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UNDERSTANDING AND MANIPULATING GANGLIOSIDE BIOSYNTHESIS
A DISSERTATION
SUBMITTED TO THE DEPARTMENT OF CHEMISTRY
AND THE COMMITTEE ON GRADUATE STUDIES
OF STANFORD UNIVERSITY
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
Chad Michael Whitman May 2010
http://creativecommons.org/licenses/by-nc/3.0/us/
This dissertation is online at: http://purl.stanford.edu/qp110yy2867
© 2010 by Chad Michael Whitman. All Rights Reserved.
Re-distributed by Stanford University under license with the author.
This work is licensed under a Creative Commons Attribution-Noncommercial 3.0 United States License.
ii
I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.
Jennifer Kohler, Primary Adviser
I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.
Justin Du Bois, Co-Adviser
I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.
Chaitan Khosla
Approved for the Stanford University Committee on Graduate Studies.
Patricia J. Gumport, Vice Provost Graduate Education
This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file inUniversity Archives.
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Abstract
Understanding and Manipulating Ganglioside Biosynthesis
By
Chad Michael Whitman
Doctor of Philosophy in Chemistry
Stanford University
The mammalian cell surface is comprised of a heterogeneous mixture of proteins
and lipids decorated by carbohydrates. These glycoproteins and glycolipids are known as
glycoconjugates. Among the many types of glycolipids imbedded within the plasma
membrane resides a class of negatively charged species known as gangliosides.
Characterized by the presence of sialic acid residues, gangliosides are responsible for
regulating the activity of many cell surface proteins and serve as recognition targets for
cell-cell communication and pathogen invasion. In Chapter 1, I introduce the
biosynthetic pathway for mammalian ganglioside synthesis. Next, I describe several
recognized biological roles of gangliosides. Finally, I present recent biochemical
methods used for elucidating biological function at the molecular level.
Gangliosides are synthesized in the endoplasmic reticulum and Golgi by
numerous membrane-bound glycosyltransferases. These glycosyltransferases catalyze
the addition of monosaccharides in a sequential fashion, resulting in a diverse assortment
of gangliosides. Ganglioside biosynthetic enzymes have been shown to associate with
one another, forming biosynthetic clusters. Previous work demonstrated that associations
among these enzymes are controlled by their N-terminal regions, which include a single-
pass transmembrane domain (TM). The formation of homo-oligomeric complexes via
TM domain interactions has also been implicated in regulating glycosyltransferase
function and localization. In Chapter 2, I characterize the interactions among TM
domains of five ganglioside glycosyltransferases. Using an assay that measures TM
domain association in SDS micelles, I discovered that three of the TM domains homo-
oligomerize, including one that forms trimers, pentamers, and higher-order oligomers.
v
To investigate the biological importance of these associations, I employed fluorescence
microscopy and western blot analysis to detect enzyme homo-oligomerization occurring
in transiently transfected mammalian cells.
Gangliosides play critical roles in the regulation of cell signaling and in pathogen-
cell recognition, but the molecular details of many of these processes are poorly
understood. Challenges to biochemical characterization are posed by the transient nature
of ganglioside-mediated interactions and the difficulty in isolating ganglioside-protein
complexes. To surmount these obstacles, I investigate the application of metabolic
oligosaccharide engineering techniques to synthesize photocrosslinking gangliosides in
mammalian cells, as described in Chapter 3. I observed that the photoactivatable GM1
ganglioside analog is recognized by cholera toxin, a well-characterized ganglioside
interactions partner. After demonstration that this complex could be covalently captured
in cells, I have begun preliminary experiments to ascertain the effects complex formation
on the retrograde trafficking mechanisms required for cholera intoxication.
The ability to introduce small structural changes into cell surface glycans through
metabolic oligosaccharide engineering methods has aided in the study of many
carbohydrate-mediated interactions. These techniques have also proved effective at
installing bio-orthogonal reactive groups to visualize glycan structures in both cells and
animals. While previous studies have illustrated the many biological roles of
glycoconjugates in mammalian cells, these studies focused primarily on global levels of
unnatural glycan incorporation and not specific glycoconjugates structures. In Chapter
4, I investigate the substrate selectivity associated with sialic acid engineering of
gangliosides in mammalian cells. I observed substantial differences in unnatural sialic
acid incorporation between cell lines derived from different mammalian species. These
differences may reflect naturally occurring dissimilarities in the ability of different
species to incorporate variant sialic acids into their glycoconjugates.
vi
Acknowledgements
I would like to thank my advisor, Jennifer Kohler, for serving as my mentor for
the past five years. Between our time at Stanford and UT Southwestern, I’ve learned a
great deal about becoming a better scientific researcher and communicator. Her guidance
has allowed me to explore many different scientific techniques in both chemistry and
biology. She has also provided me with the opportunity to mentor fellow labmates and
rotation students. I am forever grateful for the opportunity that she gave me and I only
wish her the best for many years to come.
Being a part of two universities for my graduate career, I have the awesome
opportunity to work with a large number of gifted scientists. Particularly, I would like to
thank Michelle Bond. Through our 4+ years together, we have formed an incredible
bond that has allowed us to work together and develop a deep friendship. I have an
immense amount of respect for her abilities and I hope that we can continue our
friendship as we both move to the opposite sides of the country.
While at Stanford, I had the privilege of working with a talented group of
graduate students and postdocs. Ethan Greenblatt is a great friend who continues to
update me with all of his current experiments, both public and secret, that he is pursuing.
Peter Lee was always willing to help me with my microscopy experiments and was
always the source of great late night conversations. Danielle Dube, our lab’s first
postdoc, was very generous helping me with my presentation skills. Yoshi Tanaka was a
great synthetic chemist who taught me a lot about science and Japanese culture. Meg
Desko started the group with me at Stanford and was a great person to grab coffee with
and chat for hours.
The current members of the group at UT Southwestern have also been equally as
talented as my previous ones. Nam Pham, Randy Parker, and Peter Vu (the first wave of
graduate students in the group) have been a privilege to work with to pass on my
scientific knowledge. I look forward to hearing about their endeavors for years to come
and excited for the future of lab being in their hands. Seokho Yu, Fan Yang, and Bin Li
have bestowed upon me many years of knowledge and experience that have helped me
develop my technical skills. While we have only overlapped for a few months, Rubina
vii
Tuladhar has always been a great source of endless conversation and is a very dedicated
person who will do great things in her upcoming graduate career.
There are several people at Iowa State University who influenced my path to
pursuing my doctorate degree. Prof. Joe Burnett was my instructor for several analytical
chemistry courses and encouraged me to not give up on chemistry. He has also been a
great friend who came and visited me twice while in California. Prof. Victor Lin served
as my research mentor for my senior year and if not for his advisement, I never would
have considered attending graduate school. While he has become a very busy individual,
he still takes time out of his busy schedule to catch up with me and discuss my current
and future endeavours. Brian Trewyn, a graduate student in Victor’s lab, taught me how
to be an effective researcher and how to keep one’s sanity while in graduate school. As
for my classmates, Sassan Sheikholeslami and Ali Mostrom, I’ll never forget our
countless hours spent studying and goofing off at the Cyclone Truck Stop. I’d also like
to thank all of the members of Triangle Fraternity during my undergraduate experience
that helped me develop personally and professionally.
I would also like to thank my parents, Rick and Linda Whitman, for their
incredible support over the years, providing me with countless opportunities to grow and
learn. While I know it was hard for their only son to “leave the nest” and go out West,
I’ve made it a commitment to keep in touch and keep them updated with all my
adventures. Finally, I would like to thank my fiancé Laura McNabb. She has made
many sacrifices for me to complete my degree and I can’t describe into words what she
has meant to me. I’m excited to soon being leaving Texas and returning back to
California with her to begin a new career and start our family.
viii
Table of Contents
Chapter 1 – The biosynthesis and biological roles of gangliosides ..............................1
Introduction.................................................................................................................1 Biosynthesis of gangliosides........................................................................................3 Complexes of glycosyltransferases synthesize gangliosides .........................................4 Gangliosides mediate interactions on the cell surface...................................................6 The role of NeuGc expression in gangliosides .............................................................7 Metabolic oligosaccharide engineering of gangliosides as a potential tool for studying gangliosides.................................................................................................................9 Photocapturing glycosphingolipid interactions in cells...............................................11 Conclusions ...............................................................................................................13 References.................................................................................................................14
Chapter 2 – The biosynthesis and biological roles of gangliosides ............................22
Introduction...............................................................................................................22 Results.......................................................................................................................24
In vitro assay to investigate TM interactions ..........................................................24 Investigating TM interactions in vitro ....................................................................25 Functional analysis of ganglioside glycosyltransferases .........................................29
Discussion .................................................................................................................34 The TM domains of ganglioside glycosyltransferases fail to demonstrate the formation of hetero-oligomeric species with the SN-TM assay ..............................34 The TM domains of several ganglioside glycosyltransferases mediate formation of homo-oligomers.....................................................................................................35 SialT1 and SialT2 enzymes are physically associated in cells.................................37 SialT2 demonstrates the formation of higher-order complexes in cells ...................37
Acknowledgements ...................................................................................................38 Methods ....................................................................................................................39 References.................................................................................................................47
ix
Chapter 3 – Covalently capturing the ganglioside GM1-cholera toxin complex with crosslinking..................................................................................................................52
Introduction...............................................................................................................52 Results.......................................................................................................................53
HPTLC analysis of gangliosides produced by Ac4ManNDAz-treated Jurkat cells ..53 Mass spectrometry analysis of gangliosides produced by Ac4ManNDAz-treated Jurkat cells.............................................................................................................57 Immunofluorescence microscopy analysis of CTxB trafficking in Ac44ManNDAz-treated Jurkat cells exposed to UV light .................................................................58
Discussion .................................................................................................................61 Jurkat cells can synthesize gangliosides with photoreactive chemical groups .........61 Cholera toxin subunit B recognizes GM1-SiaDAz and can be efficiently photocrosslinked....................................................................................................62 The formation of a covalent GM1-CTxB complex does not appear to affect trafficking from the plasma membrane to the TGN ................................................63
Acknowledgements ...................................................................................................64 Methods ....................................................................................................................65 References.................................................................................................................72
Chapter 4 – Exploring the incorporation of sialic acid analogs into gangliosides of mammalian cells ..........................................................................................................75
Introduction...............................................................................................................75 Results.......................................................................................................................77
Incorporation of variant sialic acids into gangliosides ............................................77 Analysis of engineered cell surface sialylation with BJAB and CHO cell lines cultured with ManNAc analogs..............................................................................81
Discussion .................................................................................................................84 Incorporation of modified sialic acids into the GM3 ganglioside of BJAB cells .....84 Incorporation of modified sialic acids into the GM3 ganglioside of CHO cells.......86 The ST3Gal5 enzyme of BJAB cells shows a reduced capacity for engineering NeuGc-containing gangliosides .............................................................................86 Future directions....................................................................................................87
Acknowledgements ...................................................................................................88 Methods ....................................................................................................................89 References.................................................................................................................99
Appendix.................................................................................................................... 105
1H NMR Data.......................................................................................................... 106 13H NMR Data......................................................................................................... 110 ESI-MS Data ........................................................................................................... 114 HPLC Data.............................................................................................................. 118 MALDI-TOF-MS Data............................................................................................ 122
x
List of Illustrations
Chapter 1 – The biosynthesis and biological roles of gangliosides ..............................1
Figure 1. Structure of a typical glycosphingolipid........................................................1 Figure 2. Chemical structure of GM1 ganglioside........................................................2 Figure 3. Biosynthesis of gangliosides .........................................................................4 Figure 4. Ganglioside glycosyltransferases are type II transmembrane proteins............5 Figure 5. Common sialic acid structures ......................................................................7 Figure 6. Examples of unnatural sialic acid analogs incorporated onto cellular glycoconjugates .........................................................................................................10 Figure 7. Common photoreactive groups used in biology...........................................12
Chapter 2 – The biosynthesis and biological roles of gangliosides ............................22
Figure 1. Reported associations among glycosyltransferases responsible for ganglioside biosynthesis ............................................................................................23 Figure 2. N-terminal sequences of ganglioside glycosyltransferases implicated in their association.................................................................................................................25 Figure 3. SDS-PAGE analysis of SN-TM homo- and hetero-oligomerization.............26 Figure 4. SDS-PAGE analysis of SN-GalT1 TM WT and mutated fusion proteins.....27 Figure 5. SDS-PAGE analysis of SN-SialT2 TM WT and mutated fusion proteins ....28 Figure 6. KDEL recruitment assay.............................................................................30 Figure 7. Cellular association of ganglioside glycosyltransferases visualized through the KDEL recruitment assay ......................................................................................32 Figure 8. Full length SialT2 enzyme shows homo-oligomerization ............................34
Chapter 3 – Covalently capture the ganglioside GM1-cholera toxin complex with crosslinking..................................................................................................................52
Figure 1. Acetylated analogs used to investigate metabolic oligosaccharide engineering of gangliosides...........................................................................................................54 Figure 2. HPTLC analysis of gangliosides from cultured Jurkat cells.........................56 Figure 3. MALDI-TOF-MS analysis of Jurkat cells cultured with Ac4ManNDAz ......58 Figure 4. Immunofluorescene analysis of Cholera toxin subunit B trafficking in supplemented Jurkat cells ..........................................................................................60
Chapter 4 – Exploring the incorporation of sialic acid analogs into gangliosides of mammalian cells ..........................................................................................................75
Figure 1. Metabolic oligosaccharide engineering of UDP-GlcNAc 2-epimerase deficient cells occurs through the introduction of ManNAc and sialic acid analogs ....78 Figure 2. Panel of ManNAc analogs used to ganglioside incorporation experiments ..79 Figure 3. HPTLC analysis of gangliosides produced by BJAB and CHO cells cultured with ManNAc analogs ...............................................................................................80 Figure 4. Flow cytometry analysis of cells surface display of modified sialic acids ....83
xi
List of Tables
Chapter 2 – The biosynthesis and biological roles of gangliosides ............................22
Table 1. Table of primers used to generate SN-TM plasmids .....................................40 Table 2. Table of primers used to generate SN-GalT1 TM mutants............................41 Table 3. Table of primers used to generate SN-SialT2 TM mutants ...........................41 Table 4. Table of primers used to generate KDEL recruitment assay plasmids ...........45
* Several sections in this Chapter were adapted from Whitman, C.M.; Bond, M.R.; Kohler, J.J. In Comprehensive Natural Products II Chemistry and Biology; Mander, L., Lui, H.-W., Eds.; Elsevier: Oxford, 2010; Volume 6, pp 175-224
1
Chapter 1 – The biosynthesis and biological roles of gangliosides*
Introduction
Glycosphingolipids are a heterogeneous class of molecules that decorate the
cellular surface of eukaryotic cells. These amphiphatic structures are anchored into the
membrane by their long hydrocarbon chains and present a variety of carbohydrate
structures to the extracellular environment (Figure 1). Various lipid scaffolds and
monosaccharides combine to form an immense diversity of glycosphingolipids; over 500
different structures have been characterized.1 Glycosphingolipid interactions play critical
roles in all aspects of cell surface recognition. As components of the plasma membrane,
glycosphingolipids utilize their glycan components to modulate the functions of many
cell surface proteins.2 Glycosphingolipids also serve as recognition targets in many cell-
cell interactions,3 Unfortunately, a variety of pathogens, such as cholera toxin and tetanus
toxin, exploit glycosphingolipids to mediate selective binding to their hosts for cell
invasion.4,5
Figure 1. Structure of a typical glycosphingolipid. Glycosphingolipids are amphiphatic molecules that are anchored into the plasma membrane by a hydrophobic ceramide, composed of a sphingosine (blue) and fatty acid (yellow). The ceramide scaffold is decorated with wide variety of glycans, initiating with either glucose or galactose (green).
Glycosphingolipids are divided into seven structural classes based upon their
glycan linkages to ceramide. While the majority of these structures are classified as
neutral glycosphingolipids, the ganglioside class comprises negatively charged species at
2
physiological pH due to the inclusion of sialic acids. While the term “ganglioside” is
often used informally to refer to any sialylated glycosphingolipids, the official
ganglioside class is derived from their common structure of Galβ1,4-Glcα-ceramide, also
known as lactosylceramide (LacCer). Figure 2 shows the chemical structure of the GM1
ganglioside. As detailed below, the biosynthesis of gangliosides starting from LacCer is
quite diverse and will vary by cell type depending on the transcriptional regulation of the
glycosyltransferases responsible for their synthesis. Changes in amount and variety of
gangliosides are observed in many eukaryotic organisms during embryonic brain
development6 and in many types of cancers.7 These changes are critical for facilitating
many ganglioside-based interactions that are occur in these settings. While changes in
ganglioside content are well-documented, the functional implications of these
fluctuations remains unclear.
Figure 2. Chemical structure of GM1 ganglioside. Gangliosides are a class of molecules that share the common structure of Galβ1,4-Glcα-ceramide, known as lactosylceramide (LacCer). Gangliosides contain at least one sialic acid residue in their structure (highlighted in red). Shown above is the chemical structure of a GM1 ganglioside.
Discovery of the genes responsible for ganglioside biosynthesis occurred during
the 1980s and 1990s and facilitated investigations into the many biological roles of these
molecules. With the dawn of chemical biology, chemist and biologists have worked
together to develop an assortment of tools to explore the molecular function and
specificity of gangliosides. For example, the synthesis of photoreactive glycolipids has
provided molecular details for ganglioside binding to tetanus toxin8 and two receptor
tyrosine kinases.9,10 While these studies have yielded information about ganglioside-
mediated interactions, there are still many unanswered questions such as determining
why eukaryotic cells modify their ganglioside composition during tumorigenesis.
3
Nonetheless, the continual advance of chemical and biological techniques will only
further improve our capability to understand the many biological roles of gangliosides.
Biosynthesis of gangliosides
The biosynthesis of gangliosides occurs through the step-wise addition of
nucleotide-activated monosaccharides by glycosyltransferases (Figure 3). Ganglioside
synthesis begins in the endoplasmic reticulum (ER) where ceramide is produced and
displayed on the cytosolic face of the ER membrane. Ceramide is then transported to the
Golgi membrane where it is initially glucosylated (GlcCer) on the cytosolic face to
generate GlcCer. Afterwards, GlcCer is translocated by unidentified mechanisms to the
luminal side of the Golgi for further modification. The attachment of galactose (GalT1)
onto GlcCer generates lactosylceramide (LacCer), the basic building block for the
ganglioside class of glycosphingolipids. Upon reaching the cis-Golgi, LacCer is initially
sialylated (SialT1) to generate GM3. These species can be further modified with
additional sialic acid (SialT2, SialT3) to generate GD3 and GT3. Transfer of these
species towards the trans-Golgi network (TGN) leads to the synthesis of more complex
gangliosides by sequential additions of N-acetylgalactosamine (GalNAcT), galactose
(GalT2), and more sialic acids (SialT4, SialT5). The exact cellular composition of
gangliosides can depend on a large range of factors, including cell type, enzyme levels,
post-translational processing of glycosyltransferases, nucleotide-sugar levels, self-
inhibition by ganglioside products, and pH.11
4
Figure 3. Biosynthesis of gangliosides. Gangliosides are synthesized in step-wise fashion through the addition of monosaccharides (see table insert) by glycosyltransferases (shown in red). Cartoon structures represent the chemical structure of individual gangliosides. Names of ganglioside species are listed below structures. Series classifications of gangliosides families are denoted at the bottom.
Complexes of glycosyltransferaes synthesize gangliosides
All ganglioside glycosyltransferases are type II transmembrane proteins,
consisting of a N-terminal cytoplasmic tail (CT), single-pass transmembrane domain
(TMD), stem region, and a C-terminal catalytic domain (Figure 4). These enzymes are
localized within the various compartments of the secretory pathway, primarily through
their N-terminal region (CT, TMD, and stem).12 Two models describe the retention of
these enzymes to the secretory pathway. The first model, the lipid bilayer thickness or
sorting model,13,14 proposes that glycosyltransferases are retained within the Golgi due to
the smaller size of the TMD compared to plasma membrane proteins. It is known that the
plasma membrane is thicker than Golgi membranes due to the increased presence of
cholesterol, which has been observed to increase lipid bilayer thickness.15 By extending
the length of the TMD domain of a Golgi-resident sialyltransferase (ST6Gal1) from 17 to
5
23 amino acids, Munro showed that the localization was redirected to the plasma
membrane.16 An inverse transposition could be used to relocate plasma membrane
proteins to the Golgi.17 Despite these initial reports, later work showed that the targeting
of ST6Gal1 was not limited to its TMD and could be directed by lumenal sequences
through enzyme homo-oligomerization,18,19 suggesting that the lipid bilayer thickness
model alone could not completely describe Golgi-localization of glycosyltransferases.
Figure 4. Ganglioside glycosyltransferases are type II transmembrane proteins. Type II transmembrane proteins are comprised of an N-terminal cytoplasmic tail, single-pass transmembrane domain, stem region, and C-terminal catalytic domain.
The second model, the oligomerization or aggregation model,20,21 proposes that
formation of oligomeric complexes of glycosyltransferases within the Golgi prevents
their delivery to secretory vessels, thus causing them to be retained. Several recent
reports involving ganglioside glycosyltransferases have provided evidence for this second
model. Work from the Maccioni group identified two independent complexes formed
from ganglioside glycosyltransferases: 1) GalT1, SialT1, and SialT2 in the proximal
Golgi22 and 2) GalNAcT and GalT2 in the TGN23 of CHO cells. The formation of these
complexes was found to be important for their localization within the secretory pathway
6
and could facilitate substrate channeling in ganglioside synthesis. Furthermore, they also
showed that the N-terminal region was sufficient for mediating complex formation. In
separate experiments, the Yu group identified a complex formed from SialT2 and
GalNAcT in the Golgi of F-11A murine neuroblastoma cells.24 These differing reports
should not be assumed as mutually exclusive, rather complex formation may be cell-type
specific and based upon the transcriptional regulation of ganglioside glycosyltransferases.
The study of homo-oligomerization of ganglioside glycosyltransferases and their
potential effects on biological function is explored in Chapter 2.
Gangliosides mediate interactions on the cell surface
Upon delivery to the extracellular plasma membrane of cells, gangliosides have
been shown to play important roles in the regulation of cell-cell interactions. Changes in
cell-surface glycosylation are frequently observed in tumor cells, which are often a target
for an immune response. Natural killer (NK) cells, a specialized lymphocyte cell, will
naturally target and suppress these cells. However, the expression of GD3 on the surface
of tumors cells activates an inhibitory response against NK cell-mediated toxicity through
Siglec-7 binding, allowing tumor cell proliferation to occur.25 Complex gangliosides are
also shown to play critical roles in the central nervous system. Nerve tissues utilize a
myelin-associated glycoprotein (MAG) to stabilize axon-myelin interactions within the
brain. Additionally, this protein is shown to inhibit nerve regeneration after severe
injuries to the central nervous system. The control of these functions is regulated by
highly specific interactions involving the complex gangliosides GD1a and GT1b.26,27
Cell surface gangliosides, such as GM3 and GD1a, also serve as both positive and
negative regulators of several receptor tyrosine kinases. The epidermal growth factor
receptor (EGFR) is a cell surface protein that binds to growth factor ligands and initiates
several downstream signaling cascades. Upregulation of EGFR signaling is thought to be
a critical event in the development of several types of tumors. This response can be
directly inhibited by interactions with GM3 at the cell surface without interfering with
epidermal growth factor (EGF) binding.28 Elevated levels of cell surface GM3 has been
shown to diminish the signaling ability of the insulin receptor (IR), implicating GM3 as a
regulator of insulin sensitivity and perhaps type II diabetes.29 Cancer cell proliferation
7
depends upon angiogenesis through signaling mechanisms involving the vascular
endothelial growth factor receptor (VEGFR). Introduction of GD1a to the surface of
human vascular endothelial cells (HUVECs) has been shown to enhance VEGFR
signaling and cell growth.30 Conversely, this signal can be attenuated by the expression of
GM3, which is the primary ganglioside expressed by these cells.31 These results highlight
the importance of gangliosides in a variety of recognition and signaling events and
highlight ability to modulate protein function through highly specific interactions.
The role of NeuGc expression in gangliosides
Sialic acids are a family of nine-carbon α-keto acids derived from N-
acetylneuraminic acid (NeuAc), N-glycolylneuraminic acid (NeuGc) and deaminated
neuraminic acid (KDN) (Figure 5). Sialic acids are typically found attached to the
nonreducing terminus of glycoproteins (both N-linked and O-linked) and
glycosphingolipids in vertebrates and “higher” invertebrates such as starfish and sea
urchins. This terminal localization allows them to serve as ligands for numerous selectins
and siglecs to mediate a large number of cell-cell adhesion processes in inflammation and
the immune response.32,33 Sialic acid-containing glycoconjugates are commonly modified
by post-glycosylational processing, leading to over 50 naturally occurring variants of
sialic acid.34,35
Figure 5. Common sialic acid structures. Sialic acids are group of nine-carbon α-keto acids derived from N-acetylneuraminic acid (NeuAc), N-glycolylneuraminic acid (NeuGc), and deaminated neuraminic acid (KDN).
The most common forms of sialic acid in mammals are NeuAc and NeuGc, which
are attached onto glycoconjugates by sialyltransferases that utilize cytidine
monophosphate (CMP) activated sugars. CMP-NeuGc is synthesized through the
hydroxylation of CMP-NeuAc by CMP-NeuAc hydroxylase (CMAH). While NeuGc is
8
abundant in most mammals, it is found at extremely low levels in human serum and
organs.7 The genomic source of humans’ inability to synthesize CMP-NeuGc occurs from
an exon deletion/frameshift in the human CMAH gene36,37 estimated to have occurred
~2.5-3 million years ago.38 While this mutation has rendered humans unable to naturally
synthesize NeuGc, it can be acquired from dietary sources, including red meat (lamb,
pork and beef) and milk, and incorporated into human glycoconjugates.39
NeuGc-containing gangliosides are key recognition elements of numerous
pathogens. E. coli infection in mammals is caused by attachment of adhesion proteins
(such as fimbriae or pilli structures) to the intestinal epithelium where the bacteria can
colonize. The K99 fimbriae, generated by E. coli K12, is known to cause severe neonatal
diarrhea to many developing mammals but not in humans; the cause has been elucidated
to occur through a NeuGc-dependent interaction with intestinal gangliosides.40 Subtilase
cytotoxin (SubAb) is an AB5 toxin secreted by Shiga toxigenic E. coli that causes
gastrointestinal disease in humans.41,42 Attachment of this toxin to humans by NeuGc is
greatly enhanced by expression of NeuGc glycoconjugates, presumably through dietary
acquisition of NeuGc. Simian virus 40 (SV40), which utilizes GM1 ganglioside for cell
surface binding, shows significant increases in binding efficiency for GM1-NeuGc over
GM1-NeuAc.43 To better understand the source of NeuGc-containing glycoconjugates in
humans, the NeuGc specificity of the ganglioside glycosyltransferase ST3Gal5 will be
explored in Chapter 4.
Changes in cell surface glycosylation are a hallmark of cancer. Specifically,
many types of tumor cells exhibit altered ganglioside expression, including introduction
of NeuGc.7 As humans do not naturally synthesize NeuGc, its cell surface presentation is
recognized as foreign by the immune system. Consequently, the human immune system
can generate antibodies that specifically recognize the GM3 antigen. In the 1920s,
Hanganutziu44 and Deicher45 independently observed the cellular expression of GM3-
NeuGc in patients treated by injections containing animal antisera. Known today as the
Hanganutziu-Deicher (HD) antigen, the molecular targeting of this antigen by antibodies
is dependent on the presence of NeuGc.46,47 Since these initial studies, researchers have
uncovered a large variety of anti-NeuGc antibodies that are commonly found in healthy
9
humans.48,49 Chickens, which are also unable to naturally synthesize NeuGc, have been
successfully utilized for generating specific anti-ganglioside antibodies.50 Unfortunately,
recent studies have postulated that anti-NeuGc antibody recognition may be a survival
mechanism used by cancer cells to enhance their own propagation.51,52
Metabolic oligosaccharide engineering of gangliosides as a potential tool for
studying gangliosides
The ability to interface chemical techniques and tools from organic chemistry into
the investigation and manipulation of living systems has enabled the expansion of
understanding how biological molecules interact together. The application of this
methodology into protein and lipid glycosylation is known as metabolic oligosaccharide
engineering. Metabolic oligosaccharide engineering refers to the ability to introduce
small structural changes into cellular glycans through the use of unnatural
monosaccharide analogs. To install these glycan modifications, synthetic analogs of
monosaccharide are added to the media of cultured cells or injected into animals.53 Once
inside, endogenous cellular machinery converts the sugar analogs to activated nucleotide
sugar donors, which can be transferred to glycoconjugate substrates. Using this
technology, a diverse class of chemical modifications (extended N-acyl chains, ketones,
azides, thiols, alkynes, diazirines) has been introduced into a variety of sugars, including
sialic acid (Figure 6).53 These non-biological reactive groups have provided unique
chemical reactivity to glycans, which can be used to better understand their function and
localization in biological organisms, without labor-intensive synthetic methods.
Furthermore, these molecules are installed into a wide variety of glycan structures,
allowing for the investigation of a large number of protein interaction partners.
10
Figure 6. Examples of unnatural sialic acid analogs incorporated onto cellular glycoconjugates. Using metabolic oligosaccharide engineering techniques, a large number of chemical modifications have been introduced into cell surface sialic acids (shown in red). The assigned names of each molecule are listed below each structure; attachment to cell surface glycoconjugate is denoted by “R”.
The application of metabolic oligosaccharide engineering as a tool to investigate
the biological roles of gangliosides is a relatively new area of study. The introduction of
bio-orthogonal reactivity into sialic acid structure has been explored as a method for
labeling of mammalian glycans. The incorporation of azides into the N-acyl chain of
sialic acid yields molecular that can be selectively reacted with various reagents inside of
cells and in animals.54 Utilizing the metabolic precursor ManNAz, Bussink et al. showed
successful engineering of GM3 with the metabolized sialic acid analog SiaNAz.55
Interestingly, they observed higher ratios of SiaNAz incorporation into gangliosides over
cell surface glycoproteins. As an initial proof-of-principle experiment, these results
underscore the potential application of monitoring ganglioside levels on the surface of
cells.
Changes in ganglioside expression have been observed in several types of cancer,
including melanoma, colon cancer, and breast cancer.7 These distinct changes in
gangliosides offer a potential target for cancer therapeutics.56,57 An emerging strategy
toward this goal takes advantage of metabolic oligosaccharide engineering to introduce
11
slight structural changes into gangliosides. Even small changes can be sufficient to make
the modified gangliosides appear foreign and immunogenic. These techniques also
ensure that the specific ganglioside response will be active only when unnatural sugar
analogs are administered, minimizing the risk of an autoimmune response. In an attempt
to improve the immunogenicity of GD3, Jennings and co-workers used metabolic
oligosaccharide engineering to introduce a butyryl group to the N-acyl side chain of both
sialic acids that could be used for targeting.58 Upon generation of a highly specific
antibody recognizing this unnatural ganglioside, they demonstrated that they could
selectively target melanoma cells cultured with the N-butyrylated ManNAc precursor,
ManNBut. More remarkably, mice harboring melanoma-derived tumors demonstrated
suppressed tumor growth when treated with combination therapy comprised of the anti-
GD3Bu and ManNBut. While these results appeared promising, the treatment was
unable to eliminate established tumors.
Guo and co-workers have employed a similar approach for development of an
immunotherapy directed against GM3 through introduction of a phenylacetyl group into
the N-acyl side chain.59-61 Using their own highly specific anti-GM3PhAc antibody, they
demonstrated selective targeting of melanoma cells cultured with N-phenylacetyl
ManNAc precursor, ManNPhAc. This targeting was found to be highly cytotoxic to the
ManNPhAc cultured cells. These results suggest that immunotherapy directed against
metabolically engineered gangliosides could provide a potential cancer treatment with
minimal toxic side effects.
Photocapturing glycosphingolipid interactions in cells
Despite some knowledge of the biological roles that glycosphingolipids play in
the plasma membrane, minimal molecular details of the observed interactions are known.
Glycan-dependent binding events are low affinity and suffer from fast off rates. As a
result, these macromolecular complexes tend to dissociate rapidly and are difficult to
isolate by common purification methods. Photocrosslinkers offer a potentially powerful
approach for capturing glycosphingolipid-protein complexes and mapping the binding
surfaces. Upon activation by specific wavelengths, these molecules transform into highly
reactive species that form covalent adducts with nearby atoms. The most common
12
photoreactive groups with demonstrated utility in biological settings include
benzophenones, aryl azides, and diazirines (Figure 7).62,63 The incorporation of
photoreactive functional groups through metabolic oligosaccharide engineering presents
an opportunity to capture and characterize glycosphingolipid-mediated interactions.
Figure 7. Common photoreactive groups used in biology. Benzophenones, aryl azides, and diazirines are some of the most common photoreactive groups that are used in biological samples. Each of these molecules can be activated with UV light to produce reactive species that form covalent adducts with nearby targets.
A variety of approaches have been employed to study ganglioside interactions
with photochemistry. Photoreactive groups have been incorporated into either the
carbohydrate moiety, to detect glycan-based interactions, or in the fatty acid chains, to
detect lipid-mediated interactions.64,65 Caveolae are plasma membrane invaginations that
are implicated in endocytosis and signal transduction pathways. Using radiolabeled,
photoreactive ganglioside analogs, several groups have reported the presence of GM1
within these domains66,67 and have also identified potential binding partners for these
molecules.66 Gangliosides are also known to be important for regulating signal
transduction events on the cell surface. To this end, photoreactive ganglioside analogs
have been successful at capturing complexes with Src-family kinases (c-Src, Lyn)68 and
receptor tyrosine kinases (insulin receptor).9 Additionally, a GM1 probe was
photocrosslinked to α- and β-tubulin,69 suggesting potential mechanistic roles for
gangliosides in the structural remodeling of the cell.
The primary advantage of installing photoreactive groups into gangliosides is
their usefulness at capturing glycan-binding events in situ to uncover underlying
molecular details. Bacterial toxins, such as tetanus toxin, invade the host cell through
molecular recognition of cell surface gangliosides. Starting with naturally occurring
GD1b, Schnaar and coworkers attached an aryl-azide group radiolabeled with 125I onto
the N-acyl chain of sialic acid and were able to capture tetanus toxin, enzymatically
13
digest the complex and isolate the binding region.8 The location of the sialic-acid binding
site was later confirmed by cocrystal structures of the toxin bound to lactose and a GD1a
analog.70,71
The metabolic breakdown of all glycosphingolipid species occurs within the
lysosome. Through the action of many glycosidases and essential cofactors, these
molecules are degraded by step-wise mechanisms into reusable metabolites for a cell.
Using a GM2 analog containing a trifluoromethyl phenyl diazirine and 14C-label,
Sandhoff and coworkers captured the GM2-activator protein (GM2AP) and indentified a
direct interaction occurring between the ceramide of GM2 and a surface loop on
GM2AP.72 These results were consistent with reported crystal structures of GM2AP that
had identified this loop as the most flexible surface of the protein and was most likely to
act as a ligand-binding domain.73,74 Chapter 3 will explore the incorporation of the
photoreactive diazirine group into the gangliosides of Jurkat cells and its applicability in
capturing the glycan-mediated interaction of GM1 to cholera toxin subunit B.
Conclusions
Gangliosides are an important class of glycosphingolipid that regulate numerous
cell surface events. The diversity of structures that can be synthesized allow for an
individual cell to maintain many of these processes simultaneously. Changes in cell
surface ganglioside expression have been shown to play important roles in embryo
development and in cancer progression, but we currently have a limited understanding of
the molecular mechanisms that underlie these important changes. Through the continual
development of biological and chemical tools, we have learned much about gangliosides
but there is still a large amount of information remaining to be uncovered and
understood.
14
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22
Chapter 2 - Exploring the molecular basis for associations among glycosphingolipid
glycosyltransferases
Introduction
In mammalian cells, the endoplasmic reticulum (ER) and Golgi are home to
numerous glycosyltransferases that transfer monosaccharides from nucleotide sugars to
proteins, lipids, and many small molecules. While there have been over 230 human
glycosyltransferases identified to date,1 the process by which complex carbohydrate
structures are assembled onto specific targets has yet to be determined in the same detail
as that of the synthesis of nucleic acids and proteins. In order to explain the regulated
synthesis of complex carbohydrate structures, there have been two important and non-
exclusive models proposed, the assembly line model2 and the multiglycosyltransferase
system.3 In the assembly line model, enzymes are arranged in a linear array that
corresponds to the order in which the modifications occur.4 In the
multiglycosyltransferase system, enzymes that are involved in the particular sequence of
a synthesis are proposed to associate together in complexes, thus allowing for the product
of one enzyme to become the substrate for the next enzyme.3
Evidence consistent with the multiglycosyltransferase model has been obtained
for the enzymes involved in ganglioside biosynthesis (Figure 1A). Gangliosides are a
specific class of glycolipid molecules that contain lactosylceramide (LacCer, Galβ1,4-
Glcα-ceramide) as the core structure and contain at least one sialic acid (Sia) residue
within the structure. Figure 1B shows the structure of the GM3 ganglioside. Three sets
of physical interactions among ganglioside-synthesizing glycosyltransferases have been
reported.5-8 The Maccioni group first reported an interaction between GalNAcT &
GalT2,6 followed by an interaction between GalT1, SialT1, and SialT2.7 Independently,
Bierberich et al. showed an interaction occurring between SialT2 and GalNAcT.5 Further
studies showed that expression of SialT2 in CHO cells causes relocalization of GalT1 and
SialT1 from early ER/Golgi compartments to more distal locations (TGN and recycling
endosomes).8 Additional experiments demonstrated that sequence information contained
within the cytoplasmic tails of these enzymes was responsible for their subcellular
localization within the secretory pathway.9
23
Figure 1. Reported associations among glycosyltransferases responsible for ganglioside biosynthesis. (A) Biosynthetic pathway of gangliosides in mammalian cells. Gangliosides are shown in black font, glycosyltransferases in red font, and reported associations are indicated by the shaded bubbles. Actual gene names for each glycosyltransferases are listed beside pathway figure. (B) GM3 ganglioside. The lactosylceramide (LacCer) core structure is shown in black and the sialic acid residue is shown in red.
Ganglioside glycosyltransferases are type II transmembrane (TM) proteins,
consisting of a N-terminal cytoplasmic tail, single-pass TM domain, stem region and C-
terminal catalytic domain. To better understand the nature of the physical association,
the Maccioni group used truncated sequences of these glycosyltransferases and showed
that the N-terminal domains containing the cytoplasmic tail, the TM domain and all or
part of the stem region were sufficient to mediate association.6-9 Upon further review of
the amino acid sequence of these proteins and considering there are numerous proteins in
nature that associate directly via TM interactions,10,11 I began to speculate that the TM
domain might be responsible for the observed association.
24
Thus, I sought to characterize the molecular basis for these observed physical
interactions, focusing primarily on the TM domains of the enzymes using in vitro assays.
In my efforts to investigate hetero-oligomerization of these enzymes, I discovered that
several of the TM regions formed homo-oligomers in various multiplicities. I then
identified the putative key amino acids that are responsible for homo-oligomerization. In
order to understand the biological relevance of these self-associating TM domains, I
expressed full-length glycosyltransferases in CHO cells and observed their behavior
using an ER retention assay published by our research group12 and by Western blot
analysis. My preliminary results indicate that several ganglioside glycosyltransferases
are able to self-associate inside of cells. Formation of these complexes may be
responsible for the correct localization and production of cellular gangliosides.
Results
In vitro assay to investigate TM interactions
To investigate whether ganglioside glycosyltransferase TM domains can mediate
oligomerization, I used hydropathy analysis13 to identify the putative TM domains of the
five ganglioside glycosyltransferases reported as interaction partners (Figure 2). Using
this information, I decided to use an established dimerization assay that tests the ability of
hydrophobic TM domains to associate within detergent (sodium dodecyl sulfate - SDS)
micelles.14 Each TM domain was expressed as a fusion protein with staphylococcal
nuclease (SN), a non-interacting globular protein (named SN-TM). The use of SN
facilitates the expression and purification of the hydrophobic TM domains (described in
Methods). The purified proteins were then analyzed by SDS-PAGE to determine
whether they associate in the detergent environment. The SDS micelles provide a
suitable mimic of a biological membrane, thus SN-TM fusion proteins that form dimers
in cellular membranes typically migrate as dimers on the SDS gel. The biologically-
relevant dimerization of the TM region of glycophorin A (GpA) has been observed using
this method. An SN-TM construct containing the GpA TM region was used as a positive
control for dimer formation. All purification steps and SDS-PAGE analyses were
performed in the presence of dithiothreitol (DTT) to prevent spurious formation of
intramolecular disulfide bonds between the TM domains.
25
Figure 2. N-terminal sequences of ganglioside glycosyltransferases implicated in their association. Putative transmembrane domains are shaded in pink.
Investigating transmembrane interactions in vitro
The individual SN-TM fusions were analyzed by SDS-PAGE (Figure 3). Both
GalNAcT (Figure 3A – Lane 1) and GalT2 (Figure 3A – Lane 2) migrated almost
exclusively as monomers. The appearance of higher-order oligomers for GalT2 was
determined to be caused by solvent effects and not due to TM association. In contrast,
GalT1 (Figure 3B – Lane 1) produces two very distinct and similarly intense bands on a
gel, correlating to monomer and dimer molecular weights. Similarly, SialT1 (Figure 3B
– Lane 2) produces two distinct bands on a gel, corresponding to the molecular weights
of a monomer and a dimer. SialT2 (Figure 3B – Lane 3) displays multiple
oligomerization states; their molecular weight suggests that these bands represent a trimer
and pentamer, along with other distinct higher-order oligomers. To prove that these
observed interactions were indeed a product of the SN-TM fusions and not due to
unremoved contaminants, I injected these samples onto a reverse phase-high performance
liquid chromatography (RP-HPLC) instrument, isolated the corresponding SN-TM
fusion, and re-analyzed the sample by SDS-PAGE and saw that the observed associations
were reproduced (data not shown).
To investigate hetero-oligomer formation, equimolar quantities of SN-TM fusion
proteins were mixed together in SDS and analyzed by SDS-PAGE.15 Interestingly, I was
unable to observe the predicted complex formation among different TM domains (Figure
3A – Lane 3, Figure 3B – Lanes 4-6). Modifying several experimental parameters,
including mixing time and temperature, also failed to produce any observable hetero-
oligomers.
26
Figure 3. SDS-PAGE analysis of SN-TM homo- and hetero-oligomerization. Molecular weight markers are shown in the left lane of each gel. GpA is the fusion of SN and the transmembrane domain of glycophorin A, which is know to form a homodimer. (A) The transmembranes of GalNAcT and GalT2 fail to demonstrate any homo-oligomerization or hetero-oligomerization when mixed together. The higher order bands present in GalT2 lanes are due to preparation in organic solvent and were demonstrated to be artifacts. (B) The SN-TM fusions of GalT1 and SialT1 are able to form stable dimers while SialT2 shows multiple oligomerization states, including a trimer. The mixing of these individual samples together fails to show any hetero-oligomerization formation.
The discovery of the self-association with several of the TM domains prompted us
to conduct a more detailed investigation into the molecular basis of these interactions.
Close examination of the amino acid sequences of the TM regions revealed the presence
of a number of polar and charged residues located within the mostly hydrophobic
environment. Several reports have demonstrated that the presence of polar residue motifs
in TM regions facilitates their association.16-20 For example, GalT1 contains a sequence
of amino acids (SxxSSxxY) that bears a striking resemblance to a TM dimerization motif
(SxxSSxxT) discovered in a selection experiment.16 Additionally, the spacing of polar
residues three amino acids apart, as seen in the TM domains of GalT1, SialT1 and SialT2,
has been shown to direct TM association.18 Polar residues within the TM domains of
several glycosyltransferases have been suggested to play important roles in dimer
formation and localization (e.g. β1,4-galactosyltransferase,21,22 α2,6-sialyltransferase I,23
and α1,3/4-fucosyltransferase III24). Considering that TM sequences of these ganglioside
27
glycosyltransferases likely form α-helical structures, the arrangement of polar residues
three amino acids apart presents them on the same face of the helix, allowing both
residues to be in simultaneous contact with a neighboring α-helix.
To test whether a similar mechanism is at work in the TM domain of GalT1, I
performed conservative mutations of each of the polar residues to a nonpolar analog
(serine and cysteine to alanine, tyrosine to phenylalanine) of the SN-TM construct and
investigated their effects by SDS-PAGE (Figure 4A). Additionally, I quantified the
relative abundance of monomer and dimers of each mutant (Figure 4B). Mutating
individual amino acids within the TM sequence showed only modest decreases in dimer
formation. Interestingly, the C28A and Y30F mutations actually showed slight increases
in dimer formation. The introduction of two separate double mutations, S23A/S26A or
S25A/C28A, produced dramatic decreases in dimer formation. This result suggests that
the disruption of three amino acid-spaced motifs can modulate the dimer formation of
GalT1. To my surprise, the double mutation of S27A/Y30F actually increased the dimer
formation, corresponding with my previous results from the single mutation Y30F.
These results suggest that the dimer formation of GalT1’s TM domain is largely due to
the presence of properly spaced polar residues.
Figure 4. SDS-PAGE analysis of SN-GalT1 TM WT and mutated fusion proteins. (A) Single and double mutations of GalT1’s TM domain cause mild to significant disruption of the homodimer. Molecular weight markers are shown in the left lane. GpA is the fusion of SN and the transmembrane domain of glycophorin A, which is know to form a homodimer. (B) Quantitative analysis of the relative ratio of monomer to dimer band pixel intensity. Color coding is used to classify mutations as slight-to-moderate dimer disruption (yellow), large dimer disruption (green) or slight increase in dimer formation (red). GalT1 TM WT is highlighted in black and GpA TM (known homodimer) is highlighted in gray.
28
I next explored how similar mutagenesis studies might affect SialT2’s ability to
self-oligomerize into trimers and higher order complexes. The TM domain of SialT2
contains four polar residues that form two pairs, each spaced three amino acids apart.
Conservative mutations of each amino acid of interest were performed as described above
and the resulting proteins were analyzed by SDS-PAGE (Figure 5A). Additionally, I
quantified the percentages of monomer and oligomer species for each mutant to further
analyze their effects (Figure 5B). The single mutation of S18A, C26A or Y29F resulted
in a modest decrease in the fraction of higher order species while the mutation of C21A
produced a dramatic decrease in oligomer formation. When double mutations were
installed, the S18A/C21A species showed a significant diminution of all higher-order
species, including the trimer. This result indicates that the self-association of SialT2’s
TM domain is governed by the cooperative action of S18 and C21. The C26A/Y29F
mutation only showed moderate destabilizing effects, similar to those observed for the
corresponding single mutants.
Figure 5. SDS-PAGE analysis of SN-SialT2 TM WT and mutated fusion proteins. (A) Single and double mutations of SialT2’s TM domain cause mild to significant disruption of the higher order oligomer formation. Molecular weight markers are shown in the left lane. (B) Quantitative analysis of the percentages of monomer and higher order band pixel intensity. Color coding is used to identify the percentage of SialT2 that forms a monomer (green), trimer (blue), pentamer (red), and higher order oligomers (yellow).
The molecular basis for self-oligomerization of SialT1 was not investigated using
the SDS-PAGE method. However, SN-TM constructs of SialT1 containing single
mutations of polar and charged residues in the TM domain were generated. Additionally,
29
I attempted to synthesize peptides of the TM sequences of GalT1, SialT1 and SialT2 but
was unable to efficiently purify them for further study.
Functional analysis of ganglioside glycosyltransferases
To investigate if the in vitro observations of TM oligomerization reflect
interactions that also occur in a cellular environment, I decided to utilize an established
subcellular relocalization assay that measures the physical interaction between two
proteins.25,26 In this assay, the full-length glycosyltransferase is cloned with one of two
different affinity tags (either myc or HA) at the C-terminus to facilitate
immunofluorescence detection of its subcellular location. One form of the enzyme is
fused with the KDEL retention signal (SEKDEL) at the C-terminus causing the protein to
be recognized by the KDEL receptor in the Golgi and become retrograde transported
back into the ER. Both the KDEL-tagged and non-KDEL tagged enzymes are then co-
transfected into CHO cells and localization is determined by immunofluorescence
microscopy. If the glycosyltransferases physically associate, the non-KDEL tagged
protein will become relocalized into the ER (Figure 6).
30
Figure 6. KDEL recruitment assay. Full length glycosyltransferases (depicted in red and purple) were expressed with different C-terminal affinity tags. One form of the enzyme also contains a KDEL retention signal (shown in yellow). Both proteins are synthesized in the ER and trafficked to the Golgi. The KDEL receptor recognizes and retrieves proteins displaying the retention signal back into the ER. If the untagged enzyme interacts with the KDEL-tagged protein, it will also be retrograde trafficked to the ER. If these two proteins do not associate, the untagged enzyme will remain in the Golgi.
Mammalian expression plasmids encoding GalT1, SialT1 and SialT2 were
generated as fusion proteins with the myc or HA affinity tags and with or without the
KDEL retention signal (described in Methods). The non-KDEL-tagged version of all
three enzymes displayed normal Golgi localization when transfected in CHO cells.
Introduction of the KDEL retention sequence onto the C-terminus efficiently relocalized
GalT1, SialT1, and SialT2 into the ER (Figure 7), thus demonstrating that these
ganglioside glycosyltransferases can be used with the KDEL recruitment assay.
To determine if GalT1, SialT1, and SialT2 are able to form self-oligomerizing
complexes inside of cells, I co-transfected CHO cells with a non-KDEL-tagged
31
glycosyltransferase and a KDEL-tagged glycosyltransferase and monitored the location
of the non-KDEL-tagged glycosyltransferase by immunofluorescence microscopy. When
GalT1-HA and GalT1-myc-KDEL were co-expressed, I observed that GalT1-HA
remained strictly in the Golgi and was not recruited into the ER (Figure 7A). When
SialT1-HA and SialT1-myc-KDEL were co-expressed, I observed that SialT1-HA was
recruited into the ER in ~50% of the cells expressing both enzymes (Figure 7B). When
SialT2-HA and SialT2-myc-KDEL were co-expressed, I observed that SialT2-HA was
recruited into the ER in ~90% of the cells expressing both enzymes (Figure 7C). These
results indicate that both SialT1 and SialT2 are likely oligomerizing inside of cells.
Next, I tested the utility of the KDEL-recruitment assay to view the reported
physical associations occurring between GalT1, SialT1, and SialT2 in CHO cells (Figure
7D).7,8 When GalT1-HA and SialT1-myc-KDEL were co-expressed, I observed that
GalT1-HA was localized in the Golgi and not recruited into the ER (Figure 7D). The
CHO cell line contains endogenous levels of GalT1 and SialT1 but is lacking the
presence of SialT2.7 Previous work has demonstrated that the complex formation of these
three enzymes is dependent on the expression of SialT2.8 Without the presence of SialT2
in this experiment, it is very unlikely that SialT1 alone can recruit GalT1. When Sial1-
HA and SialT2-myc-KDEL were co-expressed, I observed that SialT1-HA was being
recruited into the ER in most cells expressing both enzymes (Figure 7D). In this
experiment, all three enzymes are being expressed in CHO cells and I am able to observe
a physical association occurring between the two tagged glycosyltransferases. These
results suggest that the KDEL recruitment assay can show physical associations
occurring between ganglioside glycosyltransferases.
32
Figure 7. Cellular association of ganglioside glycosyltransferases visualized through the KDEL recruitment assay. Addition of the KDEL retention signal causes ganglioside glycosyltransferases to re-localize to the ER. Other glycosyltransferases that co-localize with the KDEL-tagged enzyme will also be recruited to the ER. Localization is assessed by immunofluorescence microscopy and comparison to known organelle markers. (A) GalT1 does not form strong homo-oligomerization. (B) SialT1 shows small levels of recruitment. (C) SialT2 shows recruitment. (D) GalT1 and SialT1 are not interacting with each other in CHO cells that do not contain endogenous SialT2. When SialT2 is co-expressed with SialT1, SialT1 gets recruited to ER.
Since the SialT2 enzyme displayed the most robust level of recruitment with the
KDEL assay, I sought to obtain further evidence of SialT2 homo-oligomerization
occurring within cells. Using the SialT2-HA clone, I transfected CHO cells and
investigated its possible oligomerization. Using SDS-PAGE and western blot (WB)
techniques, I saw the appearance of several higher-order bands occurring with SialT2-HA
(Figure 8A). When the samples were subjected to chemical crosslinking with
formaldehyde, the levels of both monomer and potential higher-order bands decreased.
This suggests that SialT2-HA became highly crosslinked to many other species and the
33
subsequent complex that was formed was insoluble. When I utilized
dithiobis(succinimidyl propionate) (DSP) to perform crosslinking experiments, I
observed significant higher-order oligomerization occurring, corresponding to masses
over 250 kDa. Unfortunately, I was unable to separate out these bands and identify their
exact composition. Since SDS-PAGE can be prone to the dissociation of transmembrane
proteins, I examined the lysates of SialT2-HA transfected CHO cells with perfluoro-
octanoic acid-PAGE (PFO-PAGE). This method has proven to be effective for analyzing
transmembrane proteins in their native quaternary structure.27,28 Using this technique, I
was again able to observe homo-oligomerization of SialT2 (Figure 8B). Since the KDEL
recruitment assay demonstrated a potential interaction occurring between two different
forms of the SialT2 enzyme, I tested whether co-transfection of SialT2-HA and SialT2-
GFP would reproduce these results. While I observed the presence of potential higher-
order oligomers of SialT2-GFP (and SialT2-YFP), I did not see any evidence of a
complex between SialT2-HA and SialT2-GFP by WB analysis (Figure 8C). My data
point toward the formation of a stable SialT2 self-oligomerizing product but further
analysis is required to determine its molecular basis.
34
Figure 8. Full-length SialT2 enzyme shows homo-oligomerization. Protein extracts from CHO cells transiently transfected with SialT2-HA and/or SialT2-GFP/YFP were separated by SDS-PAGE or PFO-PAGE (when noted) and analyzed by Western Blot, as described in methods. (A) SialT2-HA (MW ~ 50 kDa) shows several bands potentially corresponding to self-oligomerization. Treatment of samples with formaldehyde to induce crosslinking of SialT2-HA decreases both monomer band and all higher order bands. Treatment of samples with DSP to induce crosslinking of SialT2-HA also decreases monomer band and generates high molecular weight species. (B) PFO-PAGE analysis of SialT2-HA self-oligomerization shows potential higher-order species. (C) Transfection of SialT2-GFP produces several higher-order species. Co-transfection of SialT2-HA with SialT2-GFP does not show the production of any new higher order species.
Discussion
The TM domains of ganglioside glycosyltransferases fail to demonstrate the formation of
hetero-oligomeric species with the SN-TM assay
My initial investigations were focused toward defining the role that TM domains
play in the formation of several reported enzyme complexes of ganglioside
glycosyltransferases.5-8 My studies were performed using an established dimerization
assay that has been reported to accurately display biologically relevant TM domain
interactions.14 While this assay has proved to be robust for analyzing homo-oligomer
35
formation of TM domains,16,18-20 I did not detect any hetero-oligomer formation between
the individual TM sequences (Figure 3). My attempts to optimize the experiment did not
increase the presentation of hetero-oligomers. The investigation of these interactions
may require more sophisticated biochemical methods, such as analytical ultra
centrifugation (AUC) or multiple angle laser light scattering (MALLS).
The potential of this assay to find these interactions may be hindered for several
reasons. First, the purified SN-TM proteins may be segregated into individual SDS
micelles that are unable to mix with one another on the time scale of my experiment.
Attempting to co-express two or more SN-TM constructs at the same time might be a
possible technique to alleviate this problem. Second, and more importantly, while I am
focusing directly on the TM domain, the initial studies investigating physical associations
among ganglioside glycosyltransferases either used full-length enzymes5 or N-terminal
fragments containing the cytoplasmic tail and part of the stem region.6-8 The cytoplasmic
tails of glycosyltransferases have shown to control their localization within the secretory
pathway,29 including those involved with ganglioside biosynthesis.7,9,30 The amino acid
composition of cytoplasmic tails is also known to be a critical recognition element for
several chaperones of ganglioside glycosyltransferases, including calcium-binding
proteins31 and components of COPII vesicles.32 Stem regions have also been shown to
play critical roles in heterodimer formation of glycosyltransferases.33 The absence of
these elements in my in vitro method may hinder the observation of these enzyme
complexes, requiring more cell-based assays.
The TM domains of several ganglioside glycosyltransferases mediate formation of homo-
oligomers
While I was unable to observe hetero-oligomer formation between the TM
domains of reported ganglioside glycosyltransferase complexes, several of these enzymes
displayed the ability to form homo-oligomer complexes with the SN-TM assay: GalT1
and SialT1 formed stable dimeric complexes and SialT2 formed several higher-order
complexes (Figure 3). When I further investigated the molecular basis for GalT1
dimerization, I found that their associations were due to the presence of properly space
polar amino acids within the hydrophobic sequence; both double mutants S23A/S26A
36
and S25A/C28A showed significant diminution of the observed dimer (Figure 4). This
observation agrees with the report that spacing polar residues three amino acids apart,
thus putting them directly onto the same face of an α-helix, can mediate interactions
between TM domains.18 While the TM domain of GalT1 posses a sequence of polar
amino acids (S23XXS26S27XXY30) similar to a reported dimerization motif
(SXXSSXXT),16 performing the double mutation S27A/Y30F caused GalT1 to form a
slightly stronger dimer than the WT form (Figure 4). This suggests that Y30 is not
involved in the dimerization of GalT1 and potentially inhibits the ability to form a dimer.
When I further investigated the molecular basis for SialT2 oligomerization, I saw
that all four of the polar residues contributed to the observed self-association, with one
residue (C21) being most important (Figure 5). Introduction of the double mutant
S18A/C21A was found to be even more detrimental to SialT2 oligomerization. Cysteine
residues within TM domains have been shown to be critical for the formation of dimeric
species of several glycosyltransferases.21-24 The ability of these cysteines to form
disulfide bonds between enzymes has been shown to be essential for proper cellular
localization and glycan product synthesis. Further work will be needed to determine if
these cysteine residues are critical for these roles.
The ability of hydrophilic amino acids within TM domains to mediate the
formation of homo-oligomers has been well demonstrated and is consistent with my
results. Additionally, there are other biophysical methods that have shown value towards
understanding TM domain interactions in more biologically relevant situations, including
the TOXCAT34 and GALLEX35 assays, and would be suggested as further methods to
fully characterize both homo- and hetero-oligomerization. Overall, I feel confident that
the results seen with GalT1, SialT1, and SialT2 provide evidence that the TM domains
engage as homo-oligomers in SDS micelles. However, these results do not provide
information about oligomerization in a more native membrane context.
37
SialT1 and SialT2 enzymes are physically associated in vivo
To further investigate the potential homo-oligomeric complexes of ganglioside
glycosyltransferases observed by the in vitro method, I utilized an established KDEL-
recruitment assay (illustrated in Figure 6) that has been successful in identifying physical
associations between enzymes involved in N-linked glycan25,26 and poly-N-
acetyllactosamine synthesis.12 To confirm the ability of this assay to function with
ganglioside glycosyltransferases, I demonstrated that SialT2-myc-KDEL can recruit
SialT1-HA into the ER, confirming previous observations with these two enzymes.7,8
When I investigated homo-oligomer formation, I observed several of the ganglioside
glycosyltransferases were able to physically associate together inside of cells: SialT1
demonstrated self-association in ~50% of cells co-expressing both constructs (Figure 7B)
while SialT2 had a much higher level of self-association, being present in ~90% of cells
successfully co-expressing both constructs (Figure 7C). I did not observe any KDEL-
dependent relocalization occurring with transfected GalT1 constructs (Figure 7A). While
these results do not correlate with those found in the SDS-PAGE assay, it is plausible that
the observed TM dimer of GalT1 may be much easier to reproduce within the SN-TM
assay since I am excluding the contributions of all other regions of the enzyme.
However, since it has been shown that the N-terminal regions of ganglioside
glycosyltransferases are able to facilitate hetero-oligomer formation,6-8 I believe that their
TM domains play an undetermined role in facilitating the observed self-oligomerization
interactions observed with SialT1 and SialT2.
SialT2 demonstrates the formation of higher-order complexes in cells
When SialT2-HA was transfected into CHO cells, I was able to observe the
formation of homo-oligomeric species, including the formation of a dimer, by SDS-
PAGE (Figure 7A, C) and PFO-PAGE (Figure 7B). These results were obtained without
the use of crosslinking reagents and performed under reducing conditions (DTT). SialT2
has been observed by other groups to form a dimeric species in transfected CHO-K136
and F-11A5 cells but only under non-reducing conditions. My observations suggest that
oligomerization is not entirely controlled by disulfide bond formation since all
experiments were performed under reducing conditions. My attempts to capture these
38
complexes with common chemical crosslinking reagents provided mixed results: the use
of formaldehyde caused an overall decrease in signal without increased formation of high
molecular weight complexes, while DSP generated large molecular weight complexes
(greater than 250 kDa) that could not be resolved by SDS-PAGE (Figure 7A). These
reagents are likely causing SialT2 to form complexes with other cellular proteins and
molecules that may or may not be physiologically relevant to their actual
oligomerization. The use of crosslinking reagents that are able to trap TM interactions
directly would prove to be more relevant for these studies. One such possibility would be
to introduce photoactivatable amino acids into the transmembrane domain, such as photo-
leucine and photo-methionine, which have been used to capture protein-protein
interactions in living cells.37 Additionally, it would be very interesting to see if the SialT2
TM domain mutations, primarily S18A and C21A, would disrupt the self-association of
full length SialT2 observed in the KDEL recruitment assay and SDS-PAGE/WB
analyses.
In conclusion, I present preliminary evidence that several TM domains of
ganglioside glycosyltransferases are able to self-associate into oligomeric species both in
vitro and inside of cells. The primary molecular basis for these interactions is dependent
upon the presence of properly spaced hydrophilic amino acids. Future work is needed to
investigate the functional consequences of these associations within a cellular
environment, including effects on subcellular localization and ganglioside product
formation.
Acknowledgements
I would like to thank Don Engelman, Jennifer Czlapinski, and Suzanne Pfeffer for
their reagent gifts. I would like to thank Marguerite M. Desko and Peter L. Lee for help
helpful suggestions on microscopy and WB analyses. I would like to thank Steve Kaiser
for helpful advice and assistance with MALDI-TOF-MS analysis. I would like to thank
Kitty Lee, Cell Sciences Imaging Facility at Stanford University, for assistance with
confocal microscopy.
39
Methods
Reagents
All chemicals and reagents were purchased from Fisher Scientific (Waltham,
MA) or Sigma-Aldrich (St. Louis, MO) unless noted. Restriction enzymes, T4 DNA
Ligase, calf intestinal phosphatase were purchased from New England Biolabs (Ipswich,
MA). Mouse anti-myc antibody was purchased from the Development Studies
Hybridoma Bank (Iowa City, IA). Mouse anti-GM130 was purchased from BD
Biosciences (San Jose, CA). Mouse anti-p115 was as a generous gift of Suzanne Pfeffer
(Stanford University School of Medicine, Stanford, CA). Rabbit anti-giantin was
purchased from Covance (Princeton, NJ). Rabbit anti-calreticulin was purchased from
Stressgen (Ann Arbor, MI). Rabbit anti-HA was purchased from Abcam (Cambridge,
MA). Rat anti-HA was purchased from Roche Applied Science (Indianapolis, IN). Goat
anti-calnexin was purchased from Santa Cruz Biotechnology Inc (Santa Cruz, CA). Goat
anti-rat HRP, Goat anti-mouse Alexa 488, Goat anti-mouse Alexa 610, Goat anti-rabbit
Alexa 488, Goat anti-rabbit Alexa 610, and Goat anti-donkey Alexa 488 were purchased
from Invitrogen (Carlsbad, CA). Vectashield Mounting Medium with DAPI was
purchased from Vector Labs (Burlingame, CA).
Cloning of SN-TM constructs
pT7SN/linker and pT7SN/GpA-TM were gifts of Don Engelman (Yale
University, New Haven, CT). pT7SN/GalNAcT-TM and pT7SN/GalT2 were gifts of
Jennifer Kohler (UT Southwestern Medical Center, Dallas, TX). pCR-Blunt II-
TOPO/GalT1-HA and pcDNA3.1-Zeo/SialT2 were gifts of Jennifer Czlapinski
(University of California, Berkeley, CA). All plasmids encoding staphylococcal
nuclease-transmembrane chimeras (SN-TM) were constructed from pT7SN/linker.
Putative transmembrane sequences were identified and selected for cloning using Kyte-
Doolittle hydropathy analysis.13 PCR amplification primers for cloning were acquired
from the Stanford University Protein and Nucleic Acid Facility and are listed in Table 1.
The transmembrane sequence of GalT1 (β4GalT6, amino acids 15-40) was
amplified from the plasmid pCR-Blunt II-TOPO/GalT1-HA by PCR using primers
40
GalT1-TM 5’ and GalT1-TM 3’. The transmembrane sequence of SialT1 (ST3Gal5,
amino acids 6-31) was generated synthetically by PCR first using primers SialT1-TM 1st
Primer 5’ and SialT1-TM 1st Primer 3’, followed by using primers SialT1-TM 2nd Primer
5’ and SialT1-TM 2nd Primer 3’. The transmembrane sequence of SialT2 (ST8Sia1,
amino acids 30-53) was amplified from the plasmid pcDNA3.1-Zeo/SialT2 by PCR using
primers SialT2-TM 5’ and SialT2-TM 3’. The PCR products were cloned into the
plasmid pCR-Blunt II-TOPO using the Zero Blunt TOPO PCR Cloning Kit (Invitrogen),
whose identity was confirmed by sequencing. The inserts were excised using AvrII and
BamH1, then ligated into pT7SN/Linker that had been digested with the same enzymes
and treated with calf intestinal phosphatase. Successful ligation was confirmed by
restriction digest. The plasmids were named SN-GalT1 TM, SN-SialT1 TM and SN
SialT2-TM, respectively.
Table 1. Table of primers used to generate SN-TM plasmids. Primer names and sequences, written from 5’ to 3’, are listed.
Mutation of TM inserts
Site-directed mutagenesis was performed according to the Stratagene QuikChange
Site-Directed Mutagenesis Kit (La Jolla, CA). PCR amplification primers for cloning
were acquired from the Stanford University Protein and Nucleic Acid Facility.
Single mutations on wild-type SN-GalT1 TM were performed using the primers
listed in Table 2. Double mutations to generate S23A/S26A (using SN-GalT1 S23A),
S25A/C28A (using SN-GalT1 C28A), and S27A/Y30F (using SN-GalT1 S27A) were
performed using the primers listed in Table 2.
41
Table 2. Table of primers used to generate SN-GalT1 TM mutants. Primer names and sequences, written from 5’ to 3’, are listed.
Single mutations on wild-type SN-SialT2 TM were performed using the primers
listed in Table 3. Double mutations to generate S18A/C21A (using SN-SialT2 C21A)
and C26A/Y29F (using SialT2-TM C26A) were performed using the primers listed in
Table 3.
Table 3. Table of primers used to generate SN-SialT2 TM mutants. Primer names and sequences, written from 5’ to 3’, are listed.
42
Purification of SN-TM constructs
pT7SN/constructs were transformed into BL21(DE3) competent cells to express
the SN-TM fusion proteins. Individual colonies were taken and grown overnight in LB
media. These cultures were used to inoculate new LB cultures at a dilution of 1:100,
growing at 37 oC. After reaching an OD600 = 0.6, IPTG was added to a final
concentration of 1.0 mM and grown for two additional hours. Cells were harvested by
centrifugation for 20 min at 5,000 g, 4 oC. Cells were resuspended in 1/20 culture
volume of 50 mM Tris-HCl (pH 8.0), 5 mM EDTA (pH = 8.0), 1 mM PMSF, 1 mM
DTT. Cells were lysed with three rounds of freeze-thaw cycles. The lysate was clarified
by centrifugation for 45 min @ 20,000 g, 4 oC. To remove unwanted contaminants, the
remaining pellet was then washed first with a solution of 25 mM Tris-base (pH = 8.0), 1
M NaCl, 1 mM EDTA, 1 mM PMSF, 1 mM DTT followed by 25 mM Tris-base (pH =
8.0), 25 mM NaCl, 2 % Thesit, 1 mM EDTA, 1 mM PMSF. The pellet was washed for
three hours with each solution and clarified by centrifugation for 45 min @ 20,000 g, 4 oC. SN-TM chimeras were extracted using a solution of 25 mM Tris-HCl, 1 M NaCl, 1
mM EDTA, 1 mM PMSF, 1 mM DTT, 4.4 M urea and clarified by centrifugation. The
remaining supernatant was concentrated using an Amicon Ultra-15 centrifugal device,
5,000 MWCO (Millipore, Billerica, MA) and the buffer was exchanged with the
following of 25 mM Tris-base (pH = 8.0), 200 mM NaCl, 1 mM EDTA, 1 mM PMSF, 1
mM DTT.
Further purification was achieved using reverse phase-HPLC. Samples were
filtered with a 0.22 µM syringe filter and injected onto an Alltech Apollo C18u, 250 x 4.6
mm column. Samples were eluted using a gradient of 0 – 75 % Buffer B at a flow rate of
1.00 mL/min with Buffer A as 99.9% H2O, 0.1% TFA and Buffer B as 99.9%
acetonitrile, 0.1% TFA. Collected peaks were verified by MALDI-TOF-MS analysis
before rotary evaporation of solvent in Savant SpeedVac (ThermoFisher Scientific,
Waltham, MA). Expected masses of SN-TM constructs: GalT1 – 21383.67 Da, SialT1 –
21588.06, SialT2 – 21207.63, GalT2 – 21130.51, GalNAcT – 21113.34. All SN-TM
constructs (WT and mutants) displayed masses within 1% of expected mass. Dried
protein samples were redissolved using 25 mM Tris-base (pH = 8.0), 200 mM NaCl, 1
43
mM EDTA, 1 mM PMSF, 1 mM DTT, 2.0% Thesit. Sample concentrations were
determined using the BCA® Protein Assay Kit (Pierce, Rockford, IL).
SDS-PAGE analysis of SN-TM oligomerization
Purified SN-TM samples were mixed 1:1 with 2x SDS-PAGE loading buffer (100
mM Tris-base [pH = 6.8], 20 % glycerol, 4 % SDS, 0.2 % bromophenol blue), boiled for
10 min and separated using 12 % Tris-HCl Ready Gels (Bio-Rad, Hercules, CA). For
hetero-oligomerization experiments, samples were mixed together for 30 minutes at room
temperature before loading. Molecular weights were confirmed using Benchmark
Protein Ladder (Invitrogen). Gels were stained with either Coomassie Brilliant Blue (45
% MeOH, 45% ddH2O, 10% glacial acetic acid, 0.0025% Coomassie Brilliant Blue
R250) or SimplyBlue SafeStain (Invitrogen). Imaging of gels was achieved using an
Alpha Innotech FluorChem HD2 (Santa Clara, CA) and quantification of bands was
performed using ImageJ software (NIH, Bethesda, MD).
Cloning of full length ganglioside glycosyltransferases genes
pcDNA3.1(+)/GalT1-HA, pcDNA3.1(+)/SialT1-myc, and pcDNA3.1-Zeo/SialT2
and were generous gifts of Jennifer Czlapinski (University of California, Berkeley, CA).
All GalT1, SialT1, and SialT2 inserts were cloned with a C-terminal affinity tag (myc or
HA) and with a KDEL tag (when applicable), followed by a stop codon. The amino acid
sequences for the C-terminal epitope tags are: myc tag – EQKLISEEDL, HA tag –
YPYDVPDYA, KDEL tag – SEKDEL. PCR amplification primers for cloning were
acquired from the Stanford University Protein and Nucleic Acid Facility and are listed in
Table 4.
GalT1-myc was produced by PCR from the plasmid pCR-Blunt II-TOPO/GalT1-
HA as a template using the primers GalT1-myc 5’ and GalT1-myc 3’. GalT1-myc-
KDEL was produced by PCR from the plasmid pcDNA3.1(+)/GalT1-myc as a template
using primers GalT1-myc and myc-KDEL 3’. GalT1-HA-KDEL was produced by PCR
from the plasmid pCR-Blunt II-TOPO/GalT1-HA as a template using primers GalT1-myc
5’ and GalT1-HA-KDEL 3’. The PCR products were cloned into the plasmid
pCR4Blunt-TOPO using the Zero Blunt TOPO PCR Cloning Kit (Invitrogen), whose
44
identity was confirmed by sequencing. The inserts were excised using NheI and EcoRI,
then ligated into pcDNA3.1-Zeo that had been digested with the same enzymes and
treated with calf intestinal phosphatase. Plasmids containing the inserts were identified
by restriction digest. The plasmids were named GalT1-myc, GalT1-myc-KDEL, and
GalT1-HA-KDEL, respectively.
SialT1-HA was produced by PCR from the plasmid pcDNA3.1(+)/SialT1-myc as
a template using primers SialT1-HA 5’ and SialT1-HA 3’. SialT1-myc-KDEL was
produced by PCR from the plasmid pcDNA3.1(+)/SialT1-myc as a template using
primers SialT1-HA 5’ and myc-KDEL 3’. SialT1-HA-KDEL was produced by PCR
from the plasmid pcDNA3.1(+)/SialT1-HA as a template using primers SialT1-HA 5’
and SialT1-HA-KDEL 3’. The PCR products were cloned into the plasmid pCR4Blunt-
TOPO using the Zero Blunt TOPO PCR Cloning Kit (Invitrogen), and their identity was
confirmed by sequencing. The inserts were excised using NheI and EcoRI, then ligated
into pcDNA3.1-Zeo that had been digested with the same enzymes and treated with calf
intestinal phosphatase. Positive ligation was tested by restriction digest. The plasmids
were named SialT1-HA, SialT1-myc-KDEL, and SialT1-HA-KDEL, respectively.
SialT2-myc was produced by PCR from the plasmid pcDNA3.1-Zeo/SialT2 as a
template using primers SialT2-myc/HA 5’ and SialT2-myc 3’. SialT2-HA was produced
by PCR from the plasmid pcDNA3.1-Zeo/SialT2 as a template using primers SialT2-
myc/HA 5’ and SialT2-HA 3’. SialT2-myc-KDEL was produced by PCR from the
plasmid pcDNA3.1/SialT2-myc as a template using primers SialT2-myc/HA 5’ and
SialT2-myc-KDEL 3’. SialT2-HA-KDEL was produced by PCR from the plasmid
pcDNA3.1/SialT2-HA as a template using primers SialT2-myc/HA 5’ and SialT2-HA-
KDEL 3’. The PCR products were cloned into the plasmid pCR4Blunt-TOPO using the
Zero Blunt TOPO PCR Cloning Kit (Invitrogen) and confirmed by sequencing. The
inserts were excised using NheI and EcoRV, then ligated into pcDNA3.1-Zeo that had
been digested with the same enzymes and treated with calf intestinal phosphatase.
Positive ligation was tested by restriction digest. The plasmids were named SialT2-myc,
SialT2-HA, SialT2-myc-KDEL and SialT2-HA-KDEL, respectively.
45
Table 4. Table of primers used to generate KDEL recruitment assay plasmids. Primer names and sequences, written from 5’ to 3’, are listed.
Immunofluorescence analysis
One day prior to transfection, CHO cells were seeded at a density of 1 x 105
cells/mL in individual wells of a 6-well plate containing cover slips (10 cm2 surface
area/well). Cells were transfected using Lipofectamine 2000 Reagent (Invitrogen)
according to the manufacturers directions, using Opti-MEM (Invitrogen) as media during
the transfection. After exposure to transfection for four hours, fresh media was added
and the cells were cultured for 48 hours before harvesting.
Cells were fixed with 3.7% paraformaldehyde/PBS for 20 minutes without
washing. After several washes with PBS, the cells were permeabilized with 0.1% Triton
X-100, 1.0% BSA/PBS for 5 minutes without shaking. After several washes with PBS,
the plates were placed at RT and incubated with 1.0% BSA/PBS for 1 hour to block non-
specific binding. After blocking, the cells were incubated with primary antibody (1:500
dilution) for 2 hours at RT. The coverslips were then washed 3x with PBS and the cells
were incubated with secondary antibody (1:500 dilution) for 1 hour at RT. The
coverslips were washed again 3x with PBS. After washing, the coverslips were mounted
onto glass cover slides using Vectashield Mounting Medium with DAPI and sealed with
nail polish. Slides were stored at 4 oC until analysis. Cells were visualized using the
HCX PL APO 63x 1.32-0.60 oil objective of a Leica SP2 AOBS Confocal Laser
Scanning Microscopy equipped with 405 nm, 488 nm, and 594 nm lasers. Image analysis
was performed using Adobe Photoshop and Adobe Illustrator.
46
Western Blot Analyses of SialT2
One day prior to transfection, CHO cells were seeded at a density of 1 x 105
cells/mL in 5 mL of media in 6 cm dishes. Cells were transfected using Lipofectamine
2000 Reagent according to the manufacturers directions, using Opti-MEM as media
during the transfection. After exposure to transfection for four hours, fresh media was
added and the cells were cultured for 48 hours before harvesting. Cells were washed
three times with cold PBS and lysed using 0.5 mL of lysis buffer (50 mM Tris, pH = 7.2,
300 mM NaCl, 1.0 % Triton X-100, 1 mM DTT, 1 mM PMSF) with protease inhibitor
cocktail (antipain, turkey trypsin inhibitor, leupeptin, aprotinin) added for 30 min at 4 oC.
Formaldehyde crosslinking studies were performed directly after the 48 hour
incubation period. Cells were washed 3x with cold PBS and incubate with 0.5 % - 1.0 %
formaldehyde solution (in ddH2O) for 10 min at room temperature. The reaction was
quenched with 0.75 M glycine, incubating for an additional 10 min at room temperature.
Cells were again washed three times with cold PBS and lysed as described above. DSP
crosslinking studies were performed on harvested cell lysates. DSP (Pierce) was added at
0.0 mM, 0.12 mM, 0.25 mM, 0.62 mM and 1.24 mM concentrations to 30 µL aliquots of
lysates and incubated for 20 minutes at room temperature. The reaction was quenched
with 0.75 M glycine, incubating for an additional 10 min at room temperature before
analysis by SDS-PAGE.
For SDS-PAGE analysis, samples were mixed with an equal volume of 2x SDS-
PAGE loading buffer (100 mM Tris-base, pH = 6.8, 20 % glycerol, 4 % SDS, 0.2 %
bromophenol blue), boiled for 10 min at 95 oC and separated using 12 % Tris-HCl Ready
Gels (Bio-Rad, Hercules, CA). PFO-PAGE of SialT2 samples was performed as
described previously.28 Briefly, samples were mixed 1:1 with 2x PFO-PAGE loading
buffer (100 mM Tris-base [pH = 8.0], 20 % glycerol, 2 % PFO [pH = 8.0], 0.005%
bromophenol blue), vortexed briefly, centrifuged for 5 min at 10,000g. 7 % Tris-HCl
Ready Gels were pre-run for 60 min at 50 V (constant voltage), 4 oC before loading
samples. Electrophoresis was performed in the cold room at 140 V (constant voltage) for
2 hours using PFO-PAGE running buffer (25 mM Tris, 192 mM, 0.25% PFO at pH =
8.5), pre-chilled at 4 oC.
47
Proteins were transferred to nitrocellulose membrane and stained briefly with
Ponceau S Staining (0.1 % Ponceau S in 5 % acetic acid) to verify the presence of
transferred proteins. After rinsing with Tris-Buffered Saline with 0.1 % Tween-20
(TBST), the membrane was blocked in 5 % nonfat dry milk in TBST for one hour at
room temperature before incubation with rat anti-HA antibody (1:1250 dilution)
overnight at 4 oC. After rinsing with TBST, the membrane was blocked again with TBST
containing 5 % nonfat dry milk for hour, followed by incubation with goat anti-rat HRP
(1:5000 dilution) for one hour at room temperature. The membrane was developed using
SuperSignal West Pico chemiluminescence substrate (Pierce) and imaged on Pierce CL-
XPosure Film (Pierce). Imaging of gels was achieved using an Alpha Innotech
FluorChem HD2 (Santa Clara, CA).
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12 Lee, P. L., Kohler, J. J. & Pfeffer, S. R. Association of beta-1,3-N-
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18 Sal-Man, N., Gerber, D. & Shai, Y. The identification of a minimal dimerization
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19 Zhou, F. X., Cocco, M. J., Russ, W. P., Brunger, A. T. & Engelman, D. M.
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52
Chapter 3 – Covalently capturing the ganglioside GM1-cholera toxin complex with
photocrosslinking
Introduction
Cell surface glycans are important recognition factors for a large number of cell-
cell and cell-ligand interactions. Many of these binding events are regulated by the
presence or absence of sialic acid.1,2 Sialic acids are a family of nine carbon α-keto acids
that are found terminally attached to glycoproteins and glycosphingolipids.3 While these
molecules serve important regulatory roles for mammalian cells, they are also utilized as
receptors for invasion by many pathogens. For example, influenza viruses utilize
hemagglutinin to bind to cell surface sialic acid.4 Sialylated glycosphingolipids, known as
gangliosides, are the principal recognition elements for the invasion of several viral
pathogens, such as BK,4 SV40,5 and polyoma virus.5 They are also targets of many toxins
produced by bacterial pathogens, including tetanus toxin,6 E. coli heat-labile
enterotoxins,7 and cholera toxin.8
Explicitly determining ganglioside interaction partners and their molecular detail
remains a challenge as most carbohydrate-mediated interactions are characterized by low
affinities and fast off rates. These properties make traditional purification techniques
impractical, as the unstable complex cannot withstand the washing steps. To assist in
capturing these labile complexes, there is a need to develop new photocrosslinking tools
that capture these complexes in their native states. One conceivable solution to this
problem is to introduce photoreactive groups directly onto the glycan structure using
metabolic oligosaccharide engineering techniques. Metabolic oligosaccharide
engineering methodology has been highly successful at introducing a wide range of
chemical modifications into several monosaccharides, including sialic acid.9 Our group
has recently demonstrated the successful incorporation of a diazirine moiety into the cell
surface sialic acid binding lectin CD22 and used this crosslinker to covalently capture
CD22 homo-oligomers.10 In this case, the modification of sialic acid was achieved by
culturing mammalian cells with a diazirine containing N-acetylmannosamine analog
(ManNDAz – Figure 1), which is metabolized by cells and displayed in sialosides as
SiaDAz. The diazirine was selected instead of other crosslinkers because of its small
53
size, long UV activation wavelength, high reactivity, and non-selectivity.11 If positioned
appropriately, the diazirine will introduce a covalent bond between sialic acid and its
interaction partner upon photoirradiation. This covalently linked complex can then be
isolated, purified, and the partners identified.
While our group’s initial studies were able to capture the oligomerization of a
glycoprotein,10 I wanted to investigate the applicability of metabolic oligosaccharide
engineering to install photocrosslinking sialic acids into gangliosides. I chose to use
Jurkat cells for my investigation as they produce several ganglioside structures and
previous work showed their ability to incorporate unnatural sialic acid analogs into cell
surface gangliosides.12 I observed that Ac4ManNDAz-treated Jurkat cells were able to
produce SiaDAz-modified GM2 and GM1 gangliosides. These results demonstrated that
our unnatural analog effectively competed with endogenous sialic acid for addition into
gangliosides. Next, I verified that the modified GM1 structure bearing the
photocrosslinking moiety could be recognized by one of its known-binding partners,
cholera toxin subunit B (CTxB).8 In cooperative experiments performed with Michelle
Bond (UT Southwestern), we have confirmed that GM1-SiaDAz can be successfully
crosslinked to CTxB. To investigate association with GM1 affects CTxB endocytosis
and retrograde trafficking, I performed preliminary experiments exploring the co-
localization of CTxB in several cellular organelles using immunofluorescence
microscopy. My work shows that SiaDAz can efficiently be incorporated into cell
surface gangliosides of Jurkat cells and can be covalently linked to a known binding
partner with high efficiency.
Results
HPTLC analysis of gangliosides produced by Ac4ManNDAz-treated Jurkat cells
To investigate the potential of mammalian cells to incorporate a diazirine-
modified sugar into gangliosides, Jurkat cells were cultured with fully acetylated
ManNDAz (Ac4ManNDAz) for 72 hours. Mammalian cells are able to take up acetylated
ManNAc analogs by passive diffusion, remove the acetyl groups by non-specific
esterases, convert the ManNAc analogs to their sialic acid counterparts, and incorporate
54
the sialic acid analogs into glycoconjugates that are displayed on the cell surface.10,11,13
While ManNAc analogs serve as committed precursors for sialic acid analogs, analogs of
the 2-epimer of ManNAc, GlcNAc, are not efficiently metabolized to their sialic acid
counterparts.13,14 Thus, I chose to use the diazirine-containing analog GlcNDAz (Figure
1) as control molecule that is not expected to be metabolized to SiaDAz. Additionally,
Jurkat cells were cultured with Ac4ManNAc (Figure 1) to show that the exogenous
addition of the normal metabolites in the sialic acid biosynthesis pathway does not affect
ganglioside production. Acetylated sugars were added to the media at a final
concentration of 100 µM. After harvesting the cultured cells, I extracted the gangliosides
using established methods15-17 (described in Methods) and resolved them using high
performance TLC plates (HPTLC). To visualize gangliosides, the HPTLC plates were
stained with resorcinol, which reacts specifically with gangliosides upon heating or
visualized using CTxB immunostaining.
Figure 1. Acetylated analogs used to investigate metabolic oligosaccharide engineering of gangliosides. Jurkat cells were cultured with Ac4ManNDAz or Ac4GlcNDAz to investigate their incorporation into gangliosides. Ac4ManNAc is used as a control molecule to demonstrate that supplementation with exogenous sugar does not affect ganglioside production. Sugars were fully acetylated to improve cellular uptake; acetyl groups are expected to be cleaved inside the cell by non-specific esterases.
Jurkat cells are able to synthesize several different types of gangliosides,
including GM3, GM2, GM1 and GD1a (Figure 2A, B). The appearance of two distinct
bands for each ganglioside species is due the incorporation of different fatty acids into the
ceramide portion of the molecule. When compared to unsupplemented Jurkat cells, the
Ac4ManNDAz-treated cells exhibit significant changes in their ganglioside composition
as evidenced by the mobility changes on the HPTLC plate. Introduction of the diazirine
side chain onto sialic acid increases the hydrophobicity of the ganglioside causing it to
have a higher Rf value. The Ac4ManNDAz-treated cells continue to generate normal
55
gangliosides but at weaker at reduced levels. As expected, culturing Jurkat cells with
Ac4GlcNDAz did not cause any detectable changes in ganglioside production. This
suggests that one or more of the enzymes responsible for metabolizing GlcNDAz into
ManNDAz cannot accept this change to the N-acyl chain. I also observed that the
introduction of exogenous Ac4ManNAc to Jurkat cells does not appear to increase
production of cellular gangliosides.
To confirm the successful modification of Jurkat gangliosides with ManNDAz, I
separated the gangliosides from untreated, Ac4ManNAc-, Ac4ManNDAz-, and
Ac4GlcNDAz-treated Jurkat cells by HPTLC and probed the HPTLC plate for the GM1
ganglioside using a CTxB-Alexa Fluor 488 conjugate (Figure 2C). As a known binding
partner of GM1, CTxB expectedly binds to GM1 in unsupplemented and Ac4ManNAc-
treated Jurkat cell samples. Importantly, CTxB also recognizes additional bands in the
gangliosides from Ac4ManNDAz-treated cells. I hypothesized that these bands
correspond to GM1-SiaDAz. When gangliosides from the Ac4GlcNDAz-treated cells
were examined for binding to CTxB, there was no additional band to indicate the
presence of GM1-SiaDAz gangliosides, further confirming the inability of Jurkat cells to
metabolize GlcNDAz into gangliosides. These results also provide evidence that CTxB
recognition of GM1 is maintained upon installation of the unnatural diazirine side chain
in sialic acid.
56
Figure 2. HPTLC analysis of gangliosides from cultured Jurkat cells. Jurkat cells were cultured for 72 hours untreated or with Ac4ManNAc, Ac4ManNDAz, or Ac4ManNDAz. The cultured cells were harvested and the gangliosides were extracted. The ganglioside composition from each cultured Jurkat sample was resolved by HPTLC and analyzed by resorcinol (B) or CTxB immunostaining (C). (A) Jurkat cells are able to synthesize several different types of gangliosides, including GM3, GM2, GM1 and GD1a. (B) Ac4ManNDAz-treated Jurkat cells display modified ganglioside patterns compared to unsupplemented cells. The modified gangliosides are recognized by their mobility shift on the HPTLC plate caused by the introduction of the SiaDAz molecule. GlcNDAz, the metabolic precursor to ManNDAz, is not metabolized into SiaDAz. Treatment of cells with Ac4ManNAc does not affect ganglioside composition levels. (C) Cholera toxin subunit B is able to recognize GM1-SiaDAz. Cholera toxin subunit B binds GM1 ganglioside and can be used to confirm the presence of GM1. Ac4ManNDAz-treated Jurkat cells display two different sets of GM1 molecules: the top pair representing GM1-SiaDAz and the bottom pair representing endogenous GM1.
57
Mass spectrometry analysis of gangliosides produced by Ac4ManNDAz-treated Jurkat
cells
To confirm the successful modification of Jurkat gangliosides with SiaDAz,
extracted gangliosides were analyzed by mass spectrometry at the Complex Carbohydrate
Research Center (University of Georgia). Samples were crystallized with
trihydroxyacetophenone monohydrate (THAP) matrix and analyzed by MALDI-TOF-MS
in negative ionization mode. As shown in Figure 3, Jurkat cells cultured with
Ac4ManNDAz showed efficient incorporation of SiaDAz into GM2 and GM1
gangliosides. It was also noted that several of the peaks contained SiaDAz modified
structures without the diazirine group present (denoted by stars in Figure 3). The loss of
N2 by this MALDI-TOF-MS was consistent between several analyses (data not shown)
and is believed to occur upon ionization. These results were compared to untreated and
Ac4ManNAc-treated Jurkat samples to determine the endogenous ganglioside
composition (shown in Appendix). As observed in Figure 2B, Ac4GlcNDAz-treated cells
did not show detectable levels of SiaDAz-containing gangliosides (located in Appendix).
These results confirm that Jurkat cells can metabolize Ac4ManNDAz to SiaDAz and
incorporate SiaDAz into gangliosides at significant levels.
58
Figure 3. MALDI-TOF-MS analysis of Jurkat cells cultured with Ac4ManNDAz. Ganglioside extracts of Jurkat cells cultured with Ac4ManNDAz were crystallized with THAP and analyzed by MALDI-TOF-MS. When cultured with Ac4ManNDAz, Jurkat cells are able to incorporate SiaDAz into GM2 and GM1. Unlabeled gangliosides (GM2 – purple, GM1 – green) and SiaDAz-containing gangliosides (GM2 – orange, GM1 – cyan) are denoted above. SiaDAz-containing gangliosides subject to loss of diazirine are denoted by stars (GM2 – orange, GM1 – cyan).
Immunofluorescence microscopy analysis of CTxB trafficking in Ac4ManNDAz-treated
Jurkat cells exposed to UV light
Cholera toxin, a member of the AB5 group of toxins, is secreted by the bacterium
Vibrio cholerae and is responsible for the massive fluid loss that accompanies cholera
infection.18 The toxin is comprised of two distinct subunits: A and B. Using the B
subunit, cholera toxin binds to the host cell through cell surface GM1 with sub-
nanomolar KD.19-22 Upon binding the plasma membrane, cholera toxin is endocytosed and
trafficked through retrograde mechanisms through the endosomes, trans-Golgi network
(TGN), and finally into the endoplasmic reticulum (ER). Once in the ER, the A subunit
is cleaved in two, after which the A1 subunit is transported to the cytosol. Once there,
the A1 subunit activates the heterotrimeric G protein Gs-α, resulting in the activation of
adenylyl cyclase, generating large amounts of cAMP, which induces massive H2O and
chloride secretion by the cells.23
59
After observing that CTxB is able to recognize the unnatural GM1-SiaDAz
ganglioside synthesized by Jurkat cells, I wanted to investigate whether this interaction
would have any effects on the trafficking of CTxB from the cell membrane to the ER.
CTxB has been shown to traffic from the plasma membrane to the ER without being
complexed to the A subunit.24 Cooperative experiments performed in our group have
demonstrated that GM1-SiaDAz expressing Jurkat cells can be efficiently crosslinked to
CTxB (data not shown). Using this knowledge, I examined the intracellular trafficking of
CTxB when complexed directly to GM1 by immunofluorescence microscopy. Jurkat
cells were untreated or cultured with Ac4ManNDAz for 70 hours and then exposed to
CTxB for 10 min to allow for cell surface binding; cells were kept on ice to prevent
endocytosis. The cells were then exposed to 365 nm UV irradiation for 10 minutes to
photoactivate the diazirine into a reactive carbene to covalently bind CTxB to GM1.
Control experiments without photoirradiation were performed with untreated and
Ac4ManNDAz-treated cells. To induce CTxB endocytosis, the cellular media was
replaced with pre-warmed media and incubated for an additional 90 minutes at 37 oC.
The cells were then fixed, permeabilized and immunostained with antibodies against
CTxB and several intracellular organelles. Confocal microscopy was used to determine
subcellular localization of CTxB in the Jurkat cells.
When I analyzed the non-irradiated/unsupplemented Jurkat cells, I observed that
CTxB colocalized with early endosomes, Golgi, and ER organelles (Figure 4A). This
indicates that CTxB is able to traffic from the plasma membrane and into the secretory
pathway without forming a complex with the A subunit. Ac4ManNDAz treatment of
Jurkat cells displayed similar colocalization observations (Figure 4C), indicating that
SiaDAz-engineering of GM1 does not affect CTxB trafficking. When both untreated
(Figure 4B) and Ac4ManNDAz-treated (Figure 4D) cells were photoirradiated, direct
CTxB colocalization was observed for early endosomes and Golgi but not for the ER; I
observed a clear separation occurring outside of the nucleus between CTxB and the ER. I
also noticed that CTxB formed more punctate structures on the outside of cells after UV
exposure. Since this observation was viewed with untreated and Ac4ManNDAz-treated
cells, this suggests that the likely cause of the behavior is UV irradiation, not
crosslinking. Analysis of Jurkat cells by immunofluorescence is further complicated by
60
the immense size of the Jurkat cell nucleus: this makes it difficult to distinguish CTxB
localization between the cell surface and ER. While Jurkat cells are able to synthesize
photoactivatable gangliosides, these results indicate that use of photoirradiation to
crosslink CTxB to GM1-SiaDAz may prevent investigating intracellular trafficking by
immunofluorescence microscopy.
Figure 4. Immunofluorescence analysis of Cholera toxin subunit B trafficking in supplemented Jurkat cells. Jurkat cells, either untreated or Ac4ManNDAz, were cultured with CTxB for 10 minutes on ice. The cells were then photoirradiated for 10 min with 365 nm light. After culturing the cells for 90 minutes at 37oC, CTxB trafficking through several organelles (ER - Calnexin, cis-Golgi – GM130, trans-Golgi network (TGN46), and early endosomes – EEA1) was monitored by immunofluorescence microscopy. Immunostaining for organelle markers is shown in Column 1. Immunostaining for CTxB is shown in Column 2. Column 3 represents the overlay of the images: organelle markers are shown in red, CTxB is shown in green, and the nuclear stain (DAPI) is shown in blue. Colocalization between organelle markers and CTxB appears yellow in the merged image. (A) Untreated Jurkat cells. (B) Untreatedd Jurkat cells exposed to UV light. (C) Ac4ManNDAz-treated Jurkat cells. (D) Ac4ManNDAz-treated cells exposed to UV light.
61
Discussion
Jurkat cells can synthesize gangliosides with photoreactive chemical groups
The primary goal of my studies was to demonstrate successful engineering of
photoactivatable gangliosides in mammalian cells. Metabolic oligosaccharide
engineering is an acknowledged technique to introduce diverse chemical functionality
into cell surface sialosides.9 While our laboratory has demonstrated the successful
incorporation of a diazirine into a sialylated glycoprotein (CD22),10,13 my objective was
to investigate the applicability of this technology into cell surface gangliosides. Using
Jurkat cells, I was able to observe efficient metabolism of ManNDAz into SiaDAz in
several ganglioside species by HPTLC analysis (Figure 2B). Culturing Jurkat cells with
Ac4ManNDAz caused dramatic changes in the composition of endogenous gangliosides;
the levels of unmodified gangliosides diminished and several new gangliosides appeared.
These new species are likely SiaDAz-containing gangliosides as the introduction of the
diazirine side chain increases the hydrophobicity of the molecule, causing a mobility shift
towards the solvent front. To confirm that treatment of Jurkat cells with exogenous
ManNAc does not increase overall ganglioside production, I cultured cells with their
natural substrate, ManNAc, at the same molar concentration as with ManNDAz. As
shown in Figure 2B, cells that were untreated or Ac4ManNAc-treated showed similar
levels of ganglioside production, validating that our culturing techniques are not forcing
an increased production of gangliosides.
To confirm the existence of SiaDAz-containing gangliosides in Jurkat cells, I
submitted the total ganglioside extracts for MALDI-TOF-MS analysis (Figure 3). I
observed that Jurkat cells are able to synthesize GM2-SiaDAz and GM1-SiaDAz. During
these analyses, peaks were observed for several SiaDAz-containing gangliosides that
corresponded to a loss of N2 after irradiation by the instrument laser (Figure 3 – denoted
by stars). This was determined to be an unavoidable consequence of analyzing these
species by MALDI-TOF-MS (Roberto Sonon, personal communication). Regardless of
the loss of diazirine, this technique validated my HPTLC observations that Jurkat cells
were able to synthesize photoactivatable gangliosides.
62
When Jurkat cells were cutlured with Ac4GlcNDAz, there was no apparent
change in the ganglioside production of these cells (Figure 2B). These results were
consistent with MALDI-TOF-MS analysis of the total ganglioside extract (data not
shown). In mammalian cells, GlcNAc is coupled to UDP to generate an activated
nucleotide sugar that is subsequently epimerized by UDP-GlcNAc 2-epimerase into
ManNAc. Theoretically, this synthetic route could be possible for the conversion of
GlcNDAz into ManNDAz; however, my results suggest that this does not occur in Jurkat
cells. It appears that either the UDP-GlcNAc transferase or UDP-GlcNAc 2-epimerase is
unable to accept the introduction of the diazirine side found in GlcNDAz.
Another important aspect of my studies is that ManNDAz competes effectively
with natural Jurkat cell metabolites for incorporation into gangliosides. This was clearly
demonstrated with my HPTLC (Figure 2B, C) and MALDI-TOF-MS (Figure 3) analyses,
where significant levels of unnatural gangliosides were produced. These results are
consistent with previous work performed in our lab showing that ManNDAz can
effectively compete for sialic acid incorporation into the cell surface CD22.10,13
Additionally, I observed that culturing Jurkat cells with Ac4ManNAz, a widely used
substrate for sialic acid metabolic oligosaccharide engineering, can also compete directly
for ganglioside incorporation into Jurkat cells (data not shown).
Cholera toxin subunit B recognizes GM1-SiaDAz and can be efficiently photocrosslinked
I investigated if a well-established binding partner, CTxB, could efficiently
recognize GM1-SiaDAz. For this experiment, I resolved ganglioside extracts from
Ac4ManNDAz-treated Jurkat cells by HPTLC and immunostained against GM1 with a
CTxB-Alexa Fluor 488 conjugate (Figure 2C). The application of immunostaining
techniques for ganglioside analysis on HPTLC plates is well established and is more
sensitive than resorcinol staining.25,26 I observed that CTxB-Alexa Fluor 488 bound to
two additional bands in Ac4ManNDAz-treated Jurkat cells; these bands are likely GM1-
SiaDAz since introducing the diazirine side chain increases their Rf value. The
introduction of the diazirine side chain into GM1 does not appear to dramatically affect
this interaction, suggesting that we should be able to exploit the photoreactivity of the
diazirine to capture this complex. Indeed, cooperative experiments conducted in our
63
laboratory have demonstrated that Ac4ManNDAz-treated Jurkat cells can be efficiently
photocrosslinked to CTxB (data not shown). These results demonstrate that
photoactivatable gangliosides can be used as crosslinking reagents to capture interaction
partners.
The formation of a covalent GM1-CTxB complex does not appear to affect trafficking
from the plasma membrane to the TGN
After demonstrating that CTxB can recognize GM1-SiaDAz synthesized by
Ac4ManNDAz-treated Jurkat cells, I decided to investigate whether the formation of a
covalent complex would affect the retrograde trafficking of CTxB. CTxB has been shown
to traffic from the plasma membrane to the ER without being complexed to the A
subunit.24 I chose to monitor CTxB trafficking in Jurkat cells by performing
colocalization immunofluorescence microscopy using organelle markers for the early
endosomes, TGN, cis-Golgi, and ER. I observed that CTxB was able to efficiently traffic
from the plasma membrane to the ER within the time frame of my experiment, regardless
of culturing with Ac4ManNDAz (Figures 4A, 4C). This result confirms my observations
seen from CTxB-Alexa Fluor 488 immunostaining (Figure 2C). When I exposed the
Jurkat cells to UV irradiation to activate the diazirine, I did not observe any noticeable
changes in CTxB trafficking between unsupplemented and Ac4ManNDAz-treated cells:
CTxB trafficking was limited to the early endosomes and TGN. There was a clear
separation between the fluorescence emanating from CTxB and the ER (Figures 4B, 4D).
In addition, CTxB appeared to distribute into small puncta near the plasma membrane
that did not colocalize with several organelle markers. These results provide preliminary
evidence that photocrosslinking CTxB to GM1 does not affect its ability to traffic from
the plasma membrane toward the TGN. Due to the complications of CTxB trafficking
into the ER because of UV irradiation, I am unable to make any conclusions regarding
the trafficking of covalently formed CTxB-GM1 complexes at this time.
One of the major limitations encountered within this immunofluorescence
analysis is the small size of the cytoplasm in Jurkat cells; most of the intracellular space
is encompassed by the nucleus (Figure 4). The narrow cytosol makes it extremely
difficult to analyze the subcellular distribution of CTxB between the plasma membrane
64
and ER, as these two organelles appear adjacent. Although Jurkat cells provided a useful
starting point for our experiments because of their ganglioside patterning, they have
proven less useful for immunofluorescence experiments. An epithelial cell expressing
GM1 would be the most biologically relevant cell type and might yield a clearer picture
of CTxB trafficking. To this end, our lab is currently exploring the human intestinal
epithelial cell line T84 for its ability to produce SiaDAz-GM1, as it widely used for
studying the intracellular effects of cholera toxin infection.27
In summary, I have demonstrated that a mammalian cell line can produce
photoactivatable gangliosides that are recognized by an established binding partner.
Irradiation of these molecules can efficiently capture this ganglioside-protein interaction.
My results show promise for using this photoreactive tool to capture and investigate
ganglioside-mediated interactions. Future studies are required to investigate the
biological consequences of covalently linked GM1-CTxB in mammalian cells.
Acknowledgements
I would like to thank Kim Orth (UT Southwestern Medical Center) for sharing
Jurkat cells. I would like to thank Michelle R. Bond and Seokho Yu (UT Southwestern
Medical Center) for providing Ac4GlcNDAz. I would like to thank Yan Li (UT
Southwestern Medical Center Protein Chemistry Technology Center) for assistance with
ESI-MS analysis. I would like to thank Pablo Lopez (Johns Hopkins University) and
Aarnoud C. van der Spoel (University of Oxford) for help and advice with ganglioside
extractions. I would like to thank Parastoo Azadi and Roberto Sonon (University of
Georgia Complex Carbohydrate Research Center) for mass spectrometry of ganglioside
samples. Additionally, I would like to thank Michelle R. Bond for helpful suggestions
with my experiments and concurrent analyses involving photocrosslinking of cholera
toxin subunit B to GM1 targets.
65
Methods
Reagents
All chemicals, reagents, and general supplies were purchased from Fisher
Scientific (Waltham, MA) or Sigma-Aldrich (St. Louis, MO) unless noted. N-
hydroxybenzotriazole (HOBt) was purchased from AnaSpec (Fremont, CA). SepPak
tC18 columns (0.3 g) were purchased from Fisher Scientific. HPTLC plates (20 x 20 cm,
glass backed, 200 µm thickness) were purchased from EMD Chemicals (Gibbstown, NJ).
Matreya ganglioside standards 1508, 1510, and 1511 were purchased from Matreya LLC
(Pleasant Gap, MD). Cholera toxin subunit B-Alexa Fluor 488 conjugate was purchased
from Invitrogen (Carlsbad, CA).
Cell culture reagents, including RPMI 1640 with 2 mM L-glutamine were
purchased from Invitrogen (Carlsbad, CA). Cholera toxin B subunit B (from Vibrio
cholerae) was purchased from Sigma Aldrich. Mouse anti-GM130 and mouse anti-EEA1
were purchased from BD Biosciences (San Jose, CA). Chicken anti-cholera toxin subunit
B, rabbit anti-TGN 46, and goat anti-chicken FITC were purchased from Abcam
(Cambridge, MA). Rabbit anti-calnexin (H-70) was purchased from Santa Cruz
Biotechnology, Inc. (Santa Cruz, CA). Goat anti-mouse Alexa Fluor 546, goat anti-rabbit
Alexa Fluor 546, goat anti-mouse Alexa Fluor 633, and goat anti-rabbit Alexa Fluor 633
were purchased from Invitrogen. Vectashield mounting medium with DAPI was
purchased from Vector Labs (Burlingame, CA). 0.1% poly-L-lysine solution was
obtained from Sigma (St. Louis, MO).
Cell culturing conditions
Jurkat cells were grown and maintained in RPMI 1640 with 2 mM L-glutamine
containing 10% heat-inactivated FBS at 37oC, 5% CO2 in a water-saturated environment.
Cells were cultured at 2.0-2.5 x 105 cells/mL in media and grown for 48 hours before
passaging. Typically, cell densities were maintained between 2.5 x 105 cells/mL and 2.0
x 106 cells/ml. Cell viability was analyzed using Trypan blue dye staining with the
Countess Automated Cell Counter instrument (Invitrogen).
66
General information for the chemical synthesis of N-acetylmannosamine analogs
All chemicals were used as received from commercial suppliers without further
purification. 1-hydroxybenzotriazole hydrate was purchased from AnaSpec. All other
chemicals were purchased from Sigma-Aldrich or Fisher Scientific unless otherwise
noted. Reaction progress was monitored by analytical thin layer chromatography (TLC)
on silica gel 60 F254 glass backed plates (Fisher) and stained with ceric ammonium
molybdate. Flash column chromatography was carried out with silica gel 60 (particle
size 40-63 µm, EMD Chemicals). All 1H-NMR and 13C-NMR spectra were recorded on
a Varian 500 MHz spectrometer and are reported in δ ppm scale. 1H-NMR spectra were
referenced to D2O (4.80 ppm) or CDCl3 (7.26 ppm). 13C-NMR spectra were referenced
to CDCl3 (77.23 ppm). ESI-MS data were collected at the UT Southwestern Medical
Center Protein Chemistry Technology Center. All acetylated sugars were prepared as 10
mM stock solutions in ethanol. The purity of acetylated sugars was confirmed by HPLC
analysis before cellular treatments (spectra located in appendix).
Synthesis of Ac4ManNAc
Ac4ManNAc was synthesized as previously reported.13 Briefly, to a solution of D-
(+)-N-acetylmannosamine (301.7 mg, 1.36 mmol) in pyridine (16.4 mL, 204 mmol),
acetic anhydride (4.72 mL, 54 mmol) was added and stirred overnight on ice. The
reaction mixture was diluted by CH2Cl2 and washed successively by 1.0 M HCl,
saturated sodium bicarbonate, and brine. The organic layer was dried over magnesium
sulfate and evaporated in vacuo. The residue was purified by flash chromatography
(hexanes / ethyl acetate gradient = 5/1, 3/1, 1/1) to afford Ac4ManNAc (282 mg, 53%,
mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.65 (3H, s), 2.02 (3H, s), 2.07
(3H, s), 2.11 (3H, s), 2.18 (3H, s), 4.10 (1H, dd, J = 2.3, 12.5), 4.28 (1H, t, J = 3.7), 4.78
(1H, ddd, J = 1.6, 3.9, 9.1), 5.06 (1H, d, 4.0), 5.13 (1H, t, J = 9.8), 5.33 (1H, d, J = 4.5),
5.79 (1H, d, J = 9.0), 5.86 (1H, d, J = 1.6). 13C-NMR (125 MHz, CDCl3): δ 20.88, 20.90,
20.92, 20.96, 20.97, 21.01, 21.08, 23.56, 23.65, 49.52, 49.74, 62.20, 65.41, 65.62, 68.99,
70.29, 71.56, 73.68, 90.86, 91.90, 168.34, 168.55, 169.92, 169.93, 170.23, 170.32,
67
170.73, 170.74, 170.82. ESI-MS for C16H23NO10 [M], calculated for 389.13, found
389.12. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the appendix.
Synthesis of Ac4ManNDAz
ManNDAz was synthesized as previously reported.10 Briefly, to a solution of 4,4-
azo-pentanoic acid28 (128 mg, 1.00 mmol), D-(+)-mannosamine hydrochloride (216 mg,
1.00 mmol) and triethylamine (278 µL, 2.00 mmol) in MeOH (10 mL), 1-ethyl-3-(3-
dimethyllaminopropyl)carbodiimide hydrochloride (383 mg, 2.00 mmol) and 1-
hydroxybenzotriazole hydrate (135 mg, 1.00 mmol) were added. The reaction mixture
was stirred on ice for 10 minutes, followed by stirring at room temperature overnight.
The resulting mixture was concentrated in vacuo, roughly purified by flash
chromatography (CH2Cl2 / MeOH gradient = 1/0, 10/1, 4/1), and used directly to
synthesize the acetylated product, Ac4ManNDAz. The acetylation of ManNDAz was
performed by the same procedure described in the synthesis of Ac4ManNAc to afford
Ac4ManNDAz (84 mg, 28% over two steps, mixture of anomers). 1H-NMR (500 MHz,
CDCl3): δ 1.06 (3H, s), 1.81 (2H, m), 2.02 (3H, s), 2.07 (3H, s), 2.12 (3H, s), 2.19 (3H,
s), 4.05 (2H, s), 4.09 (1H, ddd, J = 6.2, 18, 30.5), 4.29 (1H, t, J = 4.5), 4.78 (1H, dd, J =
2.3, 8.2), 5.06 (1H, d, J = 4.0), 5.20 (1H, t, J = 9.8), 5.32 (1H, d, J = 4.4), 5.80 (1H, d, J =
9.0), 6.04 (1H, s). 13C-NMR (125 MHz, CDCl3): δ 20.18, 20.19, 20.85, 20.88, 20.91,
20.95, 20.97, 21.08, 25.50 25.54, 29.94, 30.05, 30.71, 30.84, 49.55, 49.76, 62.02, 62.15,
65.32, 69.06, 70.32, 71.57, 73.67, 90.80, 91.79, 168.33, 168.53, 169.82, 169.91, 170.20,
170.28, 170.75, 170.78, 171.62, 172.14. ESI-MS for C19H27N3O10 [M], calculated for
457.17, found 457.16. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the
appendix.
Exposure of Jurkat cells with N-acetylmannosamine analogs – Ganglioside Analysis
Jurkat cells were seeded at a density of 2.5 x 105 cells/mL in 15 cm tissue culture
plates in 60 mL of media. Prior to the addition of cells to a tissue culture plate, acetylated
sugar in ethanol or ethanol only was added and the solvent was evaporated at ambient
temperature and pressue. For each condition, 2-3 plates of cells were used. After
growing for 72 hours, cells were counted and centrifuged at 220g for 5 min in 50 ml
68
conical tubes. To ensure consistent results among all samples, equal numbers of cells
were collected for every sample; the total number of cells collected for an experiment
ranged between 1.5 – 2.0 x 108 cells. Cell pellets were stored at -80 oC overnight before
proceeding to ganglioside extraction.
Extraction of gangliosides - Total Lipid Extraction
Cell pellets were thawed to room temperature, resuspended with 300 µl of ice
cold ddH2O (W), and dounced 50 times with a Kontes tissue grinder, tube size 20. With
a glass Pasteur pipette and a 2 mL rubber bulb, the cell lysate suspension was transferred
into a 4 mL glass vial containing 800 µL of methanol (M), already stirring. 400 µL of
chloroform (C) was added to the vial and the mixture was stirred thoroughly for 2 hours
at room temperature. Samples were covered in foil to prevent exposure to light. After
stirring, the mixture was transferred by a glass Pasteur pipette into a 13 x 100 mm glass
culture tube and centrifuged at 2800g for 10 min @ 30 oC. The supernatant (containing
the total lipid extract) was transferred by a glass Pasteur pipette into a new 4 mL glass
vial and evaporated to dryness under N2 gas.
Extraction of gangliosides - Phospholipid Extraction
The dried total lipid extract was resuspended with 800 µL butanol and 1200 µL
diisopropyl ether and sonicated in a water bath for 10 minutes. The resuspended lipids
were then transferred into a 13 x 100 mm glass culture tube using a glass Pasteur pipette.
To extract undesired phospholipids from the mixture, 1000 µl of 50 mM NaCl was added
to the tube and mixed vigorously by pipetting up and down repeatedly with a glass
Pasteur pipette. The mixture was then centrifuged at 2800g for 10 min at 30 oC to
separate the two phases. Using a glass Pasteur pipette, the organic phase (top layer) was
carefully removed. The aqueous mixture (bottom layer) was then extracted two more
times using the same ratio of butanol and diisopropyl ether.
Extraction of gangliosides - SepPak purification
After the final extraction, the remaining lipid mixture was loaded onto a SepPak
tC18 column, 0.3g size. The column was first pre-treated with three 2 mL washes of
69
C/M/W (2:43:55) followed by two 2 mL washes of C/M (1:1) and ending with three more
2 mL washes of C/M/W (2:43:55). After loading of the sample, the column was washed
three times with 2 mL of C/M/W (2:43:55) followed by three 2 mL washes of C/M (1:1)
to desalt the sample and remove unwanted contaminants. Elution of gangliosides was
achieved using 2 mL of 100 % methanol. Ganglioside extracts were then transferred into
a new 4 mL glass vial and evaporated to dryness under N2.
HPTLC Analysis of Extracted Gangliosides
Extracted ganglioside samples were redissolved with 30 µL C/M/W (2:1:0.1) and
resolved on HPTLC plates. Ganglioside standards were loaded onto the plate to provide
mass references. Gangliosides were separated with chloroform:methanol:0.2% CaCl2(aq)
(80:45:10) as the running buffer. HPTLC plates were first pre-run before loading 10 µL
of ganglioside extract. After thoroughly drying the plate in a fume hood, gangliosides
were detected by resorcinol staining (0.1% resorcinol, 0.04% CuSO4 in hydrochloric
acid:water [4:1]). Plates were imaged using an Alpha Innotech FluorChem HD2 and
images were processed using Adobe Photoshop.
To specifically detect GM1, extracted gangliosides were separated by HPTLC as
described above. The plate was fully dried under 40 mbar vacuum in a dessicator for 45
minutes. The plate was then treated with 0.5% polyisobutylmethacrylate in hexanes
(diluted from a 2.0% stock solution prepared in CHCl3) for 2 minutes by immersion in
the solution. The plate was then dried thoroughly with a stream of air, followed by
immersion in 1.0% BSA/PBS for 30 minutes at room temperature. The plate was
incubated with cholera toxin subunit B, Alexa Fluor 488 conjugate solution (1:20,000
dilution, in PBS) for 50 minutes at room temperature, followed by several brief washes
with PBS. After fully drying the plate using a stream of air, the plate was imaged by
Typhoon using the 488 excitation laser and 520 emission filter. Image analysis was
performed using Adobe Photoshop. Resorcinol staining was performed (as described
above) after fluorescence imaging to confirm ganglioside composition.
70
Mass Spectrometry Analysis of Extracted Gangliosides
Dried ganglioside extracts were sent to the Complex Carbohydrate Research
Center at the University of Georgia for mass spectrometry analysis. To analyze the
overall composition of extracted gangliosides, MALDI-TOF-MS was performed.
Samples were crystallized onto a MALDI plate with trihydroxyacetophenone
monohydrate (THAP) as a matrix. Analysis of gangliosides was performed in the
negative ion mode using a Bruker microflex instrument. Spectra are presented in the
appendix.
Exposure of Jurkat cells with N-acetylmannosamine analogs – Immunofluorescence
analysis
Jurkat cells were seeded at a density of 2.5 x 105 cells/mL into individual wells of
6-well tissue culture plates in 4 mL of media. Prior to the addition of cells to the tissue
culture plate, acetylated sugar or ethanol was added and evaporated. For each condition,
3-4 individual wells were used. After growing for 70 hours, the cells were counted, spun
down at 200g for 5 min, and resuspended in original media to 5.0 x 106 cells/mL.
Coverslip preparation and poly-L-lysine coating
Coverslips were heated in a loosely covered glass beaker in 1 M HCl at 50oC for 4
hours. After cooling to room temperature, the slides were rinsed several times with
ddH2O to remove all traces of acid. The slides were then sonicated three times for 30
minutes in ddH2O, changing the solution in between. The coverslips were then sonicated
successively with 50%, 70% and 95% ethanol, each for 30 minutes. Slides were then
dried on filter paper and stored. To generate poly-L-lysine coated slides, coverslips were
coated in 0.1% (w/v) poly-L-lysine solution using a Petri dish overnight at 4oC, while
rotating. The coverslips were then washed at least 10x in ddH2O to remove poly-L-lysine
solution. Then the coverslips were rinsed with 100% ethanol and dried with filter paper
in a sterile hood. Cells were added onto the coverslips when dry.
71
Cholera toxin incubation and photocrosslinking
1000 µl of Jurkat cell suspension was transferred into individual wells of new 6-
well plates containing poly-L-lysine coated coverslips tamped onto the bottom surface.
Cells were incubated for 20 min at room temperature to allow the Jurkat cells to bind to
the lysine-coated surface. After incubation, the plates were transferred onto ice for 10
min. To each well, 5 µL of cholera toxin subunit B (1 mg/mL) was added and the plate
was gently swirled, on ice. The plates were then exposed to 365 nm light for 10 min –
approximately 2 cm away from bulb. After irradiation, the culture media was removed
and replaced with fresh pre-warmed media. Cells were incubated at 37oC, 5% CO2 for 90
minutes to allow for endocytosis of cholera toxin subunit B.
Immunofluorescence analysis
After incubation and photocrosslinking, the plates were immediately transferred
back onto ice, the supernatant aspirated, and directly fixed with 3.7%
paraformaldehyde/PBS for 20 minutes without washing. After several washes with PBS,
the cells were permeabilized with 0.1% Triton X-100, 1.0% BSA/PBS for 5 minutes
without shaking. After several washes with PBS, the plates were placed at room
temperature and incubated with 1.0% BSA/PBS for 1 hour to block non-specific binding.
After blocking, the cells were incubated with primary antibody (1:500 dilution; rabbit
anti-calnexin was used at 1:50 dilution) for 2 hours at room temperature. The coverslips
were then washed three times with PBS and the cells were incubated with secondary
antibody (1:500 dilution) for 1 hour at room temperature. The coverslips were washed
again three times with PBS. After washing, the coverslips were mounted onto glass
cover slides using Vectashield Mounting Medium with DAPI and sealed with nail polish.
Slides were stored at 4oC until analysis. Cells were visualized using the HCX PL APO
Lambda Blue 63x 1.40 oil UV objective of a Leica TCS SP5 confocal microscope
equipped with 405 nm, 488 nm, 561 nm, and 633 nm lasers. Image analysis was
performed using ImageJ (NIH), Adobe Photoshop, and Adobe Illustrator.
72
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cholera and related toxins from the plasma membrane to endoplasmic reticulm.
Mol Biol Cell 14, 4783-4793, (2003).
25 Schnaar, R. L. & Needham, L. K. Thin-layer chromatography of
glycosphingolipids. Methods Enzymol 230, 371-389, (1994).
26 Lopez, P. H. & Schnaar, R. L. Determination of glycolipid-protein interaction
specificity. Methods Enzymol 417, 205-220, (2006).
27 Wolf, A. A., Fujinaga, Y. & Lencer, W. I. Uncoupling of the cholera toxin-G(M1)
ganglioside receptor complex from endocytosis, retrograde Golgi trafficking, and
downstream signal transduction by depletion of membrane cholesterol. J Biol
Chem 277, 16249-16256, (2002).
28 Church, R. F. R. & Weiss, M. J. Diazirines. II. Synthesis and Properties of Small
Functionalized Diazirine Molecules - Some Observations on Reaction of a
Diaziridine with Iodine-Iodide Ion System. Journal of Organic Chemistry 35,
2465-2471, (1970).
75
Chapter 4 – Exploring the incorporation of sialic acid analogs into gangliosides of
mammalian cells
Introduction
The cell surface expression of sialic acid plays an integral role in facilitating
numerous molecular interactions on the eukaryotic plasma membrane.1 For example,
sialic acid is responsible for mediating adhesion of leukocytes to endothelial cells at sites
of inflammation.2 Sialic acids are commonly expressed into a wide variety of cell surface
glycoconjugates, including glycolipids. Sialylated glycolipids, known as gangliosides,
are responsible for many biological roles, including cell signaling, cell-cell
communication, and pathogen recognition.3 They are also highly expressed in the
developing mammalian brain4 and in malignancy.5 Since gangliosides are anchored
directly into the plasma membrane, studying their function and molecular details in a
native environment is often difficult.6 This hindrance precipitates the need to develop
new biological methods that can be used to examine gangliosides’ behavior in their
native environment.
One approach to investigating the roles of cell surface sialic acid is to introduce
small structural modifications by culturing cells with analogs of sialic acid or N-
acetylmannosamine (ManNAc), which is a metabolic precursor of sialic acid. Known as
metabolic oligosaccharide engineering, this technique relies on the ability of cells to
uptake these analogs through passive diffusion, process them with their natural cellular
machinery, and insert the unnatural metabolite into glycan structures in place of the
natural sugar.7,8 Pioneering work in the early 1990s by Reutter and co-workers
demonstrated that molecules in which the N-acyl chain of sialic acid was lengthened with
additional methylene units could be readily tolerated and incorporated into cell surface
glycan structures in cells and live animals.9,10 This technology demonstrated to be highly
applicable for exploring host-virus interactions,11 neuronal cell differentiation,12 and
inhibiting cell surface polysialylation.13 Since then, metabolic oligosaccharide
engineering has expanded its utility to introduce chemically reactive functional groups
(i.e. ketones,14 azides,15,16 alkynes17) into cell surface sialic acid.18 The incorporation of
these bio-orthogonal groups has provided a chemical handle for labeling sialylated glycan
76
structures in vivo.19-24 Recently, the ability to capture sialic acid-mediated interactions,
which suffer from weak affinity and fast off-rates, has been achieved by metabolic
oligosaccharide engineering techniques through the introduction of photoactivatable
groups.16,25,26
While this technology has proved effective at studying the many biological roles
of sialic acid, the majority of these studies have investigated the global incorporation of
unnatural sialic acid analogs. In humans, sialic acid is attached onto glycan chains in
α2,3-, α2,6-, and α2,8-linkages. Production of sialic acid glycoconjugates, or sialosides,
is controlled by 20 different sialyltransferases, each with individual substrate
specificities. This diversity presents a potential problem in generalizing metabolic
oligosaccharide engineering results for all types of sialosides. Because gangliosides can
be extracted from cells and independently analyzed, these molecules provide an excellent
target to study the application of metabolic oligosaccharide engineering technology into
specific sialosides. Several groups have reported the use of metabolic oligosaccharide
engineering employing ManNAc analogs to produce unnatural gangliosides in
mammalian cells (detailed in Chapter 1). ManNAz, an azide-containing analog, has
been successfully metabolized into SiaNAz, which was incorporated into cell surface
gangliosides of Jurkat cells.27 ManNProp and ManNPhAc have also been successfully
metabolized into cell surface gangliosides in several cancer lines and been utilized as
immunogenic targets for antibodies.28-31 ManNGc, which can be metabolized into
NeuGc, has been used to metabolically produce hydroxylated cell surface gangliosides in
neuronal cells to investigate myelin-axon interactions.32
To further probe the substrate flexibility that is possible with ganglioside
biosynthesis, I synthesized a panel of ManNAc analogs that are known to be metabolized
to sialosides and investigated their incorporation into gangliosides. In addition to above
mentioned ManNAc analogs, my panel was expanded to include ManNBut10,33 and
ManNDAz26 which have been previously demonstrated to become metabolized into cell
surface glycoconjugates. Because of the diversity of ganglioside structures present in
mammalian cells, I focused my experiments on production of the GM3 ganglioside,
which is the precursor for other ganglioside products. To learn more about specificity
77
differences between mammalian species, I conducted my experiments in both human
(BJAB) and hamster (CHO) cell lines. My results reveal significant differences between
these two cell lines’ ability to accept and incorporate modified sialic acids into
gangliosides.
Results
Incorporation of variant sialic acids into gangliosides
To test whether sialic acid analogs are incorporated into the ganglioside GM3, I
relied on a key set of reagents: two cell lines, BJAB K2034,35 and CHO Lec3,36 that are
impaired in sialic acid biosynthesis through their inactive UDP-GlcNAc 2-epimerase
(Figure 1). This enzyme catalyzes the conversion of UDP-GlcNAc into ManNAc and is
vital for production of sialic acid in mammalian cells. When these epimerase-deficient
cell lines are cultured in serum free conditions, they are unable to synthesize gangliosides
due to their inability to produce sialic acid. Two metabolites within the sialic acid
biosynthesis pathway, ManNAc and sialic acid, present an entry point for introducing
chemical modifications into sialylated glycans (Figure 1 – highlighted in green): in this
work, I have chose to use ManNAc analogs due to their synthetic simplicity compared to
generating sialic acid analogs. When cells are cultured with ManNAc, they regain the
ability to generate sialylated glycoconjugates, including gangliosides. Similarly, when
the cells are cultured with unnatural ManNAc analogs, any observation of gangliosides is
interpreted as reflecting successful incorporation of these unnatural analogs into
gangliosides. BJAB cells primarily generate GM3 but also are able to produce small
amounts of GM1 (confirmed by MALDI-TOF-MS analysis, data not shown). CHO cells
are only able to generate GM3 because they lack the enzymes needed to synthesize
downstream ganglioside products.37
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Figure 1. Metabolic oligosaccharide engineering of UDP-GlcNAc 2-epimerase-deficient cells occurs through the introduction of ManNAc and sialic acid analogs. BJAB K20 and CHO Lec3 cells lack UDP-GlcNAc 2-epimerase activity, rendering them unable to generate sialosides when cultured in serum free media. If these cells are cultured with downstream metabolites in this pathway, they regain the ability to generate sialosides. Two metabolites, ManNAc and sialic acid (highlighted in green), serve as entry points for introducing chemical modifications into sialosides (denoted in red).
I chose to use a panel of ManNAc analogs that have reported to be metabolized to
sialosides in one or more cell types;18 this panel includes several analogs which have
been previously shown to be metabolized to their sialic acid counterparts and
incorporated into gangliosides (Figure 2).27,28,30,31 BJAB K20 and CHO Lec3 cells were
cultured in serum free conditions with each of the ManNAc analogs for 48-72 hours.
Free hydroxyl groups present on each sugar were acetylated with acetic anhydride to
improve diffusion through the membrane.38 The acetyl protecting groups are believed to
be removed by nonspecific esterases inside the cells. After culturing with acetylated
ManNAc analogs, the cells were harvested and the gangliosides were extracted using
established methods (detailed in Methods).39-41 Gangliosides were resolved by high
performance thin layer chromatography (HPTLC) and visualized by resorcinol staining,
which specifically detects sialic acid-containing molecules.
79
Figure 2. Panel of ManNAc analogs used in ganglioside incorporation experiments. In order to investigate the incorporation of unnatural sialic acid analogs into gangliosides, a panel of unnatural N-acyl modifications were introduced into ManNAc and cultured with BJAB K20 and CHO Lec3 cells. All of these analogs have been demonstrated to be metabolized to cell surface sialosides. The color-coding scheme used for these analogs is continued throughout this chapter.
When I examined the ability of BJAB K20 cells to incorporate unnatural
ManNAc analogs into GM3, I observed that these human cells were quite permissive to
unnatural modifications to the N-acyl side chain of sialic acid (Figure 3A). Increasing
the length of the N-acyl side by additional methylene groups can easily be
accommodated, demonstrated by incorporation of ManNProp and ManNBut. This
apparent increase in N-acyl chain length also allows for efficient metabolism of ManNAz
and ManNDAz, introducing chemical functionality in the side chain of sialic acid.
However, the incorporation of ManNGc into GM3 as NeuGc appears to be significantly
reduced in comparison to the natural substrate ManNAc. Only one analog (ManNPhAc)
was not incorporated into GM3 at levels detectable by resorcinol staining. MALDI-TOF-
MS analysis (located in appendix) of extracted gangliosides from BJAB K20 cells
cultured with ManNAc, ManNGc, ManNAz, and ManNDAz confirmed the identity of
the modified GM3 molecules.
When I cultured CHO Lec3 cells with the same panel of ManNAc analogs, I
observed that the hamster cells display restricted substrate tolerance (Figure 3B). In
comparison to BJAB K20 cells, CHO Lec3 cells appear to only tolerate minimal
perturbations to the N-acyl side chain of sialic acid, evidenced from ManNProp culturing
experiments. Increasing the side chain beyond one methylene group led to diminished
80
levels of incorporation for several analogs, ManNBut and ManNAz; longer N-acyl side
chains, such as ManNPhAc and ManNDAz, were not incorporated at detectable levels.
Unlike BJAB K20 cells, CHO cells were capable of metabolizing ManNGc and
incorporating the sialic acid analog into GM3 at levels nearly comparable to ManNAc.
MALDI-TOF-MS analysis of extracted gangliosides from CHO Lec3 cells cultured with
ManNAc, ManNGc, and ManNAz confirmed successful incorporation of their sialic acid
counterparts into GM3 (located in appendix).
Figure 3. HPTLC analysis of gangliosides produced by BJAB and CHO cells cultured with ManNAc analogs. The ganglioside extracts of BJAB K20 and CHO Lec3 cells (epi -) cultured with various ManNAc analogs were resolved by HPTLC and visualized by resorcinol staining. When K20 and Lec3 cells are cultured under serum-free conditions, they lose the ability to synthesize gangliosides; if the cells are cultured with their natural substrate (ManNAc), normal ganglioside production is restored. BJAB K88 and CHO cell lines possessing normal functioning UDP-GlcNAc 2-epimerase activity were used as positive controls (epi +). (A) BJAB cells generate GM3 and GM1 gangliosides. BJAB cells are able to effectively metabolize ManNProp, ManNBut, and ManNAz to their sialic acid counterparts and incorporate them into GM3. Cells cultured with ManNGc and ManNDAz demonstrate significantly lower levels of GM3 production, while cells cultured with ManNPhAc do not produce detectable levels of GM3. (B) CHO cells produce one ganglioside: GM3. CHO cells are effectively metabolize only ManNGc and ManNProp into their sialic acid counterparts and incorporate them into GM3. Cells cultured with ManNBut and ManNAz display significantly lower levels of GM3 production, while cells cultured with ManNPhAc and ManNDAz do not produce detectable levels of GM3.
81
These observations highlight key differences in the utilities of these two cell lines
for performing metabolic oligosaccharide engineering experiments with gangliosides.
The human BJAB cell line demonstrates the substrate flexibility to introduce larger
modifications that include chemical functionalities (such as azides or diazirines) into
gangliosides but shows attenuated ability to introduce other naturally occurring forms of
sialic acid (NeuGc). Conversely, the hamster cell line shows much restricted substrate
scope but, unlike the BJAB cell line, is able to utilize ManGc as a substrate for generating
GM3-NeuGc.
Analysis of engineered cell surface sialylation with BJAB and CHO cell lines cultured
with ManNAc analogs
The HPTLC experiments performed above provide a great deal of information
regarding the ability of BJAB and CHO cells to metabolize ManNAc analogs and
incorporate their sialic acid counterparts into GM3. We were surprised to observe
significant differences between the two cell lines, and decided to perform additional
experiments to determine whether the differences reflected the overall ability of the cells
to metabolize these analogs to sialosides or if the differences were specific to ganglioside
production. To look at global cell surface sialoside production, I cultured BJAB K20 and
CHO Lec3 cells with my ManNAc analog panel and used flow cytometry to measure the
incorporation of these unnatural analogs into cell surface sialosides by lectin binding.
Lectins are a family of plant and animal carbohydrate binding proteins that are readily
used to detect specific glycan epitopes and linkages. For analysis of sialosides, α2,3-
linked sialosides were detected with the Maackia amurensis lectin (MAA) and α2,6-
linked sialosides were detected with the Sambucus nigra lectin (SNA). Currently, there
are no commercially available sources of accurately detecting α2,8-linked sialosides.
Because lectin recognition of sialosides could potentially be hindered by structural
changes to sialic acid and to include α2,8-linked structures, I examined global sialic acid
engineering using the recently published periodate oxidation and aniline-catalyzed oxime
ligation (PAL).42 This novel method uses mild periodate oxidation to convert the glycolyl
chain of sialic acid into a reactive aldehyde that can be covalently labeled by an
82
aminooxy-functionalized biotin for detection. Because the presence of cell surface
aldehydes do not occur naturally, this method can be used to specifically label sialic acid.
Because sialic acid is α2,3-linked in the GM3 ganglioside, I first compared the
ganglioside incorporation experiments in BJAB cells (Figure 3A) to the overall
incorporation of α2,3-linked sialic acid on the cell surface of BJAB cells as measured by
MAA binding (Figure 4A). The ganglioside incorporation experiments and the MAA
binding experiments resulted in similar trends: significant levels of incorporation were
observed with extended N-acyl alkyl chains (ManNProp and ManNBut) and with the
introduction of azide group (ManNAz), while the hydroxyl- and diazirine-modified
analogs (ManNGc and ManNDAz) yielded slightly lower levels of incorporation. Next, I
examined the incorporation of analogs into α2,6-linked sialic acid glycoconjugates on the
surface of BJAB cells (Figure 4B). I observed the same trends for analog incorporation
with one exception: ManNGc. When BJAB K20 cells were cultured with ManNGc, I
observed low levels of MAA binding (Figure 4A) but the levels of SNA binding exceed
those produced by the natural substrate (ManNAc) (Figure 4B). However, the amount of
ManNGc incorporation into the total cell surface sialosides, as measured by PAL, was
substantially lower (Figure 4C). These results suggest NeuGc is efficiently incorporated
into α2,6-linked sialosides, but less efficiently incorporated into α 2,3linked sialosides.
Because I observe robust levels of SNA binding, it seems unlikely that the low levels of
MAA binding result from inefficient metabolism of ManNGc to the nucleotide sugar
donor, CMP-NeuGc. Rather, I speculate that the incorporation differences result from
differences in the ability of individual sialyltransferases to tolerate modifications to sialic
acid.
Next, I used lectin binding to examine the ability of different ManNAc analogs to
be metabolized to sialosides and displayed on the surface of CHO cells (Figure 4D). My
observations were consistent with the ganglioside incorporation experiments conducted
in CHO cells: the highest levels of incorporation were only seen with small modifications
to the N-acyl chain (ManNGc and ManNProp) while further increases in size diminished
the levels of incorporation (ManNBut, ManNAz, and ManNDAz). These results were
also fairly consistent with total cell surface sialoside measurements made by PAL (Figure
83
4E). I was unable to examine α2,6-linked glycoconjugates since CHO cells are devoid of
ST6Gal1 enzyme, the primary enzyme responsible for α2,6-linked glycoconjugates found
on N-linked glycosylation.43
Figure 4. Flow cytometry analysis of cell surface display of modified sialic acids. Cell surface sialylation of engineered BJAB K20 and CHO Lec3 cells (epi -) was analyzed by lectin binding (MAA - α2,3-sialic acid linkages, SNA - α2,6-sialic acid linkages) and periodate oxidation and aniline-catalyzed oxime ligation (PAL – measures total cell surface sialylation). BJAB K88 and CHO cell lines possessing functional UDP-GlcNAc 2-epimerase activity were used as positive controls (epi +). Experiments were performed in biological triplicate; duplication of the entire experiment yielded similar results. (A) MAA lectin analysis of cultured BJAB cells. (B) SNA lectin analysis of cultured BJAB cells. (C) PAL analysis of cultured BJAB cells. (D) MAA lectin analysis of cultured CHO cells. (E) PAL analysis of cultured CHO cells. SNA binding was not assayed in CHO cells because they lack production of α2,6-linked sialosides.
84
Discussion
Metabolic oligosaccharide engineering is an emerging technique that provides the
ability to introduce small structural changes into individual monosaccharides to probe the
biological roles and responsibilities of cell surface glycosylation.18 To date, this
technique has provided significant information regarding glycoproteins while the focus
on glycolipid engineering, specifically gangliosides, has only recently begun to be
explored.28-31 My goal was to explore the substrate flexibility of ganglioside
sialyltransferases with metabolic oligosaccharide engineering using a panel of ManNAc
analogs reported to become metabolized into cell surface sialosides. My experimental
system relied on two UDP-GlcNAc 2-epimerase deficient cell lines (BJAB K20 and CHO
Lec3) that, when devoid of serum rich media, are only able to generate sialic acid-
containing gangliosides when supplemented with ManNAc (Figure 1).34-36 This unique
system provides a direct readout of the successful incorporation of unnatural sialic acid
analogs into gangliosides.
Incorporation of modified sialic acids into the GM3 ganglioside of BJAB cells
I cultured BJAB K20 cells with a panel of ManNAc analogs (Figure 2) and
visualized their incorporation of the resulting sialic acid analog into gangliosides by
HPTLC (Figure 3A). BJAB K20 cells appear to tolerate extension of the N-acyl chain on
GM3 with additional methylene units using ManNProp and ManNBut. This promiscuity
in hsaST3Gal5 allows for the introduction of bio-orthogonal chemical groups into GM3,
as evidenced with BJAB K20 cells cultured with ManNAz and ManNDAz. Interestingly,
the relative production of GM3-SiaNAz in BJAB K20 cells appears to be consistent to
normal GM3 levels. This raises the possibility that many reported ManNAz labeling
experiments3,15,19-24,44 could be labeling gangliosides along with glycoproteins.
Generating SiaDAz-labeled GM3 presents an excellent opportunity to investigate the
molecular basis for many recently reported associations involving this ganglioside, as
described in Chapter 1.3,44 When comparing the relative levels of the GM3 analog being
produced, it appears that increasing the size of the N-acyl chain decreases ganglioside
production (Figure 3). This observation was found to be consistent with the flow
cytometry measurements made on α2,3-linked glycoconjugates (Figure 4A) and α2,6-
85
linked glycoconjugates (Figure 4B). Because the introduction of N-acyl chain
modifications on sialic acid can hinder lectin recognition, I analyzed the cell surface
sialoside production by PAL and observed a consistent global pattern of glycan
engineering. These results indicate that metabolic oligosaccharide engineering of
gangliosides is generally comparable to similar experiments targeting glycoproteins in
human cells.
While BJAB K20 cells were able to metabolize several ManNAc analogs into
GM3, I observed diminished incorporation of ManNGc (discussed below) and was
unable to detect the utilization of ManNPhAc (Figure 3A). ManNPhAc was included
within this panel due to its successful incorporation into GM3 in two human tumor cell
lines: K562 and SK-MEL-28.30 In these experiments, a highly specific antibody
generated against this epitope was used to detect its presence on the cell surface. Because
antibody recognition is more sensitive than resorcinol staining, it is plausible that
ManNPhAc engineering is occurring at extremely low levels. The metabolic engineering
experiments performed here were done with a higher concentration (100 µM) of the
ManNAc analog than used previously with ManNPhAc (40 µM); this decision was based
upon our group’s previous work investigating optimal levels of ManNDAz incorporation
into BJAB K20 cells.25 It is possible that reducing the concentration of ManNPhAc in the
culture media might allow for detectable synthesis of GM3-SiaPhAc. Examination of
α2,3-linked (Figure 4A), α2,6-linked (Figure 4B) and total cell surface sialosides (Figure
4C) demonstrated minimal incorporation levels of ManNPhAc into sialylated glycan
structures, indicating that this trend is likely occurring with all types of sialyltransferases.
Another possibility for the observed differences could involve the type of cancer cell line
that was chosen. Due to the mutational differences that exist between these cancerous
cell lines, it is difficult to identify the exact mechanism behind these differing
observations. BJAB K20 cells were chosen for these studies for their ability to provide a
direct readout of metabolic oligosaccharide engineering without competition from
endogenous metabolites.
86
Incorporation of modified sialic acids into the GM3 ganglioside of CHO cells
I cultured CHO Lec3 cells with a panel of ManNAc analogs (Figure 2) and
visualized their incorporation into gangliosides by HPTLC (Figure 3B). CHO Lec3 cells
appear to tolerate minor extensions onto the N-acyl chain on GM3, observed with
ManNGc and ManNProp. Only trace amounts of gangliosides could be detected with the
longer extensions from ManNProp and ManNAz, while both ManNPhAc and ManNDAz
were unable to be detected. These results were fairly consistent with the flow cytometry
measurements made on α2,3-linked glycoconjugates (Figure 4D) and total cell surface
sialosides (Figure 4E). The ability of CHO cells to introduce ManNAz into cellular
glycans is well documented; CHO cells are able to incorporate ManNAz into cellular
glycoproteins,45,46 including α2,3-linked glycoconjugates 47 (shown here in Figure 4D).
However, CHO cells appear to be inefficient at incorporating the azide functionality into
gangliosides (Figure 3B). It appears that the narrow substrate flexibility of ST3Gal5 in
CHO cells hinders their usage to study ganglioside-based interactions with metabolic
oligosaccharide engineering techniques.
The ST3Gal5 enzyme of BJAB cells shows a reduced capacity for engineering NeuGc-
containing gangliosides
When I cultured BJAB K20 cells with ManNGc, I observed that synthesis of the
corresponding GM3 analog (GM3-NeuGc) was considerably lower than BJAB K20 cells
cultured with ManNAc (Figure 3A). This trend appears to be consistent with the entire
α2,3-sialyltransferase family, as evidenced by MAA-lectin analysis (Figure 4A). One
possible explanation for these results is that BJAB K20 cells are inefficient at producing
adequate levels of CMP-NeuGc for sialoside synthesis. However, NeuGc engineering of
BJAB K20 cells into α2,6-linked glycoconjugates was observed to be higher than BJAB
K20 cells engineered with NeuAc (Figure 4B). This implies that the decreased synthesis
of α2,3-linked sialosides is not likely due to reduced production of CMP-NeuGc, since I
observed the synthesis of α2,6-linked NeuGc structures. Furthermore, the production of
NeuGc-containing sialosides in CHO cells was efficiently recognized by the MAA lectin,
suggested that the reduced level of MAA binding is not simply due to MAA’s inability to
87
recognize NeuGc-containing glycans. Indeed, previous reports have demonstrated that
SNA and MAA lectins do not discriminate between NeuAc and NeuGc.48 Instead, my
results point to an interpretation that human α2,3-sialyltransferases, specifically here
ST3Gal5, are able to utilize CMP-NeuGc as efficiently as α2,6-sialyltransferases. To
confirm my hypothesis, I am currently planning experiments to analyze the production of
CMP-NeuGc in BJAB K20 cells cultured with ManNGc. These cells will be harvested
for their nucleotides and analyzed by high performance anion exchange chromatography
(HPEAC) to quantify the synthesis of CMP-NeuGc. These results will be compared to
BJAB K88 and BJAB K20 cells cultured with ManNAc to determine if there are
adequate levels of CMP-NeuGc being produced. These results will help determine if the
production of GM3-NeuGc is hampered by inefficient synthesis of CMP-NeuGc or by the
substrate specificity of ST3Gal5. More direct measurements of ST3Gal5 specificity are
also underway.
Future directions
Overall, I observed that the human BJAB cells and hamster CHO cells display
significant differences in their ability to incorporate modified sialic acid into
gangliosides. These differences are extremely evident between ManNGc-, ManNAz- and
ManNDAz-treated cells. BJAB K20 cells are able to synthesize GM3-SiaNAz and GM3-
SiaDAz more efficiently than CHO Lec3 cells. Conversely, CHO Lec3 cells are able to
synthesize GM3-NeuGc more efficiently than CHO Lec3 cells. These results could be
caused from subtle structural variations in the catalytic domains and immediate
surrounding areas of ST3Gal5. Therefore, my hypothesis is BJAB K20 cells transfected
with choST3Gal5 would regain the ability to synthesize GM3-NeuGc and CHO Lec3
transfected with hsaST3Gal5 would obtain the ability to synthesize GM3-SiaNAz and
GM3-SiaDAz. To test this theory, I am currently planning to perform these
aforementioned transfections and analyze the production of GM3 by flow cytometry. To
confirm that the transfection of ST3Gal5 into these cells does not simply result in
overproduction of GM3, I am planning to perform control experiments where I transfect
the endogenous gene into each cell type. Due to the incomplete sequencing of the
hamster genome, I was unable to generate a mammalian expression plasmid for hamster
88
ST3Gal5; in its place, I have chosen to use the mouse ST3Gal5 for my investigations.
Additionally, this hypothesis will be tested by recombinantly expressing ST3Gal5
enzymes and performing in vitro experiments with unnatural sialic acid analogs.
Furthermore, it would be worthwhile to perform these experiments at different ManNAc
analog concentrations to see if the efficiency of metabolic oligosaccharide engineering
changes.
In conclusion, I have demonstrated that metabolic oligosaccharide engineering
techniques can be applied to introduce small perturbations into the N-acyl side chain of
the sialic acid residue of the GM3 ganglioside. My results also illustrate significant
differences in the incorporation of sialic acid analogs into GM3 in different cell lines; the
most notable difference was observed with α2,3-NeuGc incorporation. These differences
may reflect species-specific or cell type-specific differences. Further analyses are
currently underway to determine if the attenuated synthesis of GM3-NeuGc in BJAB K20
cells versus CHO cells is caused by an impairment of CMP-NeuGc synthesis or by
differences in ST3Gal5 specificity.
Acknowledgements
I would like to thank Michael Pawlita (German Cancer Research Center) and
James Paulson (The Scripps Research Institute) for sharing BJAB K20 and BJAB K88
cells, Mark Lehrman (UT Southwestern Medical Center) for sharing CHO cells, and
Pamela Stanley (Albert Einstein College of Medicine) for sharing CHO Lec3 cells. I
would like to thank Yan Li (UT Southwestern Medical Center Protein Chemistry
Technology Center) for mass spectrometry analysis of sugars, Parastoo Azadi and
Roberto Sonon (University of Georgia Complex Carbohydrate Research Center) for mass
spectrometry of ganglioside samples, and Angela Mobley (UT Southwestern Medical
Center Flow Cytometry Core Facility) for help with flow cytometry.
89
Methods
General information for the chemical synthesis of N-acetylmannosamine analogs
All chemicals were used as received from commercial suppliers without further
purification. 1-hydroxybenzotriazole hydrate was purchased from AnaSpec. N-butyric
anhydride, phenylacetic acid and propionic anhydride were purchased from TCI America.
All other chemicals were purchased from Sigma-Aldrich or Fisher Scientific unless
otherwise noted. Reaction progress was monitored by analytical thin layer
chromatography (TLC) on silica gel 60 F254 glass backed plates (Fisher) and stained with
ceric ammonium molybdate. Flash column chromatography was carried out with silica
gel 60 (particle size 40-63 µm, EMD Chemicals). All 1H-NMR and 13C-NMR spectra
were recorded on a Varian 500 MHz spectrometer and are reported in δ ppm scale. 1H-
NMR spectra were referenced to D2O (4.80 ppm) or CDCl3 (7.26 ppm). 13C-NMR
spectra were referenced to CDCl3 (77.23 ppm). ESI-MS data were collected at the UT
Southwestern Medical Center Protein Chemistry Technology Center. All acetylated
sugars were prepared as 10 mM stock solutions in ethanol. The purity of acetylated
sugars was confirmed by HPLC analysis before cellular treatments (spectra located in
appendix).
Synthesis of Ac4ManNAc
Ac4ManNAc was synthesized as previously reported.13 Briefly, to a solution of D-
(+)-N-acetylmannosamine (301.7 mg, 1.36 mmol) in pyridine (16.4 mL, 204 mmol),
acetic anhydride (4.72 mL, 54 mmol) was added and stirred overnight on ice. The
reaction mixture was diluted by CH2Cl2 and washed successively by 1.0 M HCl,
saturated sodium bicarbonate, and brine. The organic layer was dried over magnesium
sulfate and evaporated in vacuo. The residue was purified by flash chromatography
(hexanes / ethyl acetate gradient = 5/1, 3/1, 1/1) to afford Ac4ManNAc (282 mg, 53%,
mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.65 (3H, s), 2.02 (3H, s), 2.07
(3H, s), 2.11 (3H, s), 2.18 (3H, s), 4.10 (1H, dd, J = 2.3, 12.5), 4.28 (1H, t, J = 3.7), 4.78
(1H, ddd, J = 1.6, 3.9, 9.1), 5.06 (1H, d, 4.0), 5.13 (1H, t, J = 9.8), 5.33 (1H, d, J = 4.5),
5.79 (1H, d, J = 9.0), 5.86 (1H, d, J = 1.6). 13C-NMR (125 MHz, CDCl3): δ 20.88, 20.90,
90
20.92, 20.96, 20.97, 21.01, 21.08, 23.56, 23.65, 49.52, 49.74, 62.20, 65.41, 65.62, 68.99,
70.29, 71.56, 73.68, 90.86, 91.90, 168.34, 168.55, 169.92, 169.93, 170.23, 170.32,
170.73, 170.74, 170.82. ESI-MS for C16H23NO10 [M], calculated for 389.13, found
389.12. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the appendix.
Synthesis of Ac5ManNGc
Monoacetylated ManNGc was synthesized as previously reported.53 Briefly, to a
solution of D-(+)-mannosamine hydrochloride (216 mg, 1.00 mmol) and sodium
bicarbonate (1.68 g, 20 mmol) in water (8.6 mL, 480 mmol) chilled on ice, acetoxyacetyl
chloride (537 µL, 5.00 mmol) was added dropwise and the reaction was stirred for 3
hours on ice, monitoring reaction progress by TLC (ethyl acetate / acetic acid / water =
3/2/1) using nihydrin to detect unreacted starting material and orcinol-sulfuric acid to
detect sugars. After filtering through Celite in a glass Pasteur pipette, the filtrate was
neutralized with 1.0 M HCl (pH ~ 7, added dropwise). The resulting mixture was
concentrated in vacuo, roughly purified by flash chromatography (ethyl acetate /
isopropanol / water = 27/8/4), and used directly to synthesize the fully acetylated product,
Ac5ManNGc. The acetylation of monoacetylated ManNGc was performed by the same
procedure described in the synthesis of Ac4ManNAc to afford Ac5ManNGc (159 mg,
36%, mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.97 (3H, s), 2.03 (3H, s),
2.07 (3H, s), 2.15 (3H, s), 2.18 (3H, s), 4.03 (1H, dd, J = 1.9, 12.6), 4.23 (1H, d, J = 4.7),
4.58 (2H, s), 4.65 (1H, ddd, J = 1.9, 4.4, 9.0), 5.04 (1H, d, J = 3.8), 5.14 (1H, t, J = 10.3),
5.30 (1H, d, J = 4.3), 6.01 (1H, d, J = 1.3), 6.37 (1H, d, J = 9.2). 13C-NMR (125 MHz,
CDCl3): δ 20.84, 20.86, 20.87, 20.87, 20.89, 20.91, 20.96, 21.06, 49.16, 49.61, 61.96,
62.04, 63.25, 63.30, 65.15, 65.26, 68.99, 70.31, 71.40, 73.63, 90.62, 91.63, 167.41,
168.00, 168.32, 168.53, 169.53, 169.58, 169.75, 169.79, 170.29, 170.32, 170.61, 170.62.
ESI-MS for C18H25NO12 [M], calculated for 447.14, found 447.15. 1H-NMR, 13C-NMR,
and ESI-MS spectra are presented in the appendix.
Synthesis of Ac4ManNProp
ManNProp was synthesized as previously reported.52 Briefly, to a solution of D-
(+)-mannosamine hydrochloride (300 mg, 1.39 mmol) in MeOH (20 mL) and 3.0 M
91
NaOH (0.5 mL), propionic anhydride (1.0 mL, 7.80 mmol) was added dropwise on ice
while stirring for several hours, monitoring reaction progress by TLC. After completion,
1.0 M HCl was added dropwise to neutralize the solution (pH ~ 7). After the solvent was
evaporated in vacuo, dried with several washes of toluene, filtered with cotton in Pasteur
pipette, and evaporated under vacuum. The resulting mixture was roughly purified by
flash chromatography (CH2Cl2 / MeOH = 1/0, 10/1, 4/1) and used directly to synthesize
the acetylated product, Ac4ManNProp. The acetylation of ManNProp was performed by
the same procedure described in the synthesis of Ac4ManNAc to afford Ac4ManNDAz
(284 mg, 51%, mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.19 (3H, t, J =
7.9), 2.00 (3H, s), 2.07 (3H, s), 2.11 (3H, s), 2.19 (3H, s), 2.34 (2H, m), 4.06 (1H, dd, J =
1.4, 12.0), 4.28 (1H, t, J = 4.8), 4.80 (1H, ddd, J = 1.6, 4.4, 9.2), 5.06 (1H, d, J = 4.0),
5.18 (1H, t, J = 10.3), 5.22 (1H, d, J = 4.5), 5.65 (1H, d, J = 9.3), 6.04 (1H, d, J = 4.8). 13C-NMR (125 MHz, CDCl3): δ 9.92, 10.11, 20.88, 20.90, 20.92, 20.94, 20.95, 20.99,
21.09, 29.84, 29.99, 49.28, 49.54, 53.64, 54.65, 62.07, 62.19, 65.40, 65.58, 69.06, 70.26,
71.56, 73.62, 90.88, 91.93, 168.36, 168.53, 169.90, 170.22, 170.29, 170.72, 173.94,
174.59. ESI-MS for C17H25NO10 [M-H]-, calculated for 402.15, found 402.15. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the appendix.
Synthesis of Ac4ManNBut
ManNBut was synthesized as previously reported.52 Briefly, to a solution of D-
(+)-mannosamine hydrochloride (300 mg, 1.39 mmol) in MeOH (5.0 mL) and 3.0 M
NaOH (0.5 mL), butyric anhydride (1.0 mL, 6.13 mmol) was added dropwise on ice
while stirring for several hours, monitoring reaction progress by TLC. After completion,
1.0 M HCl was added dropwise to neutralize the solution (pH ~ 7). After the solvent was
evaporated in vacuo, dried with several washes of toluene, filtered with cotton in Pasteur
pipette, and evaporated under vacuum. The resulting mixture was roughly purified by
flash chromatography (CH2Cl2 / MeOH = 1/0, 10/1, 4/1) and used directly to synthesize
the acetylated product, Ac4ManNBut. The acetylated of ManNBut was performed by the
same procedure described in the synthesis of Ac4ManNAc to afford Ac4ManNBut (266
mg, 45%, mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.00 (3H, t, J = 7.3), 1.71
(2H, m), 2.01 (3H, s), 2.07 (3H, s), 2.11 (3H, s), 2.19 (3H, s), 2.25 (2H, t, J = 7.5), 4.06
92
(1H, dd, J = 4.9), 4.28 (1H, t, J = 4.8), 4.68 (1H, ddd, J = 4.6, 9.5, 12.6), 5.06 (1H, d, J =
4.0), 5.18 (1H, t, J = 10.3), 5.33 (1H, d, J = 4.4), 5.65 (1H, d, J = 9.4), 6.04 (1H, s). 13C-
NMR (125 MHz, CDCl3): δ 13.72, 13.82, 19.3, 19.49, 20.88, 20.90, 20.93, 20.95, 21.10,
38.73, 38.90, 49.24, 49.56, 62.06, 62.89, 65.37, 65.55, 69.09, 70.29, 71.60, 73.65, 90.84,
91.94, 168.37, 168.51, 169.88, 170.22, 170.72, 173.16, 173.80. ESI-MS for C18H27NO10
[M-H]-, calculated for 416.16, found 416.17. 1H-NMR, 13C-NMR, and ESI-MS spectra
are presented in the appendix.
Synthesis of Ac4ManNPhAc
To a solution of phenylacetic acid (136 mg, 1.00 mmol), D-(+)-mannosamine
hydrochloride (216 mg, 1.00 mmol) and triethylamine (280 µL, 2.00 mmol) in MeOH (10
mL), 1-ethyl-3-(3-dimethyllaminopropyl)carbodiimide hydrochloride (388 mg, 2.00
mmol) was added. The reaction mixture was stirred on ice for 10 minutes, followed by
stirring at room temperature overnight. The resulting mixture was concentrated in vacuo,
roughly purified by flash chromatography (CH2Cl2 / MeOH = 1/0, 10/1, 4/1), and used
directly to synthesize the acetylated product, Ac4ManNPhAc. The acetylation of
ManNPhAc was performed by the same procedure described in the synthesis of
Ac4ManNAc to afford Ac4ManNPhAc (117 mg, 25%, mixture of anomers). 1H-NMR
(500 MHz, CDCl3): δ 1.94 (3H, s), 2.01 (3H, s), 2.04 (3H, s), 2.16 (3H, s), 3.99 (1H, ddd,
J = 2.2, 6.0, 13.0), 4.06 (1H, dd, J = 2.7, 9.7), 4.66 (1H, ddd, J = 4.3, 9.4, 13.7), 4.97 (2H,
s), 4.99 (1H, t, J = 5.6), 5.02 (1H, d, J = 3.7), 5.28 (1H, d, J = 4.4), 5.63 (1H, t, J = 9.0),
5.96 (1H, d, J = 1.6), 7.34 (3H, m), 7.42 (2H, m). 13C-NMR (125 MHz, CDCl3): δ 20.81,
20.83, 20.84, 20.86, 20.89, 20.92, 21.04, 43.80, 43.99, 49.43, 49.46, 61.94, 61.96, 65.19,
65.24, 69.21, 70.16, 71.35, 73.37, 90.58, 91.71, 127.63, 127.78, 129.22, 129.37, 129.41,
129.49, 134.48, 168.33, 168.36, 169.65, 169.70, 170.20, 170.24, 170.68, 170.70, 171.35.
ESI-MS for C22H27NO10 [M], calculated for 465.16, found 465.16. 1H-NMR, 13C-NMR,
and ESI-MS spectra are presented in the appendix.
Synthesis of Ac4ManNAz
ManNAz was synthesized as previously reported.48 Briefly, to a solution of
azidoacetic acid (360 mg, 3.00 mmol), D-(+)-mannosamine hydrochloride (432 mg, 2.00
93
mmol) and triethylamine (560 µL, 4.00 mmol) in MeOH (20 mL), 1-ethyl-3-(3-
dimethyllaminopropyl)carbodiimide hydrochloride (766 mg, 4.00 mmol) and 1-
hydroxybenzotriazole hydrate (270 mg, 2.00 mmol) were added. The reaction mixture
was stirred on ice for 10 minutes, followed by stirring at room temperature overnight.
The resulting mixture was concentrated in vacuo, roughly purified by flash
chromatography (CH2Cl2 / MeOH = 1/0, 5/1, 3/1), and used directly to synthesize the
acetylated product, Ac4ManNAz. The acetylation of ManNAz was performed by the
same procedure described in the synthesis of Ac4ManNAc to afford Ac4ManNAz (437
mg, 50%, mixture of anomers). 1H-NMR (500 MHz, CDCl3): δ 1.96 (3H, s), 2.03 (3H, s),
2.08 (3H, s), 2.15 (3H, s), 4.02 (2H, s), 4.09 (1H, ddd, J = 2.0, 8.0, 14.4), 4.20 (1H, d, J =
1.8), 4.58 (1H, ddd, J = 4.2, 9.3, 13.5), 5.04 (1h, d, J = 3.9), 5.19 (1H, t, J = 10.1), 5.20
(1H, d, J = 4.3), 6.01 (1H, s), 6.65 (1H, d, J = 18.1). 13C-NMR (125 MHz, CDCl3): δ
20.80, 20.84, 20.87, 20.91, 20.95, 20.98, 21.06, 49.44, 49.90, 52.55, 52.74, 61.84, 61.93,
65.08, 65.27, 69.02, 70.41, 71.62, 73.55, 90.42, 91.47, 166.94, 167.52, 168.30, 168.54,
169.76, 170.30, 170.35, 170.73. ESI-MS for C16H22N4O10 [M], calculated for 430.13,
found 430.14. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the appendix.
Synthesis of Ac4ManNDAz
ManNDAz was synthesized as previously reported.10 Briefly, to a solution of 4,4-
azo-pentanoic acid27 (128 mg, 1.00 mmol), D-(+)-mannosamine hydrochloride (216 mg,
1.00 mmol) and triethylamine (278 µL, 2.00 mmol) in MeOH (10 mL), 1-ethyl-3-(3-
dimethyllaminopropyl)carbodiimide hydrochloride (383 mg, 2.00 mmol) and 1-
hydroxybenzotriazole hydrate (135 mg, 1.00 mmol) were added. The reaction mixture
was stirred on ice for 10 minutes, followed by stirring at room temperature overnight.
The resulting mixture was concentrated in vacuo, roughly purified by flash
chromatography (CH2Cl2 / MeOH gradient = 1/0, 10/1, 4/1), and used directly to
synthesize the acetylated product, Ac4ManNDAz. The acetylation of ManNDAz was
performed by the same procedure described in the synthesis of Ac4ManNAc to afford
Ac4ManNDAz (84 mg, 28% over two steps, mixture of anomers). 1H-NMR (500 MHz,
CDCl3): δ 1.06 (3H, s), 1.81 (2H, m), 2.02 (3H, s), 2.07 (3H, s), 2.12 (3H, s), 2.19 (3H,
s), 4.05 (2H, s), 4.09 (1H, ddd, J = 6.2, 18, 30.5), 4.29 (1H, t, J = 4.5), 4.78 (1H, dd, J =
94
2.3, 8.2), 5.06 (1H, d, J = 4.0), 5.20 (1H, t, J = 9.8), 5.32 (1H, d, J = 4.4), 5.80 (1H, d, J =
9.0), 6.04 (1H, s). 13C-NMR (125 MHz, CDCl3): δ 20.18, 20.19, 20.85, 20.88, 20.91,
20.95, 20.97, 21.08, 25.50 25.54, 29.94, 30.05, 30.71, 30.84, 49.55, 49.76, 62.02, 62.15,
65.32, 69.06, 70.32, 71.57, 73.67, 90.80, 91.79, 168.33, 168.53, 169.82, 169.91, 170.20,
170.28, 170.75, 170.78, 171.62, 172.14. ESI-MS for C19H27N3O10 [M], calculated for
457.17, found 457.16. 1H-NMR, 13C-NMR, and ESI-MS spectra are presented in the
appendix.
Cell culturing experiment reagents
RPMI 1640 with 2 mM L-glutamine, α-minimum Eagle’s medium with
glutamine, ribonucleosides, and deoxyribonucleosides, Opti-MEM, fetal calf serum,
penicillin/streptomycin, dPBS, PBS (pH = 7.4), FITC-streptavidin, aminooxy-biotin, and
propidium iodide were purchased from Invitrogen. Nutridoma SP and BSA Fraction V
were purchased from Roche Applied Science. Aniline was purchased from Sigma-
Aldrich. Glycerol was purchased from Fisher Scientific. SNA-FITC was purchased from
EY Labs. MAA-biotin was purchased from Vector Labs. DTAF-streptavidin was
purchased from Jackson Immunoresearch. Cell counting was performed on the
Invitrogen Countess Automated Cell Counter. Flow cytometry experiments were
performed on a BD Biosciences FACSCaliber flow cytometer
Cell culturing conditions
BJAB K20 and K88 cells were grown and maintained in RPMI 1640 with 2 mM
L-glutamine containing 10% fetal calf serum, 100 U/ml penicillin, and 100 µg/ml
streptomycin at 37oC, 5% CO2 in a water-saturated environment. Cells were cultured at
2.5 x 105 cells/mL in media and grown for 48 hours before passaging. Typically, cell
densities were maintained between 2.5 x 105 cells/mL and 2.0 x 106 cells/mL. Cell
viability was analyzed using Trypan blue dye staining with the Countess Automated Cell
Counter instrument.
CHO and CHO Lec3 cells were grown and maintained in α-minimum Eagle’s
medium w/ glutamine, ribonucleosides, and deoxyribonucleosides containing 10% fetal
calf serum at 37oC, 5% CO2 in a water-saturated environment. Cells were cultured at 2.5
95
x 104 cells/mL in media and grown for 72 hours before passaging. Typically, cell
densities were maintained between 2.5 x 104 cells/ml and 2.0 x 106 cells/mL.
Serum free exposure and supplementation with monosaccharides
Initially, CHO and CHO Lec3 cells were plated at 5.0 x 104 cells/ml and allowed
to grow for 24 hours. Cells were then washed 3 times with PBS, then cultured with
OptiMEM media for serum free exposure. Using 10 mM ethanol stocks of each sugar,
ethanol or acetylated sugar was added to each plate while swirling to make a final
concentration of 100 µM. After growth in the presence of the appropriate
monosaccharides for 48 hours, the cells were removed from the plate by exposure to
trypsin for 5 minutes, counted, and harvested by centrifugation at 220g for 5 min in 50 ml
conical tubes.
To generate serum free conditions for BJAB K20 and K88 cells, the cells were
grown in RPMI 1640 with 2 mM L-glutamine containing 1x Nutridoma SP, 50 U/ml
penicillin, and 50 µg/ml streptomycin. Cells were cultured for two passages at 2.5 x 105
cells/ml in media for 72 hours at a time before supplementation with monosaccharides.
Prior to the addition of cells to a tissue culture plates, acetylated sugar or ethanol were
added and the ethanol was pre-evaporated. After cell counting, BJAB cells were plated at
2.5 x 105 cells/ml in serum free media for each monosaccharide condition. After growing
for 72 hours, cells were counted and harvested by centrifugation at 220g for 5 min in 50
ml conical tubes.
To ensure consistent results among all samples, equal numbers of cells were
collected for every samle; the toal number of cells collected for an experiment ranged
between 3.0 – 4.0 x 107 cells for CHO/CHO Lec3 analysis and 1.0 – 1.2 x 108 cells for
BJAB K88/K20 analysis. Cell pellets were stored at -80 oC overnight before proceeding
to ganglioside extraction.
Extraction of gangliosides - Total Lipid Extraction
Cell pellets were thawed to room temperature, resuspended with 300 µl of ice
cold ddH2O (W), and dounced 50 times with a Kontes tissue grinder, tube size 20. With
96
a glass Pasteur pipette and a 2 mL rubber bulb, the cell lysate suspension was transferred
into a 4 mL glass vial containing 800 µL of methanol (M), already stirring. 400 µL of
chloroform (C) was added to the vial and the mixture was stirred thoroughly for 2 hours
at room temperature. Samples were covered in foil to prevent exposure to light. After
stirring, the mixture was transferred by a glass Pasteur pipette into a 13 x 100 mm glass
culture tube and centrifuged at 2800g for 10 min @ 30 oC. The supernatant (containing
the total lipid extract) was transferred by a glass Pasteur pipette into a new 4 mL glass
vial and evaporated to dryness under N2 gas.
Extraction of gangliosides - Phospholipid Extraction
The dried total lipid extract was resuspended with 800 µL butanol and 1200 µL
diisopropyl ether and sonicated in a water bath for 10 minutes. The resuspended lipids
were then transferred into a 13 x 100 mm glass culture tube using a glass Pasteur pipette.
To extract undesired phospholipids from the mixture, 1000 µl of 50 mM NaCl was added
to the tube and mixed vigorously by pipetting up and down repeatedly with a glass
Pasteur pipette. The mixture was then centrifuged at 2800g for 10 min at 30 oC to
separate the two phases. Using a glass Pasteur pipette, the organic phase (top layer) was
carefully removed. The aqueous mixture (bottom layer) was then extracted two more
times using the same ratio of butanol and diisopropyl ether.
Extraction of gangliosides - SepPak purification
After the final extraction, the remaining lipid mixture was loaded onto a SepPak tC18
column, 0.3g size. The column was first pre-treated with three 2 mL washes of C/M/W
(2:43:55) followed by two 2 mL washes of C/M (1:1) and ending with three more 2 mL
washes of C/M/W (2:43:55). After loading of the sample, the column was washed three
times with 2 mL of C/M/W (2:43:55) followed by three 2 mL washes of C/M (1:1) to
desalt the sample and remove unwanted contaminants. Elution of gangliosides was
achieved using 2 mL of 100 % methanol. Ganglioside extracts were then transferred into
a new 4 mL glass vial and evaporated to dryness under N2.
97
HPTLC Analysis of Extracted Gangliosides
Extracted ganglioside samples were redissolved with 30 µL C/M/W (2:1:0.1) and
resolved on HPTLC plates. Ganglioside standards were loaded onto the plate to provide
mass references. Gangliosides were separated with chloroform:methanol:0.2% CaCl2(aq)
(80:45:10) as the running buffer. HPTLC plates were first pre-run before loading 10 µL
of ganglioside extract. After thoroughly drying the plate in a fume hood, gangliosides
were detected by resorcinol staining (0.1% resorcinol, 0.04% CuSO4 in hydrochloric
acid:water [4:1]). Plates were imaged using an Alpha Innotech FluorChem HD2 and
images were processed using Adobe Photoshop.
Mass Spectrometry Analysis of Extracted Gangliosides
Dried ganglioside extracts were sent to the Complex Carbohydrate Research
Center at the University of Georgia for mass spectrometry analysis. To analyze the
overall composition of extracted gangliosides, MALDI-TOF-MS was performed.
Samples were crystallized onto a MALDI plate with trihydroxyacetophenone
monohydrate (THAP) as a matrix. Analysis of gangliosides was performed in the
negative ion mode using a Bruker microflex instrument.
Flow cytometry - Lectin
Cells were resuspended at 1.875 x 106 cells/ml in PBS and aliquoted into a v-
bottom 96-well plate in 200 µl amounts. Analyses were performed in triplicate with three
separate cultures of each condition. Cells were washed 3 times with 200 µL of 0.1%
BSA/PBS and centrifuged at 2,000 rpm, 4 oC for 4 minutes. For MAA binding
experiments, the cells were incubated with 50 µL of 10 µg/ml of MAA-biotin in 0.1%
BSA/PBS for 30 minutes on ice. After being washed 3 times with 200 µL of 0.1%
BSA/PBS, the cells were incubated with 50 µL of 20 µg/ml of streptavidin-FITC for 30
min on ice. After being washed 3 times with 200 µL of 0.1% BSA/PBS, the cells were
resuspended in 400 µl of 0.1% BSA/PBS. Propidium iodide, used to identify dead cells
versus live cells, was added at a final concentration of 50 µg/ml into each tube before
analysis. The cells were analyzed using a FACSCaliber flow cytometer. Live cells
98
(10,000 cells/sample) were identified by their forward scatter versus side scatter plot; all
propidium iodide positive cells (dead cells) were excluded from analysis. FITC
fluorescence was measured on the FL-1 channel of the instrument.
For SNA binding experiments, the cells were incubated with 10 µg/ml of SNA-
FITC in 0.1% BSA/PBS for 30 minutes on ice. After being washed three times with
0.1% BSA/PBS, the cells were resuspended in 400 µl of 0.1% BSA/PBS. Propidium
iodide, used to identify dead cells versus live cells, was added at 50 µg/ml into each tube
before analysis.
Flow cytometry - PAL
Cells were resuspended at 1.0 x 106 cells/ml in PBS and aliquoted into a v-bottom
96-well plate in 200 µl amounts, in biological triplicate. Cells were washed 2x with PBS
(pH = 7.4) and centrifuged at 2,000 rpm, 4oC for 4 minutes. Cells were then resuspended
in 200 µl 1.0 mM NaIO4/PBS (pH = 7.4) and incubated for 30 min on ice. After
incubation, 50 µl of 5.0 mM glycerol/PBS (pH = 7.4) was added and incubated for 30
min on ice to quench the oxidation reaction. Cells were pelleted by centrifugation at
2,000 rpm, 4oC for 4 min to remove the supernatant and resuspended with 200 µl of 5.0%
FBS/PBS (pH = 6.7). The cells were then washed 2 times with 5.0% FBS/PBS (pH =
6.7) and centrifuged at 2,000 rpm, 4 oC for 4 minutes. After washing, the cells were
resuspended in 200 µl of 0.1 mM aminooxy-biotin, 10.0 mM aniline 5.0 % FBS/PBS (pH
= 6.7) and incubated for 90 min on ice. After incubation, cells were washed three times
with 5.0% FBS/PBS (pH = 6.7) and centrifuged at 2,000 rpm, 4oC for 4 minutes. After
washing, the cells were resuspended in 200 µl of 3.2 µg/ml DTAF-Streptavidin 5.0%
FBS/PBS for 30 min on ice. Cells were then washed three times with 5.0% FBS/PBS
(pH = 6.7) and centrifuged at 2,000 rpm, 4oC for 4 minutes. After being washed three
times with 0.1% BSA/PBS, the cells were resuspended in 400 µl of 5.0% FBS/PBS (pH =
6.7). The cells were analyzed using a FACSCaliber Flow Cytometer. Live cells (10,000
cells/sample) were identified by their forward scatter versus side scatter plot. DTAF
fluorescence was measured on the FL-1 channel of the instrument.
99
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Appendix
106
1H NMR Data
107
108
109
110
13C NMR Data
111
112
113
114
ESI-MS Data
115
116
117
118
HPLC Data
119
120
121
122
MALDI-TOF-MS Data
123
124
125
126
127
128