO
N
3
OH
AcO
OAc
425
Compound 6.5 1,3,4-Tri-O-Acetyl-2-Azido-2-Deoxyglucose (Ac32AzGlc)
O
AcO
N
3
OH
AcO
OAc
426

Abstract (if available)

Abstract Post translational modifications (PTMs) are covalent additions appended to the side chains of amino acids that afford an extra level of biological and chemical complexity to the genome. These modifications can range from the addition of a single methyl group to the installation of a large oligosaccharide. Nevertheless, PTMs have been shown to have the ability to change the function, activity or localization of their target proteins. Furthermore, PTMs can be dynamically regulated and can be added enzymatically or chemically thus allowing them to act as signaling mechanisms or as metabolic sensors. In order to interrogate the consequences of PTMs, many methods have been developed to visualize and identify substrate proteins. One of these methods involves the treatment of live cells with a small molecule memetic of a metabolic precursor, containing an abiotic azide or alkyne, for its enzymatic incorporation onto target proteins and subsequent analysis using copper catalyzed azide alkyne cycloaddition (CuAAC), or click chemistry. These small molecules are termed metabolic chemical reporters (MCRs) and have been extensively utilized in the study of protein glycosylation. Described here is the development and optimization of the first MCRs to be specific for a single type of protein glycosylation called O-GlcNAc. O-GlcNAc modification is defined by the enzymatic addition of a single monosaccharide termed N-acetyl-glucosamine (GlcNAc) to serine and threonine residues of intracellular proteins. With this novel MCR in hand, the modification of the apoptotic regulator proteins called the caspases was revealed and was subsequently shown to affect the cleavage and activation of the protein during cell death. In addition, described here are three N-propargyloxycarbamate MCRs for the investigation of carbohydrate salvage pathways, an MCR for visualizing the metabolic cross-talk between carbohydrate metabolism and protein acetylation, and finally the small molecule 2-azido-2-deoxy-glucose, a reporter that revealed the unexpected promiscuity of O-GlcNAc transferase, the enzyme responsible for the addition of O-GlcNAc.

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Asset Metadata

Creator Chuh, Kelly Nicole (author)

Core Title Development of metabolic chemical reporters for the investigation of protein glycosylation

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Chemistry

Publication Date 09/21/2017

Defense Date 07/17/2017

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag chemical reporters,click chemistry,glycosylation,metabolic chemical reporters,OAI-PMH Harvest,O-GlcNAc,post translational modifications

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Pratt, Matthew (committee chair), Chen, Lin (committee member), Qin, Peter (committee member)

Creator Email KChuh@usc.edu,KNChuh@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-431442

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Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA