The Investigation of Peptide and Protein-Glycosaminoglycan Binding

Interactions using Fluorescent Probes

By

Anthony F. Rullo

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Graduate Department of Chemistry

University of Toronto

Copyright by Anthony F. Rullo 2012

ii

Abstract

The Investigation of Peptide and Protein-Glycosaminoglycan Binding

Interactions using Fluorescent Probes

Doctor of Philosophy

Graduate Department of Chemistry

University of Toronto

Anthony F. Rullo 2012

The structural complexity of glycosaminoglycans (GAGs) such as heparin and heparan sulfate

(HS) and their numerous biological roles, brings forth the need to develop new methods, capable

of studying GAGs and their interactions with peptides and proteins under native settings. This

thesis explores the development of chemical tools to study heparin/HS binding interactions under

physiologically relevant conditions using fluorescence. In chapter 2, we designed peptide-based

quinolinium probes to study the structural requirements of cationic peptides required for high

affinity peptide-heparin interactions. These fluorescent probes enabled the study of peptide-

heparin interactions at nM concentrations allowing the calculation of peptide-heparin binding

constants. It was observed that peptides with positive charge displayed on one face of an α-helix

in a continuous arrangement bound to heparin with the highest affinity and that heparin likely

prefers to bind to these peptides while remaining in an extended conformation.

In chapter 3, we set out to study an important biological role of HS which involves the binding

and sequestering of proteins at the cell surface, facilitating endocytosis. HS has been implicated

iii

in the mechanism of cell penetrating peptide (CPP) cell uptake, with different CPPs showing

different degrees of HS dependence on uptake as well as different mechanisms of entry. The role

of HS in the mechanism of CPP uptake was investigated in chapter 3 using fluorescent peptide-

based probes incorporating fluorophore/quencher pairs. These were used to identify and

characterize the ability of heparin/HS to bind and cluster with CPPs to form colloidally stable

aggregates. It was shown that the CPP Antp formed much more stable clusters with heparin than

the TAT peptide despite both peptides having similar binding affinity for a single heparin chain.

These findings were used to explain the cell surface HS dependence of Antp on cell uptake via

endocytosis in contrast to the low dependance of TAT on HS and its uptake via translocation. A

general model relating the ability of a CPP to cluster surface HS to its preferred mechanism of

cell entry was proposed. In chapter 4, a strategy to selectively, and site specifically acylate

carbohydrate binding proteins was developed using thioester-based affinity conjugates. It was

possible to label maltose binding protein, a periplasmic protein, with high yield and selectivity at

a single lysine residue proximal to the maltose binding site. Selective protein labeling could be

carried out in bacterial cell extracts and in live bacterial cells. This strategy can potentially be

applied to develop protein-based carbohydrate biosensors as well as profile carbohydrate binding

proteins in biological samples.

iv

Acknowledgments

I would sincerely like to thank Professor Mark Nitz for the incredible opportunity to pursue my

doctoral studies under his supervision. Your mentorship and scientific ability has had a very

positive and profound impact on my development as a graduate student and on my transition

from a student to a young scientist. I am also very thankful to the members of my committee

professor Rebecca Jockusch and professor Andrew Woolley for their invaluable advice and

support.

Thank you to the members of the Nitz group and especially to Rodolfo Gomez. I have very

much enjoyed our numerous insightful discussions ranging from kinetics and organic synthesis

to the everyday trials and tribulations of life as a doctoral student. I wish you the best of luck in

your future endeavors and I know that you will always be ready to replace luck with impressive

perseverance and technical skill should the situation require.

A special thanks to my friend Andrew Beharry who’s ability as a doctoral student and passion

for science has been an inspiration throughout my studies at the University of Toronto. It has

been a pleasure developing as a doctoral student in chemistry alongside you and I very much

look forward to being a colleague of yours in the future.

The work presented in this thesis is the culmination of five years of dedication, dogged

determination, and devotion and is a testament to the special people in my life who introduced

me to these virtues. My dad and first mentor John, who personifies strength and discipline,

taught me at an early age the importance of staying the course and the meaning of endurance. I

learned by his example to never quit and to persevere in the face of adversity. Anything I ever

have achieved or will achieve in life is attributed to him and his example. My mother Anna’s

selflessness and sacrifice throughout the years made this PhD possible. I will be forever indebted

to you and cannot describe with words the extent of my gratitude and appreciation. To my Amy,

your love and support has been an integral part of my doctoral studies. You have been on the

front line alongside me throughout this entire process from the frequent frustrations, to the joy of

making exciting discoveries and positive contributions to the field. I can’t imagine having done

this without you.

v

I would also like to give a special acknowledgment to my nanna Serafina. I could never do

justice in describing the magnitude of the significance you have had in my life and the quality of

the human being you are by simply putting black letters down on a white page. Through the

many adventures of growing up, I could never have hoped for a more nurturing, loyal, and noble

protector and friend than the one God gave me as a grandmother through you. Your devotion to

your grandchildren was not in vain and is largely responsible for the many successes we have all

enjoyed including the attainment of my doctoral degree in chemistry and the work presented in

this thesis. It is with true sadness I bring this chapter of my life to a close without being able to

share it with you and without you deriving the pleasure from it that you, most of all, deserved to

enjoy. You will serve as an inspiration to me always and will forever be a critical part of the

foundation upon which I am built. This thesis is dedicated to you.

vi

Table of Contents

Abstract……………………………………………………………………………………. ii

Acknowledgments…………………………………………………………………………. iv

Table of Contents…………………………………………………………………………. vi

List of Abbreviations……………………………………………………………………… viii

List of Tables………………………………………………………………………………. x

List of Figures……………………………………………………………………………... xi

List of Schemes……………………………………………………………………………. xiv

Chapter 1. Introduction………………………………………………………………….. 1

1.1 Carbohydrates in glycobiology…………………………………………………………. 1

1.2 Glycosaminoglycans……………………………………………………………………. 3

1.3 Heparin and heparan sulfate……………………………………………………………. 5

1.4 Lectins………………………………………………………………………………….. 6

1.5 Lectin-carbohydrate interactions……………………………………………………….. 8

1.6 Heparin/HS-Protein Interactions……………………………………………………….. 10

1.7 Techniques to study HS/heparin interactions…………………………………………... 15

1.8 The development of probes for HS/heparin……………………………………………. 16

1.9 Purpose of study……………………………………………………………………….. 22

References………………………………………………………………………………….. 23

Chapter 2. The effects of the spatial display of positive charge on peptide

heparin binding affinity…………………………………………………………………...

27

2.1 Introduction……………………………………………………………………………. 27

2.2 Results………………………………………………………………………………….. 30

2.3 Discussions and Conclusions…………………………………………………………… 41

2.4 Materials and Methods…………………………………………………………………. 43

References………………………………………………………………………………….. 47

Chapter 3. Peptide-Glycosaminoglycan Cluster Formation

Involving Cell Penetrating Peptides………………………………………………………

50

3.1 Introduction…………………………………………………………………………….. 50

3.2 Results………………………………………………………………………………….. 53

Heparin sepharose chromatography…………………………………………….. 54

Dynamic light scattering………………………………………………………… 54

vii

Cluster stability by fluorescence spectroscopy………………………………….. 55

3.3 Discussions and Conclusions…………………………………………………………… 62

3.4 Materials and methods………………………………………………………………….. 66

References………………………………………………………………………………….. 70

Chapter 4. The synthesis of thioester carbohydrate

conjugates as selective lectin labelling agents............................................................

75

4.1 Introduction…………………………………………………………………………….. 75

4.2 Results and Discussion…………………………………………………………………. 76

Synthesis of thioester constructs………………………………………………… 76

MBP labeling in vitro…………………………………………………………… 77

MBP labeling in cellular extracts and live cells………………………………… 87

4.3 Concluding remarks…………………………………………………………………….. 91

4.4 Materials and Methods…………………………………………………………………. 92

References………………………………………………………………………………….. 100

Chapter 5. Summary and Perspective…………………………………………………… 103

5.1 The importance of the spatial display of charge in peptide/protein interactions

with GAGs…………………………………………………………………………………..

104

5.2 Peptide-Cluster formation involving cell penetrating peptides………………………… 105

5.3 The synthesis of thioester carbohydrate conjugates as selective lectin

labelling agents..........................................................................................................

108

5.4 Future directions……………………………………………………………………….. 111

References………………………………………………………………………………….. 113

Appendix…………………………………………………………………………………… 116

viii

List of Abbreviations

ATIII antithrombin III

CD Circular Dichroism

ConA concavalin A

CPP Cell Penetrating Peptide

DABCYL dimethylaminoazocarboxylic acid

DCC Dicyclohexylcarbodiimide

DIEPA Diisopropylethylamine

DLS Dynamic light scattering

DMF Dimethylformamide

E.coli Escherichia coli

ESI electrospray ionization

FN Fibronectin

FPLC fast protein liquid chromatography

FRET Fluorescence resonance energy transfer

GAG Glycosaminoglycan

Gal D-Galactose

GalNAc N-acetyl-D-galactosamine

GFP Green fluorescent protein

Glc D-Glucose

GlcN D-Glucosamine

GlcNAc N-acetyl-D-glucosamine

GPI Glucosylphosphatidylinositol

HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HHBP helical heparin binding peptide

HOBt N-hydroxybenzotriazole

HPLC High Pressure liquid chromatography

HS Heparan Sulfate

IPTG Isopropyl β-D-1-thiogalactopyranoside

ITC Isothermal titration calorimetry

KD Equilibrium dissociation constant

LB lysogeny broth

Man Mannose

MALDI-TOF matrix assisted laser desorption ionization-time of flight

MS mass spectrometry

ix

m/z mass/charge

NA N-acetylated

NMR Nuclear Magnetic Resonance

NS N-sulfated

SPR surface Plasmon resonance

TFA Trifluoroacetic acid

TIS Triisopropylsilane

VEGF vascular endothelial growth factor

WGA wheat germ agglutinin

Xyl Xylose

x

List of Tables

Table 1-1. A summary of the different classes of animal lectins…………………………………….. 8

Table 1-2. Titrations of thrombin with heparin oligosaccharides of increasing length………………. 12

Table 2-1. Sequences of Synthesized Heparin Binding Peptides........................................................... 34

Table 2-2. Experimentally Determined Dissociation Constants and

Structural Binding Characteristics of Seven Synthetic Peptides.............................................................

37

Table 3-1. The sequences of the three cationic heparin binding peptides used in this study……….. 53

xi

List of Figures

Figure 1-1. The structures of a glycoglycerolipid (left) and glycosphingolipid (right)…………………. 2

Figure 1-2. The three structures of the protein-carbohydrate linkages found in glycoproteins

N-linked (Top left), O-linked (Bottom left), GPI anchors (right)………………………………………..

3

Figure 1-3. The chemical structures of common members of the glycosaminoglycan family………. 4

Figure 1-4. A schematic representation outlining the biosynthesis of heparin and HS……………….. 6

Figure 1-5. A schematic representation of the polyelectrolyte effect…………………………………. 10

Figure 1-6. The structure of the specific pentasaccharide sequence on heparin recognized

by AT III. The critical sulfate at position O-3 is represented by R2. R = H R

2 = H or SO3

-.

RE = reducing terminus NRE = non reducing terminus………………………………………………...

12

Figure 1-7. An illustration of the reduction in dimensionality experienced by ATIII

and thrombin in the presence of heparin…………………………………………………………………

13

Figure 1-8. A scheme representing the proposed mechanism of heparin rate enhanced

AT III inhibition of thrombin due to ternary complex formation……………………………………….

13

Figure 1-9. A schematic representation of the heparin catalyzed conformational change of

fibronectin(FN)……………………………………………………………………………………………

14

Figure 1-10. A synthetic receptor approach to heparin probe design…………………………………. 19

Figure 1-11. The chemical structure of the quinolinium tethered heparin probe.

R = the α-carbon of an amino acid……………………………………………………………………….

19

Figure 1-12. The coiled coil motif applied as a ratiometric probe for heparin.

The cationic heparin binding site containing lysine residues at the e and g positions

are indicated by arrows…………………………………………………………………………………...

20

Figure 1-13. The selectivity of the probe for heparin over other polyelectrolytes…………………….. 21

Figure 1-14. Self-assembling fluorescent receptors for GAGs…………………………………………. 22

Figure 2-1. The preferred conformations of glucosamine and uronic acid residues within the

heparin/HS chain in aqueous solution.........................................................................................................

28

Figure 2-2. The proposed mechanism of quinolinium probing of peptide-heparin

binding interactions……………………………………………………………………………………….

30

Figure 2-3. Helical wheel and ribbon-ball models of the five helical heparin binding peptides

synthesized presented as α-helices..............................................................................................................

31

Figure 2-4. Fluorescence Titration of peptide 1 with heparin, 50 µM peptide 1 titrated with heparin 50

mM phosphate buffer 150 mM NaCl pH 7.................................................................................................

33

xii

Figure 2-5. CD spectra of seven heparin binding peptides (75 µM) with/without heparin (500 µM)

(50 mM phosphate buffer 150 mM NaCl pH 7...........................................................................................

36

Figure 2-6. Binding isotherms of synthesized peptides to heparin.......................................................... 38

Figure 2-7. Schematic representation of the proposed 1Gap peptide interaction with heparin............. 39

Figure 2-8. Peptide 1 and 1Gap titrations with heparin 14 mer oligosaccharide

(50 mM phosphate buffer pH 7, 150 mM NaCl…………………………………………………………

40

Figure 2-9. Nonlinear least squares analysis of binding of peptide 1R (25 nM) with heparin

(50 mM phosphate buffer, 150 mM NaCl, pH 7)........................................................................................

40

Figure 2-10. The synthesis of quinolinium-peptide conjugates................................................................. 44

Figure 3-1. A schematic representation of the different uptake mechanisms

by which TAT (translocation) and Antp (endocytosis) predominantly enter

the cell highlighting a potential role played by cell surface GAGs………………………………………

52

Figure 3-2. Heparin sepharose affinity chromatogram of methoxycoumarin labeled

HHBP, TAT, and Antp……………………………………………………………………………………

56

Figure 3-3. The hydrodynamic radii of clusters present in peptide-heparin solutions………………… 57

Figure 3-4. Fluorescence quenching of labeled peptide heparin solutions……………………………. 58

Figure 3-5. The quenching observed due to peptide-peptide interactions upon dilution of coumarin –

HHBP peptide with unlabeled HHBP peptide (total peptide concentration 10 µM) in the presence of a

fixed concentration of heparin (10 µM)………………………………………………………………….

59

Figure 3-6. Fluorescence of labeled peptide solutions in the presence of heparin…………………….. 60

Figure 3-7. Fluorescence of peptide solutions in the presence of excess heparin……………………… 61

Figure 3-8. Fluorescence change upon addition of excess heparin to preformed

heparin-HHBP aggregates………………………………………………………………………………...

62

Figure 3-9. Our proposed model that employs GAG clustering to explain differences

in CPP cell uptake mechanisms………………………………………………………………………….

65

Figure 4-1. Structures of thioester constructs…………………………………………………………… 78

Figure 4-2. Change in UV maxima (nm) during labeling………………………………………………. 79

Figure 4-3. Stability of construct 3M: 3M (10 µM) in HEPES buffer (10mM, pH 8) at 25 °C.............. 79

Figure 4-4. Stability of construct 2M: construct 2M (10 µM) in HEPES buffer (10 mM, pH 8)........... 80

Figure 4-5. UV spectra of isolated DAC labelled MBP(3M)…………………………………………… 81

Figure 4-6. ESI-MS timecourse of MBP labelling reaction with 3M from T=0 to T= 20hr.( 10 µM 3M

(HEPES buffer

xiii

(10 mM, pH 8, 20 hr, 22 °C……………………………………………………………………………… 82

Figure 4-7. UV spectra of isolated DAC labelled MBP(2M)…………………………………………… 83

Figure 4-8. A chart of the tabulated MBP labeling yield and selectivity

achieved using constructs 1M 2M and 3M………………………………………………………………

84

Figure 4-9. C-18 HPLC chromatogram of DAC labelled MBP trypsin digest......................................... 85

Figure 4-10. ESI-MS/MS spectrum of DAC labelled MBP tryptic peptide collected by HPLC............. 86

Figure 4-11. Structure of maltose bound MBP (PDB: 1MBD)………………………………………... 87

Figure 4-12. SDS-PAGE analysis of MBP labeling in crude bacterial cell extracts………………….. 88

Figure 4.13. Confocal microscopy, flow cytometry, and SDS-PAGE analysis of MBP

labeling in bacterial cells………………………………………………………………………………….

89

Figure 4-14. Confocal images based on z-stacking analysis of an E.coli cell following incubation

with 3M for 3hrs and PBS washing steps illustrating the cellular uptake of 3M.......................................

90

Figure 4-15. Flow cytometry histograms generated from fixed bacterial cells

following incubation with 3M. Live bacterial cells were incubated with 10uM 3M

for 3hr followed by PBS washing, formaldehyde crosslinking/fixation, and

washing (0.2% TWEEN-20, PBS) to remove unreacted 3M......................................................................

91

xiv

List of Schemes

Scheme 4-1. Synthesis of the thioester labeling conjugates………………………………………….. 77

Scheme 4-2. The labelling of MBP using ligand directed thioester conjugates..................................... 92

Scheme 4-3. Synthesis of glycosyl hydrazide thiols…………………………………………………... 92

Scheme 4-4. Synthesis of glycosyl thioester conjugates……………………………………………… 95

1

Chapter 1 Introduction

1.1 Carbohydrates in glycobiology

Carbohydrates are unique when compared to other classes of biomolecules such as nucleic acids

and proteins in their structural complexity. Polysaccharides are made up of monomeric

carbohydrate building blocks containing multiple functional groups allowing for a tremendous

degree of structural variability (1). In addition to the extensive degree of branching that can

result during the biosynthetic polymerization reactions, an additional layer of structural

complexity exists due to the stereocenter at the reducing terminus of each monomeric residue.

This results in the anomeric linkage of each sugar in the polysaccharide existing in either an α or

β configuration. The resulting polysaccharides can be further functionalized through the action of

numerous modification enzymes carrying out acetylation, sulfation, phosphorylation, and

oxidation reactions at various O and N positions of the sugar ring (1). How nature utilizes the

incredibly large structural diversity inherent to carbohydrates is poorly understood and remains a

major question in glycobiology. Carbohydrates serve numerous important structural and

regulatory roles in biology, existing alone as in the case of cellulose making up the cell wall of

plants, and starch which acts as a fuel storage or linked to various biomolecules in the form of

glycoconjugates found in the extracellular matrix. There are four main classes of naturally

occurring glycoconjugates: glycolipids, glycoproteins, GPI-anchors, and proteoglycans. These

glycoconjugates play important roles in cellular communication processes and in cell adhesion.

Glycolipids consist of a lipophillic 1,2-di-O-diacylglycerol or N-acylsphingosine moiety linked

to a hydrophilic carbohydrate moiety (1) (Fig 1-1). Examples of the two types of glycolipids,

glycoglycerolipids and glycosphingolipids, are shown in the figure below and are commonly

found in the plasma membranes of both plants and mammals. Glycosphingolipids are present in

eukaryotic membranes and are commonly found in neuronal cells in the form of galactosyl

cerebroside. More complex glycosphingolipids, called gangliosides, contain a common β-

lactosyl moiety within the ceramide unit (2).

2

Glycoproteins, in their soluble form, can be found in the cytosol, and in subcellular organelles.

They are also present in all cell membranes inserted into the lipid bilayer with the polysaccharide

portion extending extracellularly. Glycoproteins are linked to the polysaccharide chain through

N-glycosidic,O-glycosidic, or ethanolamine phosphate linkages (Fig1-2). N-linked glycoproteins

are linked to the polysaccharide via the side chain of an asparagine residue to the reducing

termini of N-acetylglucosamine (1). All N-linked glycoproteins have a conserved

pentasaccharide core structure linked to the protein via N-acetylglucosamine with the

sequence: Man-(α1,6)[Man-(α1,3)]Man-(β1,4)-GlcNAc-(β1,4)GlcNAc-Asn. The asparagine is

part of a consensus sequence consisting of Asn-Xaa-Ser/Thr where Xaa can be any amino acid

except proline (3).

Figure 1-1. The structures of a glycoglycerolipid (left) and glycosphingolipid (right).

O-linked glycoproteins, the most common of which are known as mucins, are linked through a

serine or threonine side chain to the reducing termini of a GalNAc sugar residue and these

glycans often make up the protective layer on the surface of epithelial cells (1, 2, 4). Linkages

between proteins and phosphorylated carbohydrates involving an ethanolamine linker unit occur

3

in glycosylphosphatidylinositols (GPI-anchors) which anchor the protein to the cell membrane

(2). Proteoglycans are a special class of glycoproteins containing a large protein bound

polysaccharide known as glycosaminoglycan.

Figure 1-2. The three structures of the protein-carbohydrate linkages found in glycoproteins. N-

linked (Top left), O-linked (Bottom left), GPI anchors (right).

1.2 Glycosaminoglycans

Glycosaminoglycans (GAGs) are a family of unbranched, acidic polysaccharides. Common

members of the GAG family include hyaluronan, chondroitin sulfate, dermatan sulfate, heparin,

and heparan sulfate (HS). The disaccharide repeating unit making up each of these GAGs is

composed of N-acetylglucosamine or N-acetylgalactosamine linked to a uronic acid residue with

either the D-glucuronic acid or L-iduronic acid configuration (Fig 1-3). Hyaluronan is composed

of N-acetylglucosamine and D-glucuronic acid. Chondroitin sulfate is composed of N-

acetylgalactosamine and D-glucuronic acid with variable sulfation at the O4 and O6 positions of

the galactosamine residue and the O2 position of the glucuronic acid residue. Like chondroitin

sulfate, dermatan sulfate also contains N-acetylgalactosamine however dermatan sulfate contains

only L-iduronic acid and has variable sulfation only at the O4 position of the N-

acetylgalactosamine residue in addition to potential sulfation at the O2 position of L-iduronic

acid. Heparan sulfate (HS) and heparin are the most highly sulfated GAGs containing either D-

glucuronic or L-iduronic acid linked to glucosamine with highly variable sulfation at the N2, O3,

4

and O6 of the glucosamine residue and the O2 position of the uronic acid residue. Heparin and

heparan sulfate differ in both their size and in the distribution of sulfates along the

polysaccharide. Heparin is highly sulfated throughout its length, in contrast to heparan sulfate

which has regions of high sulfation known as NS (N-sulfated) domains separated by regions of

low or no sulfation known as NA (N-acetylated) domains. Heparan sulfate is also larger and

more polydisperse than heparin with an average MW of ~ 30 kDa compared to heparin with an

average MW of ~ 15 kDa (5, 6).

Figure 1-3. The chemical structures of common members of the glycosaminoglycan family.

5

1.3 Heparin and Heparan sulfate

The polydispersity of heparin and heparan sulfate is largely conferred during GAG biosynthesis

in the Golgi. The biosynthesis of GAGs is initiated by the formation of a tetrasaccharide

composed of N-acetylglucosamine, galactose, and xylose with the sequence β-GlcNAc(1→3)-β-

Gal(1→3)-β-Gal(1→4)-β-Xyl-1, O-linked to a serine or threonine residue on the core protein

(Fig 1-4) (7). This is followed by the addition of either an N-acetylglucosamine which initiates

the biosynthesis of heparin or heparan sulfate or by the addition of N-acetylgalactosamine which

initiates the biosynthesis of chondroitin sulfate or dermatan sulfate (8). In the event of

HS/heparin biosynthesis, a transferase enzyme elongates the chain from the non-reducing

terminus with alternating glucuronic acid (GlcA) and N-acetylglucosamine GlcNAc residues

which are activated as UDP-sugar nucleotides (9). Chain elongation is followed by chain

modification starting with N-deacetylation and N-sulfation of the GlcNAc residues. It is the

apparently random and the incomplete nature of N-deacetylation by N-deacetylase enzymes that

is partially responsible for the heterogeneity of the polymer as this modification primes the

activity of the other modifying enzymes. The specificity of the N-deacetylase enzyme is also

responsible for the distribution of sulfates into discrete domains observed in the structure of HS

(5). The next step in the pathway involves the C-5 epimerization of D-glucuronic to L-iduronic

acid followed by O-sulfation of iduronic acid at C-2. Finally, 6-O-sulfotransferase enzymes

transfer a sulfate group to position 6 of the glucosamine residue which may be acetylated or

sulfated at the N position. At this stage in the biosynthetic pathway, 3-O-sulfotransferase

enzymes partially modify the polymer at various glucosamine residues at the C-3 position of

residues which already contain both N-2 and O-6 sulfation (GlcNS6S) (10). Following

biosynthesis, the heparin polymer is cleaved from its serglycin core protein by a glucuronidase

resulting in the production of GAG heparin which is primarily found in mast cells. HS remains

linked to the protein in the form of proteoglycan which can be secreted or reside in the plasma

membrane.

Heparin and heparan sulfate are involved in numerous biological processes. Heparin, is a

commonly administered clinical anticoagulant (11) and plays a direct role in the blood

coagulation cascade by facilitating the antithrombin III inhibition of the proteases thrombin and

factor Xa (12). Heparan sulfate, is involved in many cellular processes including the mediation

6

of tyrosine kinase receptor dimerization events, cell-cell communication through interactions

with cell surface adhesion molecules and regulation of leukocyte trafficking by modulating the

presentation of chemokines on the cell surface (13-15). Another important role of HS is to

sequester extracellular proteins such as hormones and growth factors to the cell surface where

cellular uptake via energy dependant endocytotic mechanisms can take place (16). The numerous

roles heparin and heparan sulfate play in these complex processes occur through binding

interactions with specific proteins.

Figure 1-4. A schematic representation outlining the biosynthesis of heparin and HS.

1.4 Lectins

One of the earliest observed examples of carbohydrate-mediated recognition was the mediation

of viral infection by sialic acid (17, 18). The initial step in viral infection involves adhesion of

the virus to the cell surface, which was shown to be mediated by the binding of viral

hemagglutinin to sialic acid on the cell surface. The discovery of the hepatic Gal/GalNAc

binding receptor triggered the earliest investigations into the potential of carbohydrates to act as

biological signals. It was found that in ceruloplasmin metabolism, serum glycoproteins with

exposed galactose residues following desialyation were quickly taken up by the liver and cleared

from the serum (19). This allowed for the discovery of hepatic carbohydrate binding receptors in

7

mammals and subsequently the discovery of Man-6-phosphate receptors for lysosomal enzymes,

and alveolar macrophage Man-receptors (17, 20).

Carbohydrate binding proteins, commonly known as lectins, have many biological roles. There

are thousands of examples of lectins in nature and they have been isolated from all orders of life.

Animal lectins were historically divided into two classes C-type (calcium dependent for

carbohydrate binding) and S-type (calcium independent) (21). The S-type lectins are found

ubiquitously in various tissues and organs and have a high degree of homology in contrast to C-

type lectins which differ structurally and are more organ and tissue specific. Examples of S-type

lectins include bovine spleen S-lectin which recognizes N-acetyllactosamine and human dimeric

S-lac lectin which binds to lactose. Examples of C-type lectins include E-selectin found in

endothelial cells, L-selectin in leukocytes, and P-selectin in platelets. These proteins play critical

roles in the immune response such as mediating the rolling of leukocytes along the blood vessel

wall through low affinity interactions with sialylated glycoproteins. There are now eight well

known classes of animal lectins, the first four include the calnexin family, M-type, L-type, and P-

type lectins which operate intracellularly playing important roles in facilitating the correct

folding and trafficking of proteins throughout the protein homoeostasis pathway. This is

followed by the four groups of lectins that operate extracellularly: C-type, R-type, Siglecs, and

galectins which are all found either associated with the plasma membrane or secreted (Table 1-1)

(22). Periplasmic binding proteins represent another class of carbohydrate binding proteins

exemplified by glucose and galactose binding proteins (GBP) and maltose binding protein

(MBP) isolated from bacteria, all of which play important roles in intracellular sugar transport

and metabolism. Lectins isolated from plants include the legume lectins such as Concavalin A

(ConA) which binds to glucose and mannose containing receptors such as rhodopsin and certain

immunoglobulins and the cereal lectins such as wheat germ agglutinin (WGA) which binds to N-

acetylglucosamine. The biological roles of plant derived lectins are not well understood.

8

Table 1-1. A summary of the different classes of animal lectins (22)

Lectin family Saccharide

ligands

Subcellular

location

Examples of functions

Calnexin Glc1Man9 ER Protein sorting in the

endoplasmic reticulum.

M-type lectins Man8 ER ER-associated degradation of

glycoproteins.

L-type lectins Varies ER, ERGIC, Golgi Protein sorting in the

endoplasmic reticulum.

P-type lectins Man 6-

phosphate, others

Secretory pathway Protein sorting post-Golgi,

glycoprotein trafficking, ER-

associated degradation of

glycoproteins, enzyme targeting.

C-type lectins Sialic acid/ others Cell membrane,

extracellular

Cell adhesion (selectins),

glycoprotein clearance, innate

immunity (collectins).

Galectins -Galactosides Cytoplasm,

extracellular

Glycan crosslinking in the

extracellular matrix.

I-type

lectins(siglecs)

Sialic acid Cell membrane Cell adhesion.

R-type lectins Varies Golgi, Cell

membrane

Enzyme targeting, glycoprotein

hormone turnover.

F-box lectins GlcNAc2 Cytoplasm Degradation of misfolded

glycoproteins.

Figolins GlcNAc, GalNAc Cell membrane,

extracellular

Innate immunity.

Chitinase-like

lectins

Chito-

oligosaccharides

Extracellular Collagen metabolism .

F-type lectins Fuc-terminating

oligosaccharides

Extracellular Innate immunity.

Intelectins Gal,

galactofuranose,

pentoses

Extracellular/cell

membrane

Innate immunity. Fertilization

and embryogenesis.

1.5 Lectin-carbohydrate interactions

The binding of carbohydrates by lectins generally involves an extensive H-bonding network

between the protein and the carbohydrate often mediated by water molecules (21). The burial of

carbohydrate hydrophobic surface area, release of water from the surfaces of the binding pair,

and the development of van der Waals contacts between the protein side chains and the

carbohydrate all contribute to the free energy of lectin-carbohydrate binding (21). Lectin-

carbohydrate binding is typically associated with negative enthalpy and entropy in addition to

negative changes in heat capacity. This was exemplified by isothermal titration calorimetry

(ITC) studies of concavalin A and a legume lectin from Dioclea grandiflora which were each

9

shown to bind to various mannosides containing one to three residues with negative changes in

heat capacity and negative overall changes in entropy (23). Unfavorable configurational entropy

was associated with binding in each case as expected, due to the loss of translational and

vibrational freedom which accompanies binding; however, the binding in each case was

observed to involve favorable changes in the entropy of solvation.

Lectins which commonly exist as oligomers in solution typically have shallow, solvent exposed

binding pockets resulting in relatively weak binding interactions being made to monovalent

carbohydrates. The energetics of these interactions are typically described by dissociation

constants in the µM-mM range (24). However, lectin binding to oligosaccharides can result in

significant enhancements in the apparent binding affinity when compared to the valence

corrected monovalent interaction due to a phenomenon known as “the glycoside cluster effect”

(24). This effect has in part been attributed to the enhancement in the apparent lectin-

carbohydrate binding affinity that can result from linked equilibrium between lectin-

carbohydrate complex formation and the aggregation of oligomeric lectins with oligosaccharides

stabilized by protein-protein interactions. Additionally, chelate binding interactions which

commonly play a role in multivalent interactions can contribute to the glycoside cluster effect

(24). Multivalent binding interactions between oligomeric lectins and the multiple binding sites

on the target oligosaccharide results in the simultaneous formation of several microscopic

complexes stabilized by a combination of intermolecular and intramolecular (chelate)

interactions. Chelate binding interactions between the multivalent lectin and multivalent

oligosaccharide occur with a larger more favorable free energy than the sum of the

corresponding intermolecular lectin-carbohydrate interaction free energies. This is due to the

lower entropic cost associated with decreasing the rotational and vibration degrees of freedom of

the multivalent ligand and receptor prior to binding (25). The increase in the apparent binding

affinity of multivalent lectin/oligosaccharide interactions in the absence of aggregation is

described by an avidity constant and is composed of a combination of the free energy

contributions from these intermolecular and chelate binding events in addition to an entropically

favorable component known as avidity entropy (25). Avidity entropy reflects the microscopic

degeneracy of each possible microscopic complex that can form in a multivalent interaction, is

always favorable and is independent of the magnitude of the intramolecular interaction.

10

1.6 Heparin/HS-Protein Interactions

The interactions between heparin/HS and biologically relevant peptides and proteins are largely

governed by electrostatic contacts. Due to the high negative charge density inherent to

heparin/HS, they exist in solution surrounded by a shell of counterions. Upon binding to cationic

regions on the cognate protein or peptide, the counterions surrounding heparin/HS and the

protein are displaced (Fig1-5). The entropically favorable release of counterions driving the

binding interaction is known as the polyelectrolyte effect (26). The extent to which this effect

drives the interaction is typically assessed through determining the relationship between the

dissociation constant for protein/peptide-HS/heparin binding and salt concentration. The binding

interaction between thrombin and heparin is largely driven by the polyelectrolyte effect with the

binding affinity shown to have a high salt dependence (27). Determination of the dissociation

constants between thrombin and heparin at varying concentrations of NaCl revealed that at least

five ionic contacts are made upon thrombin binding to heparin.

Figure 1-5. A schematic representation of the polyelectrolyte effect

11

This can be calculated using the following equation:

log KD = log KD,nonionic + m(1-f) log[Na+]

In this equation, KD represents the apparent dissociation constant containing contributions from

both ionic and non-ionic interactions, KD,nonionic represents the dissociation constant in the

absence of contributions from the polyelectrolyte effect, m represents the number of Na+ ions

released upon heparin protein binding and f represents the fraction of anionic charge on the

polyelectrolyte not charge neutralized by Na+ ions (27). A steeper slope indicates that a greater

number of electrostatic contacts are involved in the interaction while the y intercept gives the

dissociation constant for protein-heparin binding in the presence of 1M NaCl which describes the

contribution of H-bonding and hydrophobic interactions to protein-heparin binding affinity.

A consequence of the largely electrostatic nature of protein-heparin/HS interactions in-vivo

results in the majority of these interactions being non-specific. Such non-specific interactions

occur without a defined binding stoichiometry and do not require the tight ligand/receptor

complexation that accompanies specific interactions. Specific interactions occur with defined

stoichiometry and typically involve H-bonds and hydrophobic contacts which operate at shorter

distances and require a more geometric complementarity at the binding interface. Titrations of

thrombin with heparin oligosaccharides of increasing length indicate that the apparent binding

affinity increases with increasing length (27) (Table1-2). This binding behavior is consistent

with a non-specific binding model describing the interaction between thrombin and heparin.

Longer heparin oligosaccharides contain more non-specific overlapping binding sites

accompanying the longer negatively charged heparin chain. This has the effect of increasing the

number of possible single protein-single heparin chain microscopic complexes that can form

reflected in a higher affinity apparent dissociation constant.

12

Table 1-2. Titrations of thrombin with heparin oligosaccharides of increasing length (27).

Oligosaccharide

chain length

KD obs (µM)

4mer 71 ± 6

6mer 7.7±0.9

8mer 3.3±0.3

10mer 2.4±0.2

14mer 1.9±0.4

18mer 1.8±0.3

Although the majority of heparin/HS interactions with proteins are non-specific, there are a few

reported cases of binding specificity. One well known example of binding specificity occurs

between antithrombin III (ATIII) and heparin. Binding has been shown to occur between ATIII

and a specific pentasaccharide sequence in heparin containing a sulfate at the O-3 position of the

internal glucosamine residue (Fig 1-6). The absence of sulfation at this position results in a

significant decrease in binding affinity with the dissociation constant increasing over three orders

of magnitude from 300 nM to 500 µM (28). It has been proposed that the deletion of this critical

sulfate disrupts a co-operative network of hydrogen bonds between ATIII and heparin.

Figure 1-6. The structure of the specific pentasaccharide sequence on heparin recognized by AT III.

The critical sulfate at position O-3 is represented by R2. R = H, R

2 = H or SO3

-. RE = reducing

terminus, NRE = non reducing terminus

Heparin facilitates the ATIII inhibition of thrombin through a 50000 fold enhancement in the rate

of inhibition (29). The long, relatively flat, and narrow heparin chain binds to both ATIII and

thrombin acting as a molecular bridge, which reduces the ATIII-thrombin search time as the

proteins are brought out of solution in 3-dimensions to 1-dimensional heparin. This is known as

a reduction in a dimensionality (Fig1-7). Stopped flow analysis of the AT III-thrombin inhibition

13

reaction in the presence of heparin suggested the presence of a ternary complex consisting of

heparin bound to AT-IIII and thrombin (Fig1-8) (29).

Figure 1-7. An illustration of the reduction in dimensionality experienced by ATIII and thrombin

in the presence of heparin

Figure 1-8. A scheme representing the proposed mechanism of heparin rate enhanced ATIII

inhibition of thrombin due to ternary complex formation (29). AT.H and T.H represent protein-

heparin complexes with the respective protein, T.AT represents protein-protein complex without

bound heparin, T-AT-H represents the ternary complex and T-AT the irreversibly inhibited

thrombin protein linked to AT-III. The equilibrium constants calculated for each reversible step

are indicated.

Heparan sulfate has been proposed to play an important role in the binding of fibronectin, an

extracellular matrix protein, to vascular endothelial growth factor (VEGF) resulting in an

increase in cell proliferation and differentiation. Studies conducted using heparin as a model to

study HS revealed that heparin catalyzed a conformational change in fibronectin from a closed

14

(Fc) to an open extended (F0) conformation facilitating binding to VEGF (Fig1-9) (30).

Experimental evidence showed that heparin was no longer associated with fibronectin in the

open state and that catalysis occurred through initial high affinity binding by heparin to the

closed state of fibronectin. This was then followed by a conformational change to the open state

of fibronectin, which has a lower binding affinity for heparin resulting in heparin dissociation

(Fig1-9) (30). The origins of how heparin catalyzes the rate of conformational change remain

unclear. However, the observations that a non-stoichiometric amount of heparin is required for

catalysis and that the rate of conformational change from closed to open FN binding sites

increases with heparin concentration without changing the final open FN concentration support a

catalytic role played by heparin.

In summary, heparin and HS function in at least three different ways when interacting with

proteins. The first is through electrostatic contacts resulting in largely non specific interactions,

although instances of specific interactions between HS/heparin and certain proteins have been

reported. The second is through a reduction in dimensionality where heparin/HS reduce the

search time required for protein-protein interactions and the third function is as a potential

catalyst of protein conformational change

Figure 1-9. A schematic representation of the heparin catalyzed conformational change of

fibronectin (FN) (30). KC

A is the dissociation constant for heparin binding to the closed

conformation of FN, kc is the rate constant for conformation change from closed to open FN

conformation, K0A is the dissociation constant for heparin binding to the open conformation of FN.

15

1.7 Techniques to study HS/heparin interactions

There are a variety of techniques used to study the interactions between Heparin/HS and various

peptides and proteins. Surface Plasmon resonance (SPR) is becoming increasingly popular for

the quantitative analysis of peptide/heparin interactions yielding kinetic rate constants in addition

to equilibrium binding constants (31). This technique requires the immobilization of one binding

partner to a sensor chip while the other binding partner is passed over the chip. A binding event

results in a change in the angle of reflected light from the host or guest tethered sensor chip with

the magnitude of this change proportional to the fraction of host or guest bound. Although an

effective method for determining relative binding affinities between peptides and GAGs, SPR

has the drawback of measuring the interaction under non-native conditions in addition to

requiring a significant amount of time for analyte immobilization and chip optimization (5).

The use of affinity chromatography is a popular qualitative approach used to study

peptide/protein-heparin interactions where again, one of the binding partners, usually heparin is

immobilized to a solid support and the peptide or protein is passed through the column. The

peptide or protein is eluted with a salt gradient and the retention time is proportional to the

relative binding affinity of the peptide or protein for heparin. This is a rapid technique and an

efficient method to assess relative binding affinity for heparin, however it has the major

limitation of only assessing the electrostatic component to binding (5).

Isothermal Titration Calorimetry (ITC) can provide information on the thermodynamic binding

parameters behind heparin/HS-protein/peptide interactions including the ∆H, KD, and the

stoichiometry of binding also allowing for the calculation of ∆S (32). This technique measures

the heat released or absorbed that accompanies the binding interaction between the two binding

partners. ITC has the limitations of requiring high (mg) amounts of analyte in order to reliably

detect changes in heat flow which can lead to precipitation and aggregation in the case of

peptide/protein interactions with heparin which can occur at higher protein concentrations.

Another limitation of the technique is that it can only monitor dissociation constants in the range

of 10-4

to 10-8

(5).

16

NMR can provide very useful structural and conformational information regarding

protein/peptide-heparin/HS interactions however it is also relatively insensitive requiring large

amounts of material which again can result in solubility problems. This technique is very limited

for determining low dissociation constants which are common to heparin binding interactions in

addition to being further complicated by the ability of heparin/HS to form aggregates with

several proteins and peptides at higher concentrations. This can lead to significant peak

broadening from T2 spin relaxation effects (33).

An extremely sensitive approach to studying heparin/HS interactions involves fluorescence

spectroscopy, where the binding event is coupled to a change in fluorescence response. This can

be achieved using intrinsic fluorescence in the analysis of protein interactions due fortuitous

tyrosine and tryptophan residues within the protein (34). Often these studies can reveal

information regarding the protein conformation accompanying HS/heparin binding. Extrinsic

fluorescence may also be used to monitor binding and obtain equilibrium binding data through

the tethering of fluorescent dyes to the peptide/protein (35, 36). The advantage of this approach

is that it requires very low concentrations of material for the analysis but the tethered

fluorophores can potentially enhance or disrupt the native interaction.

1.8 The development of probes for HS/heparin

The importance of HS and heparin in the regulation of numerous biological processes in

addition to the role of heparin as a clinical anticoagulant, have inspired the development of

sensitive and selective probes for heparin/HS. Heparin is often administered during

cardiopulmonary bypass surgery to prevent thrombosis (11) and must be carefully monitored to

prevent a potentially fatal overdose. The analysis is complicated by the complex and

heterogeneous structure of heparin making quantification challenging. Heparin levels are

typically monitored indirectly by determination of the activated clotting time (ACT) of whole

blood through measurement of the mass deposition or viscokinetic changes (5, 37). The

development of direct heparin probes with increased sensitivity and potentially rapid and direct

quantification of serum heparin levels could contribute to the accuracy and effectiveness of the

ACT method.

17

Heparan sulfate is associated with all amyloid deposits present in numerous protein folding

diseases such as Alzheimer’s, and prion disease (38). HS has also been observed to be

differentially regulated on the surface of numerous cancer and tumor cells (39). Recently, the

differential regulation of HS was observed in amyloid burdened tissue and ovarian tumor cells in

mice using radiolabelled antibodies that bind selectively to HS (40). These antibodies present a

powerful imaging strategy for the identification of such pathological conditions. The

development of HS probes capable of reversibly binding to HS/heparin under biologically

relevant conditions could find a number of useful applications including the imaging of HS on

the surface of live cells with spatial and temporal resolution and the ability to quantify the

concentrations of HS/heparin on cells or in tissue cultures. Such probes would require high

selectivity and sensitivity for heparin/HS in the presence of numerous other biomolecules while

being soluble and resistant to aggregation under aqueous conditions. For quantitative

applications, these probes would also require ratiometric readout capabilities. Ratiometric

probes are required for accurate heparin quantification because the signal ratio is independent of

the probe concentration. The ratio of fluorescence at two wavelengths can then be correlated to a

concentration of heparin which is useful for measuring rapid fluctuations in heparin

concentration and in scenarios where it may be difficult to control the probe concentration

Currently, there are examples of heparin probes that have been designed in attempts to meet

these criteria (35, 36, 41, 42). These probes employ electrostatic interactions to facilitate high

affinity binding to heparin and an absorbance or fluorescence read-out to confer high sensitivity

for potential detection of HS/heparin in a biological setting. The probes are equipped with

reporter systems that either employ chromophore displacement or tether an environmentally

sensitive chromophore to the probe to provide a read out of probe-heparin interactions. Each

type of reporter system has limitations. Direct tethering of a dye can enhance or inhibit the

interactions of heparin with the probe and potentially induce probe aggregation. Chromophore

displacement readouts can suffer from a low dynamic change in the signal response if the

chromophore to be displaced upon probe-heparin binding interacts with the probe with too low

an affinity. However if the chromophore binds too tightly to the probe, the affinity of the probe

for heparin will be reduced. These opposing factors result in the need to optimize chromophore-

probe binding affinity prior to studies with heparin.

18

An example of a heparin probe employing a chromophore displacement readout is the

calorimetric heparin sensor developed by Ansyln et al. (42). The synthetic receptor (Fig1-10)

incorporates ammonium functional groups and phenyl dimethylamino boronic acid moieties

well known for their ability to rapidly form boronic esters with the diols of carbohydrates (42).

Heparin binding displaces a receptor bound pyrocatechol violet chromophore resulting in a UV

absorbance change. Titrations with heparin revealed this construct to have mM affinity for

heparin and almost no ability to recognize hyaluronan. Despite respectable selectivity, this probe

suffers from relatively low binding affinity and requires a 1:1 buffer/methanol co-solvent for

solubility, far from physiological conditions. This limits the application of this probe for

detection of heparin in cell or tissue culture.

Due to the primarily electrostatic interaction of heparin probes, the affinity of the probe for

heparin is highly sensitive to the ionic strength of the solution. This complicates the application

of heparin probes for heparin detection under physiologically relevant ionic strength where the

heparin binding affinity can drastically decrease. A peptide based approach to probing heparin

under physiological ionic strength was reported using heparin binding peptides tethered to a

chloride sensitive quinolinium ion (Fig1-11) (36). The synthesis of the probe also allowed for

the positioning of the quinolinium reporter at any position in a potential heparin binding

sequence.

19

Figure 1-10. A synthetic receptor approach to heparin probe design.

Figure 1-11. The chemical structure of the quinolinium tethered heparin probe. R = the α-carbon

of an amino acid.

The quinolinium probe was shown to selectively detect heparin over chondroitin sulfate at low

µM heparin concentrations relevant to the serum concentration range of heparin administered

clinically (1.2 to 7.3 µM). However, the presence of numerous proteins with nanomolar affinity

and the lack of a ratiometric fluorescent readout of this probe limits its application for the

quantification of heparin or HS in biological media.

20

A ratiometric peptide based probe for heparin was developed using diethylaminocoumarin

(DAC) excimer fluorescence (35). The probe was designed based on a dimeric coiled-coil motif

(Fig 1-12) with each peptide labeled with 7-diethylaminocoumarin-3-carboxylic acid at the N-

terminus. The peptides were designed to form a coiled-coil only in the presence of heparin by

incorporating lysine residues at the positions flanking the hydrophobic coil-coil interface.

Coiled-coil formation is accompanied by a red shift in fluorescence from 477 nm to 560 nm due

to the formation of DAC excimers. The ratio of the fluorescence at 560 nm and 477 nm is

proportional to the concentration of heparin titrated into solution under conditions of excess

probe to heparin. The coiled-coil probe was observed to bind to heparin with high affinity

having a dissociation constant in the low µM range which decreased further to low nM upon

preforming a dimeric peptide with a disulfide linkage. Selective recognition of heparin was also

observed over other polyelectrolytes (Fig1-13). A major limitation of this probe is that under

high heparin:probe ratios more complicated binding behavior was observed, with the ratiometric

fluorescence no longer being proportional to heparin concentration. Due to the high

concentration of heparan at the surface of many cell types, the complicated binding profile of this

probe limited its utility in biological samples.

.

Figure 1-12. The coiled coil motif applied as a ratiometric probe for heparin. The cationic heparin

binding site containing lysine residues at the e and g positions are indicated by arrows.

21

Figure 1-13. The selectivity of the probe for heparin over other polyelectrolytes. The probe (2 µM)

was titrated with heparin (■), DNA (▼), chondroitin sulfate A (▲), or polyglutamic acid (●) in

phosphate buffered saline (pH 7.0). (Reprinted with permission from Ref (35))

The design of most heparin probes are done with the intention of probing HS, using heparin as a

proof of concept due to its less complex structure, availability and lower cost. Examples of HS

selective probes in the literature are scarce, however, the development of a self assembling

fluorescence receptor system that can distinguish HS from heparin and other GAGs using pattern

based recognition has been reported (41). The GAG is recognized using a series of self

assembling cyclodextrin-quinolinium tethered lithicholic acid receptors functionalized with

amino or guanidine groups on the cyclodextrin (CD) motif. The cations interact with the GAG

of interest and the binding event is coupled to an increase in quinolinium fluorescence. The

series of probes differ in their relative fluorescence response to GAG binding due to structural

differences in the functionality of the CD and in the tethering of the quinolinium fluorophore to

the lithicholic acid (Fig1-14). This results in the generation of a fluorescence intensity profile

unique to the structural nature of the GAG. Although this approach is very useful to determine

the purity of heparin or heparan samples, such an approach is limited due to the supramolecular

nature of the probe and potential competing binding interactions.

22

Figure 1-14. Self-assembling fluorescent receptors for GAGs. The receptors consist of a

polycationic cyclodextrin host (1a-e) and a fluorescent reporter tethered to a lithocholic acid guest

(2a-c).

1.9 Purpose of study

Establishing the relationship of HS and protein/peptide structure to the affinity and selectivity of

the binding interaction is critical for the development of probes for HS and to better understand

the nature of HS interactions. These probes would provide important tools to understand how

the cell regulates key biological processes through modulation of the highly variable structure of

HS. Our initial investigations into the role of the spatial display of charge in cationic peptide-

heparin binding were conducted to determine the preferred binding modes of heparin/HS to

peptides in addition to the optimal spatial display of positive charges for high affinity binding.

The results and impact of our findings on future HS probe design are presented in Chapter 2 and

5. In Chapter 3, investigations into the role of aggregation in heparin interactions with cationic

cell penetrating peptides (CPPs) is explored. Experimental data allowed us to propose a model

explaining why CPPs that bind heparin with similar affinity enter the cell through different

mechanisms and with a different dependence on cell surface HS. In Chapter 4, the development

of a strategy to selectively label carbohydrate binding proteins using an affinity based labeling

technique employing thioester acylation chemistry is described. The selective labeling of

maltose binding protein, as a proof of concept, is presented and its applicability to the design of

HS protein biosensors discussed. Concluding remarks and the impact of this work on the field of

glycobiology will be addressed in Chapter 5.

23

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39. Nackaerts, K., Verbeken, E., Deneffe, G., Vanderschueren, B., Demedts, M., and David,

G. (1997) Heparan sulfate proteoglycan expression in human lung-cancer cells,

International Journal of Cancer 74, 335-345.

40. Smits, N. C., Kurup, S., Rops, A. L., ten Dam, G. B., Massuger, L. F., Hafmans, T.,

Turnbull, J. E., Spillmann, D., Li, J.-p., Kennel, S. J., Wall, J. S., Shworak, N. W.,

Dekhuijzen, P. N. R., van der Vlag, J., and van Kuppevelt, T. H. (2010) The Heparan

Sulfate Motif (GlcNS6S-IdoA2S)3, Common in Heparin, Has a Strict Topography and Is

Involved in Cell Behavior and Disease, Journal of Biological Chemistry 285, 41143-

41151.

41. Jagt, R. B. C., Gómez-Biagi, R. F., and Nitz, M. (2009) Pattern-Based Recognition of

Heparin Contaminants by an Array of Self-Assembling Fluorescent Receptors,

Angewandte Chemie International Edition 48, 1995-1997.

42. Zhong, Z., and Anslyn, E. V. (2002) A Colorimetric Sensing Ensemble for Heparin,

Journal of the American Chemical Society 124, 9014-9015.

27

Chapter 2 The effects of the spatial display of positive charge on peptide

heparin binding affinity0F

1

2.1 Introduction

In solution, heparin/HS exists primarily as an extended helical structure with conserved

glycosidic bond angles, yet heparin possesses an inherent degree of conformational flexibility (1-

3). This flexibility is due to the similar energy of and rapid interconversion between the chair and

skew boat conformations of the iduronic acid residues which are abundant in heparin and the S-

domains of heparan sulphate (4) (Fig 2-1). The conformational flexibility inherent to the heparin

chain allows for diverse spatial displays of negative charge which facilitates interactions with

proteins. Insight into the possible conformations assumed by heparin/HS upon protein and

peptide binding has been gained through studies of heparin alone using NMR and x-ray

crystallography (1-3). Information regarding the conformation taken up by heparin upon

complexation to proteins has also been obtained through x-ray crystallography of protein-heparin

complexes using heparin derived oligosaccharides. This was done to study the structure of

heparin complexes with the protease inhibitor protein ATIII (5).

Investigations into the structural requirements of the protein for high affinity binding to heparin

were first made by Cardin and Weintraub (6) who proposed the existence of consensus

sequences. Alignment of heparin binding sequences from apolipoprotein B, apolipoprotein E,

vitronectin, and platelet factor 4 revealed the consensus sequences XBBXBX and XBBBXXBX

where X is a hydropathic residue and B is a basic residue. The modeling of these consensus

sequences as α helices showed that positive charge was largely displayed on one face of the helix

suggesting that the spatial display of positive charge is potentially an important determining

factor for high affinity binding to heparin (6).

1 Reproduced with permission from Rullo, A., Nitz, M (2009). Importance of the Spatial Display of

Charged Residues in Heparin-Peptide Interactions. Biopolymers, 93, 290-298. The introduction of the

main text has been modified and figures have been added from the supplementary information. The text

from the materials and method sections in addition to the results and discussion section remain

unchanged.

28

Figure 2-1. The preferred conformations of glucosamine and uronic acid residues within the

heparin/HS chain in aqueous solution. RE refers to the reducing terminus of the sugar and NRE

refers to the non-reducing terminus of the sugar.

Engineered peptides incorporating these two heparin binding consensus sequences XBBXBX

and XBBBXXBX showed increases in α-helicity in the presence of heparin as determined by

circular dichroism spectroscopy indicating that heparin stabilizes the helical form of these

peptides upon binding (7,8). Molecular modeling of heparin binding sequences from apoE and

platelet factor 4 as α-helices revealed that a distance of 20 Å was present between clusters of

positive charge however, in these examples, charge density was found on opposite sides of the α-

helix. This led the authors to propose that heparin preferred to coil around an α-helical peptide

target or binding interface on a protein in order to maximize contacts with these basic regions

(9). It was also noted that such coiling could induce a conformational change in the heparin

binding protein.

Attempts to unravel predictive factors for favourable protein heparin binding have implicated the

importance of peptide primary sequence, and how positive charge is presented spatially.

However, essential to understanding the heparin-protein interaction is knowledge of the

relationship between the display of charge, binding affinity, and conformation before and after

the binding event. To address the structural requirements that govern α-helical peptide/protein-

29

heparin interactions in biological systems, we designed seven peptides that would differ only in

the spatial distribution of positive charges upon taking up an α-helical conformation. The binding

event could be monitored by changes in secondary structure experienced by the peptides upon

heparin binding using circular dichroism spectroscopy (CD).

To determine the structural requirements for high affinity binding between heparin and its

cognate protein or peptide, a sensitive method would be required to measure heparin-peptide

binding affinity under physiologically relevant ionic strength and at low peptide concentrations.

Towards this end, a quinolinium ion, was chosen to probe the binding event through covalent

tethering to the N-terminus of each of the peptides. The quinolinium ion has been employed

previously to probe peptide-heparin interactions through covalent tethering to a heparin binding

peptide (10). It was shown that the quinolinium tethered peptide was collisionally quenched by

chloride ions in solution, with fluorescence intensity decreasing with increasing concentrations

of NaCl. After the addition of heparin, the quinolinium dye experiences a relief of quenching

and a “turn on” increase in fluorescence following peptide-heparin complexation (10). This is

due to the fact that heparin binding decreases the local concentration of chloride ions

surrounding the quinolinium due to negative charge repulsion, increasing the quantum yield of

the quinolinium ion (Fig 2-2). The increase in fluorescence observed upon titrating heparin into

a solution of fixed peptide concentration, proportional to the fraction of peptide bound to

heparin, allowed for the calculation of peptide-heparin dissociation constants (10).

30

Figure 2-2. The proposed mechanism of quinolinium probing of peptide-heparin binding

interactions.

2.2 Results

The synthesized peptides designed differ in their spatial arrangement of positive charges when

folded into a 28 residue α-helix (Fig2-3, Table 2-1). Lysine residues were substituted at various

positions from a to g in the helical heptad repeat with each peptide containing 8 lysine residues

separated by alanine residues at the remaining positions. The high helical propensity of alanine

and lysine residues promote an α-helical conformation in the peptide (8, 12, 13). The binding

interaction is expected to be predominantly electrostatic in nature as has been observed in

numerous glycosaminoglycan protein interactions (4, 12, 14). At the N-terminus of each peptide,

two glycine residues were installed to separate the attached fluorescent quinolinium reporter

from the α-helix. Figure 2-3 shows the sequences of peptides modeled in an α-helical

conformation. Peptide 1 has lysines at the a and d positions of the heptad repeat forming a

continuous face of positive charge along the α-helix. Peptide 2 has lysine residues at positions a

and d of the first two heptad repeats and at c and f positions of the remaining two heptads,

positioning positive charge on opposite faces of the α-helix. Peptide 3 was constructed with

lysines gradually spiralling around the α-helix. Peptides 4 and 5 were constructed with lysines

broadly distributed along the helix (Table 2-1).

31

In order to assess the α-helical content of the peptides, circular dichroism spectroscopy (CD) was

used to characterize the peptide conformations in the presence and absence of saturating heparin.

Although this measurement does not provide a high resolution structure of the amino acid side

chains it does provide a useful estimation of the spatial position of the positively charged

residues when the peptides are highly helical. Helicity of the peptides was evaluated by

measuring the mean residue ellipticity (MRE) value θ222 deg cm2dmol

-1 of each peptide. The

maximum theoretical α-helicity for a 30 amino acid peptide is 36,670 deg cm2dmol

-1(15).

Figure 2-3. Helical wheel and ribbon-ball models of the five helical heparin binding peptides

synthesized presented as α-helices. The models are provided as a guide to the position of the side

chains in each peptide if it were to adopt an ideal α-helix.

32

It was observed that peptide 1 was the least helical before the addition of heparin with an MRE

corresponding to 49% helicity (Fig2-5(A), Table 2-2). This is not surprising as peptide 1 contains

three i, i+4 electrostatically repulsive interactions which have been shown to be destabilizing to

an α-helix (16, 17). Peptides 3 and 5 have an intermediate amount of helical content prior to

heparin binding, corresponding to approximately 70% helicity. Both peptides 3 and 5, lack the

repulsive i, i+4 interactions explaining the higher degree of a-helicity compared to peptide 1.

Peptide 2 and 4 are greater than 90% helical as has been observed in literature for alanine rich

peptides containing non-terminal charged amino acid side chains (18). Interestingly, peptide 2

with i, i+4 repulsions is more helical than peptides 3 and 5 which lack these repulsive effects.

This is likely the result of a balance between peptide backbone desolvation by lysine residues

versus their repulsive effects (19). Upon binding heparin, peptide 1 experienced a significant

increase in α-helicity becoming 95% helical (Fig2-5 (B)). The remaining peptides exhibited no

change or very little change in helicity upon heparin binding (Table 2-2).

To determine the effect of lysine distribution on peptide affinity for heparin, each peptide was

titrated with heparin and the fluorescence emission of the quinolinium reporter was observed at

430 nm. The binding energy behind the interaction is highly dependent on the ionic strength of

the solution and conditions similar to those found in physiological environments were chosen (50

mM phosphate buffer, 150 mM NaCl). Because of the polyelectrolyte driving force behind the

peptide interaction, lower ionic strength buffers will significantly strengthen the peptide heparin

interaction. The fluorescence data were fit to a 1:1 binding isotherm and dissociation constants

extracted. The titration experiments were performed at fixed peptide concentrations significantly

below the dissociation constants (KD) for the complexes. The concentration of peptide binding

sites in the heparin solution was estimated from an equivalence point titration at high

concentrations of peptide 1 (Fig 2-4). This titration led to an estimated binding site molecular

weight of 4000 Da approximately corresponding to a dodecasaccharide containing 18 sulfate

groups. The concentration of peptide binding sites in the heparin solution is used for the

calculation of the dissociation constants due to the structural heterogeneity of heparin with

respect to sulfation, sequence, and length. The quantification of binding sites is based on the

assumption that heparin consists of multiple, equivalent, overlapping non-interacting binding

sites for the basic peptides. The equivalence point titration, defines the concentration of peptide

binding sites under conditions where all the heparin chains in solution are saturated with peptide

33

[peptide] >> KD. Although useful, this determination results in a systematic error in the

concentration of peptide binding sites calculated, as under the high concentrations of peptide

required in an equivalence point titration, weak binding sites on the heparin chains are occupied.

Figure 2-4. Fluorescence Titration of peptide 1 with heparin, 50 µM peptide 1 titrated with heparin

50 mM phosphate buffer 150 mM NaCl pH 7. A blank buffer trace was subtracted from each

fluorescence spectrum. Excitation at 336 nm. ■ 0 µM, ● 10 µM, ▲ 20 µM, ▼ 30 µM, ♦ 50 µM

peptide binding sites on heparin.

34

Table 2-1. Sequences of Synthesized Heparin Binding Peptides

Peptide Description

Sequence Quin-GG-(abcdefg)4a

1 Charges on one

face

XGGKAAKAAAKAAKAAAKAAKAAAKAAKAAA

2 Charges on two

opposite faces

XGGKAAKAAAKAAKAAAAAKAAKAAAKAAKA

3 Charges align as

a gradual spiral

down the helix

XGGKAAKAAAAKAAKAAAAKAAKAAAAKAAK

4 Charges broadly

distributed along

helix

XGGKAAKAAAAAKAAKAAKAAKAAKAAKAAA

5 Charges broadly

distributed along

helix

XGGKAAAAKAKAAAAKAKAAAAKAKAAAAKA

1 Gap 6 alanine gap in

peptide interior

charges along

one face

XGGAAAKAAKKAAKAAAAAAKAAKKAAKAAA

1R Charges along

one face

XGGRAARAAARAARAAARAARAAARAARAAA

a X refers to the quinolinium fluorophore reporter.

The overestimation of the binding site concentration when applied to titrations at low peptide

concentrations, significantly below KD, results in an apparent KD that is larger (weaker affinity)

than the microscopic KD. To determine dissociation constants that best represent a 1:1 peptide-

heparin binding event, it is necessary to carry out KD determination at low peptide

concentrations. This is due to the fact that the apparent dissociation constants calculated will

increase when titrations are carried out at high peptide binding densities (20). At low

concentrations of peptide ([peptide] << KD), the binding event takes place under conditions of

excess heparin and as a result, the majority of binding sites on heparin remain unbound upon

completion of the titration. The quinolinium fluorescence probe employed to report on the

peptide heparin binding interaction, effectively operates at very low peptide concentrations. This

allows the titrations to be carried out at lower concentrations than those determinations which

use tryptophan/tyrosine fluorescence (9, 21). Although approximation of the number of peptide

binding sites using an equivalence point titration is inherently flawed for determining intrinsic

microscopic dissociation constants, it allows for comparison of apparent dissociation constants of

a 1:1 peptide-heparin binding event experienced by different heparin binding peptides.

35

A significant difference in binding affinity was observed among the peptides which differ in the

spatial distribution of positive charge. The strongest lysine based peptide, peptide 1, with lysines

at the a and d position had an apparent KD of 640 nM for heparin (Fig2-6 (A)). The binding

affinity for heparin significantly dropped upon decreasing the positive charge density along one

face of the α-helix. Peptides 2 and 3 each had a dissociation constant of 25 µM. Peptide 2 had

lysines in two patches on opposite faces and opposite ends of the α-helix while peptide 3

displayed lysines in a gradual spiral-like distribution along the helix. The two weakest heparin

binding peptides were peptides 4 and 5 with dissociation constants of 49 and 44 µM (Fig 2-

6(B)). Peptides 4 and 5 had lysines more diffusely spaced along the α-helix. The highest affinity

binding for heparin by peptide 1 suggests that heparin can make the most contacts with a peptide

displaying high charge density localized in a linear fashion.

Two more synthetic peptides 1Gap and 1R were produced based on peptide 1 to further evaluate

high affinity heparin binding peptides. Peptide 1Gap had lysines positioned in two clusters along

one face of the α-helix separated by a six alanine sequence. This peptide was constructed to

optimize contacts with sulfate groups based on the spacing of sulfate residues present in the

solution structure of heparin (Fig2-7) (2). CD analysis was carried out before and after the

addition of heparin to determine the helical content. The 1Gap peptide was 61% helical before

binding heparin (Fig2-5(A)). The reduced helicity in this peptide is likely due to i, i+4

repulsions. The 1Gap peptide experienced an increase in helicity after binding heparin becoming

95% helical (Fig2-5 (B)). Determination of the binding affinity of this peptide for heparin gave

rise to a calculated KD of 3.4 µM which was surprisingly higher (lower affinity) than peptide 1

(Fig2-6A). To investigate the possible origins of this unexpected decrease in heparin binding

affinity by the 1Gap peptide, titrations were conducted using a 14mer oligosaccharide of heparin

corresponding to the approximate molecular weight of the peptide binding site calculated by an

equivalence point titration.

36

Figure 2-5. (A) CD spectra of seven heparin binding peptides (75 µM) without heparin (50 mM

phosphate buffer 150 mM NaCl pH 7). ■ Peptide 1, ▼ Peptide 2, ♦ Peptide 3, ► Peptide 4, ◄

Peptide 5, ● 1R, ▲ 1Gap; (B) CD spectra of the three highest affinity heparin binding peptides (75

µM) with heparin (500 µM), 50 mM phosphate buffer 150 mM NaCl pH 7. ■ peptide 1, ● 1R, ▲

1Gap. The CD spectra of the remaining peptides did not change significantly upon heparin

addition.

37

Table 2-2. Experimentally Determined Dissociation Constants and Structural Binding Characteristics of

Seven Synthetic Peptides

Peptide KD (µM) θ222 w/o

Heparin

(% helicity)a

θ222 with

Heparin

(% helicity)a

∆θ222

1 0.64 ± 0.07 17,770 (49%) 34,910 (95%) 17,140

2 24.9 ± 0.6 35,080 (95%) 34,720 (95%) -360.0

3 25 ± 1 27,830 (76%) 28,200 (76%) 360.0

4 49 ± 4 33,230 (91%) 36,620 (100%) 3,380

5 44 ± 4 24,670 (67%) 24,740 (67%) 70

1Gap 3.4 ± 0.3 22,350 (61%) 36,050 (98%) 13,700

1R 0.0034 ±

0.0006

36,840 (~100%) 43,960 (~100%) 7,120

a The maximum theoretical a-helicity for a 30 residue peptide is 36,670 deg cm-2 dmol-1

As observed previously (22) the binding affinity dropped drastically for both peptides 1 and

1Gap when binding the heparin oligosaccharide compared to full length heparin. Interestingly,

the calculated dissociation constants for oligosaccharide binding by peptide 1 and 1Gap are

within experimental error of each other with an apparent KD of ~ 80 µM (Fig2-8). Peptide 1R

has arginines substituted for lysines positioned at the a and d positions along one face of the α-

helix. This peptide is expected to bind heparin with higher affinity due to the 2.5 times more

favorable enthalpy associated with arginine-heparin interactions versus that of lysine (23). CD

analysis of the 1R peptide before binding heparin revealed that 1R has the highest degree of α-

helicity of all the peptides studied. Arginine is known to have a higher helical propensity than

lysine. Arginine residues have also been shown in literature to increase the degree of α-helicity

by shielding the backbone hydrogen bonds from water through favorable interactions between

the charged side chain and the carbonyl oxygen at i, i+4 (18). Upon binding heparin, the 1R

peptide became more helical with an MRE of θ222 = 43, 960 deg cm2

dmol-1

(Fig2-5(B)). In

addition to the high degree of helicity, 1R bound heparin with the highest affinity of all the

peptides with KD 3 nM (Fig2-9).

38

Figure 2-6. Binding isotherms of synthesized peptides to heparin. (A) ■ peptide 1 (100 nM), ● 1Gap

(100 nM); (B) ■ Peptide 2 (1 µM), ● Peptide 3 (1 µM), ▲ Peptide 4 (1 µM),▼ Peptide 5 (1 µM).

All titrations carried out in 50 mM phosphate buffer, 150 mM NaCl, pH 7. Each point of the

titration represents individually prepared solutions to avoid photobleaching effects.

39

Figure 2-7. Schematic representation of the proposed 1Gap peptide interaction with heparin.

Engineering of peptide sequence based on the alignment of lysine groups on the peptide with

sulphate groups on a solution structure of heparin solved previously. Sulfate esters and lysine -

amino groups are emphasized with spheres.

40

Figure 2-8. Peptide 1 and 1Gap titrations with heparin 14 mer oligosaccharide (50 mM phosphate

buffer pH 7, 150 mM NaCl. Peptide 1 ♦, Peptide 1Gap ■

Figure 2-9. Nonlinear least squares analysis of binding of peptide 1R (25 nM) with heparin (50 mM

phosphate buffer, 150 mM NaCl, pH 7). Each point on the titration represents individually

prepared solutions to avoid photobleaching effects.

4000

9000

14000

19000

24000

29000

0 50 100 150

Emis

sio

n 4

30

nm

14mer (µM)

41

2.3 Discussion and Conclusions

The heparin binding affinity possessed by the synthesized helical peptides was shown to be

highly dependent on the spatial orientation of the positively charged lysine amino acids. Of the

lysine based peptides, peptide 1, with charge distributed along one face of an α-helix, bound

heparin with the highest affinity. NMR studies have shown heparin in solution to exist

predominantly in an extended helical conformation (2). Circular dichroism supports the α-

helical conformation of peptide 1 upon interacting with heparin clearly indicating an increase in

helicity upon heparin binding. This increase in helicity most likely results from relief of i, i+4

electrostatic repulsions in the α-helix upon interaction with heparin. In the case of the remaining

lysine-based peptides, binding affinity drops significantly as positive charge density is

distributed away from a linear arrangement along one face of the α-helix. Positive charge

distributed on two faces in the case of peptide 2, along a spiral in the case of peptide 3, and

broadly distributed in the cases of peptide 4 and 5 results in a significant decrease in heparin

binding affinity. These lower affinity peptides are highly helical and the degree of α-helicity

associated does not change upon binding to heparin. These observations show that the peptides

do not adopt an alternate conformation to make a greater number of ionic contacts with heparin.

In addition, these observations suggest that alternative heparin conformations that may wrap

around the α-helix to facilitate a greater number of ionic contacts with the peptide are

energetically unfavorable. The observed decrease in binding affinity with peptides 2-5 is likely

the result of heparin remaining extended and making fewer contacts while the peptides maintain

their favorable α-helical conformation. The 1R peptide with the lysines of peptide 1 replaced

with arginine residues bound heparin with a significantly higher affinity than its lysine

counterpart. This was expected, as Lindhardt et al. have demonstrated a more favourable

enthalpy for the interaction of arginine versus lysine with heparin and have attributed this

observation to the softer electrostatic interactions between the arginine guanidino and heparin

sulfate groups as well as to stronger hydrogen bonds between these functional groups (23). In

addition peptide 1R was significantly more helical than peptide 1 prior to binding heparin and

experienced a smaller conformational change upon binding heparin than peptide 1, thus the

preorganized structure of peptide 1R likely contributes to its higher heparin affinity.

42

Surprisingly, the 1Gap peptide displayed a fourfold decrease in heparin binding affinity

compared with peptide 1. In addition to the potentially optimized ionic contacts with heparin,

peptide 1Gap is also more helical prior to binding heparin than peptide 1 and thus should have a

reduced conformational entropic cost upon heparin binding. The binding experiments using a

14mer of heparin suggest possible reasons for this unexpected decrease in observed binding

affinity by 1Gap for heparin. The significant decrease in binding affinity observed for both

peptides 1 and 1Gap in binding an isolated 14mer heparin oligosaccharide was expected.

Previous work has demonstrated the significance of heparin length on binding affinity. It was

observed that the HIV-1 Tat protein immobilized to glutathione agarose columns bound 22 mer

heparin with a KD of 30 nM while it bound to a 6mer of heparin with a KD of 700 µM showing a

23-fold decrease in binding affinity with decreasing heparin oligosaccharide chain length (22).

Michaelis constants describing heparin binding to thrombin calculated in studies looking at

heparin mediated AT III inhibition of thrombin exhibited the same trend, showing decreases in

binding affinity from 7 nM, to 100 nM and 6 µM upon binding with heparin of molecular

weights 15,000, 4300, and 3200 Da, respectively (21). Studies using an internally quenched

fluorogenic peptide with engineered XBBBXXBX Cardin motifs of the sequence:

[ARKKAAKA]ARKKAAKAARKKAAKAARKKAAKAQ exhibited a KD for 4500 Da heparin

of 8 nM. This dropped to 732 nM upon binding studies with a synthetic pentasaccharide analog

of heparin (24). This approximately 100-fold decrease in binding affinity is similar to what was

observed upon titrating peptide 1 and 1Gap with 14mer heparin. The decrease in observed

binding affinity upon reduction in the heparin chain length is a statistical effect due to a

reduction in the number of overlapping binding sites on the heparin chain (25). The dissociation

constants between peptides 1 and 1Gap not only increase, but become indistinguishable upon

binding the heparin 14mer. This suggests that the higher binding affinity observed upon peptide

1 binding full length heparin was a result of a greater number of overlapping binding sites for

this peptide because the charge is uniformly displayed along the α-helix. The space between

positive charges on peptide 1Gap reduced the peptides ability to sample as many overlapping

binding sites along the heparin chain, as charged clusters need to line up with the clusters of

charge along the heparin chain, resulting in an observed weaker affinity of interaction. The

14mer heparin oligosaccharide is approximately the same length as peptides 1 or 1Gap when

these peptides are in an α-helical conformation. Thus, a large number of overlapping binding

43

sites cannot exist on this oligosaccharide and therefore the statistical effect of overlapping

binding sites on the observed dissociation constant is reduced.

The wide range of affinities observed in peptide-heparin binding interactions clearly

demonstrates that spatial display of positive charge is a very significant factor behind affinity of

peptide/protein and heparin interactions. The calculated apparent equilibrium dissociation

constants span three orders of magnitude despite the peptides possessing the same amino acid

composition and overall positive charge. The results suggest that heparin most favourably

participates in high affinity electrostatic interactions as an extended helical structure enabling it

to make contacts with localized continuous arrangements of positive charge density. Despite the

peptides not containing known heparin binding consensus sequences, high affinity binding was

observed. This report sheds new light on important factors in peptide heparin interactions and

offers insight into current heparin sensor development efforts (11,26). We have demonstrated

how binding affinity is highly tunable through subtle structural modifications. Tunable binding

affinity coupled with sensitive fluorescence would be a formidable combination in achieving the

sensitivity and selectivity required for highly desirable in vivo probing of heparin and heparan

sulfate glycosaminoglycans.

2.4 Materials and Methods

All peptides were synthesized by microwave assisted solid phase peptide synthesis on a 0.1

mmol scale using Fmoc protected amino acids and rink amide resin as the solid support. All

amino acids and coupling agents were purchased from Anaspec. Unfractionated heparin was

purchased from Celsus laboratories and 14mer heparin oligosaccharide was purchased from

Dextra laboratories. The quinolinium fluorophore 2-((N-methyl quinolinium)-6-yloxy) acetic

acid chloride (11) appended peptide constructs was synthesized according to Fig2-10:

44

Figure 2-10. The synthesis of quinolinium-peptide conjugates

Fluorophore-Peptide Coupling

2-((N-methyl quinolinium)-6-yloxy)acetic acid chloride (0.4 mmol) was first dissolved in DMF

(5 mL) by adding 0.4 mmol of KPF6 and heating the suspension at 500C for 10 min. Following

heating, N-hydroxysuccinimide (NHS) (0.4 mmol) and diisopropylcarbodiimide (DIC) (0.4

mmol) were added and the solution was heated again for 10 min at 50oC. The activated

fluorophore was cooled to room temperature. The solution was then added to the solid supported

peptide (0.1 mmol) and allowed to shake for 2 h. The efficiency of the coupling steps was

assessed by adding a sample of the beads to a solution of picryl sulfonic acid and observing the

emergence of red color indicating the presence of free amines.

Peptide Deprotection and Purification

The peptide was cleaved from the resin with a solution of trifluoroacetic acid TFA:TIS:H2O in a

9.5:0.25:0.25 ratio for 4 h. The solid phase resin was removed by filtration and the resulting

solution was concentrated, and the peptide precipitated by addition of cold diethyl ether followed

45

by centrifugation for 20 min. The resulting pellet was resuspended and washed with diethyl

ether. After drying, the pellet was dissolved in water and purified by preparatory reverse-phase

C18 HPLC using an acetonitrile/H2O (0.1% TFA) gradient of 0%–70% acetonitrile over 50 min.

The purified sample was lyophilized and later dissolved in deionized water. The concentration of

the peptide solutions were determined by measuring the A336 employing an extinction coefficient

of the attached quinolinium dye of 4006 M-1

cm-1

. MALDI-TOF MS analysis of the purified

peptides gave the expected masses in single protonated and sodiated forms.

Circular Dichroism Spectroscopy

All spectra were recorded on a JASCO 710 spectrometer in a 1 cm path length cuvette at room

temperature. Peptides (75 µM) were dissolved in buffer (50 mM phosphate buffer, pH 7.0, 150

mM NaCl) and spectra were then recorded from 190 nm to 260 nm. Blank spectra containing

buffer alone or buffer with Heparin were subtracted from the peptide spectra. Spectra were

recorded in the absence and presence of saturating levels of heparin (500 µM peptide binding

sites) added to fully complex the peptide. All CD spectra were acquired with a sensitivity of 100

mdeg, 1 nm resolution, and a 2 s acquisition time.

Steady State Fluorescence Spectroscopy

Measurements were performed on a Fluorolog1-3 spectrofluorometer, in 0.5 x 0.5 x 3 cm3

quartz fluorescence cuvettes. Excitation/ Emission monochrometers were set to 336/430 nm

respectively, and slit widths were adjusted to ensure counts per second over the course of the

titration fell between 2 x 103 and 3 x 10

6. All spectra were recorded in 50 mM phosphate, 150

mM NaCl buffer at pH 7.0. Individually prepared solutions were used to avoid the effects of

photobleaching. The solutions were prepared by mixing calculated aliquots of aqueous sodium

chloride, deionized water, and phosphate buffer from prepared stock solutions to afford the

desired final concentration in the fluorescence cuvette. A fluorescence scan was taken of the

solution to allow for buffer subtraction from the subsequent fluorescent scans containing

mixtures of peptide and heparin. Following this, an aliquot of the peptide was added from a stock

solution of known concentration to the cuvette. A defined amount of heparin was added from a

heparin stock solution of 50 mg/mL (164 units/mg from porcine intestinal mucosa) in deionized

46

water stored at 22oC and the fluorescence was recorded. The heparin stock concentration was

defined according to the number of peptide binding sites (see results) using an equivalence point

titration of peptide 1. Equivalence point titrations were carried out using peptide concentrations

at least ten times larger than the dissociation constant of the complex. The concentration and

volume of heparin required to fully complex this peptide allowed for a back calculation of the

concentration of peptide binding sites contained in the heparin stock solution (12.5 mM peptide

binding sites). By using this experimentally calculated concentration of binding sites and the

mass of heparin dissolved in a given volume of water, the molecular weight of the binding site

can be calculated. Nonlinear least squares fitting analysis of the binding data was generated using

Origin graphing software employing the following equations to determine KD:

Equation 1: pep][hep]

complex KD Equation 2:

Equation 3: [complex]2 – [pepT][complex]-[hepT][complex]-KD[complex] + [hepT][pepT] = 0

Substituting Eq (2) into Eq (3) and applying a quadratic equation [Eq. (4)] leads to Eq. (5)

solving for Fobs.

Equation 4: y = [complex] =

Equation 5: y = P1 x P2)- P1-x-P2)

(P3-P4)

P1 P4

[pep] = free peptide, [hep] = free peptide binding sites on heparin, [ pep T] = total peptide, where

the fluorescence observed Fobs = y, the total concentration of peptide P1 and heparin binding sites

(x) are known as are the maximum fluorescence (P3) and the minimum fluorescence (P4).

Iterative curve fitting yielded the dissociation constant KD (P2).

47

References

1. Mulloy, B., and Forster, M. J. (2000) Conformation and dynamics of heparin and heparan

sulfate, Glycobiology 10, 1147-1156.

2. Mulloy, B., Forster, M. J., Jones, C., and Davies, D. B. (1993) N.M.R. and molecular-

modelling studies of the solution conformation of heparin, Biochemical. Journal. 293,

849-858.

3. Mulloy, B., Forster, M. J., Jones, C., Drake, A. F., Johnson, E. A., and Davies, D. B.

(1994) The effect of variation of substitution on the solution conformation of heparin: a

spectroscopic and molecular modelling study, Carbohydrate Research 255, 1-26.

4. Capila, I., and Linhardt, R. J. (2002) Heparin–Protein Interactions, Angewandte Chemie

International Edition 41, 390-412.

5. Jin, L., Abrahams, J. P., Skinner, R., Petitou, M., Pike, R. N., and Carrell, R. W. (1997)

The anticoagulant activation of antithrombin by heparin, Proceedings of the National

Academy of Sciences 94, 14683-14688.

6. Cardin, A., and Weintraub, H. (1989) Molecular modeling of protein-glycosaminoglycan

interactions, Arteriosclerosis, Thrombosis, and Vascular Biology 9, 21-32.

7. Jayaraman, G., Wu, C. W., Liu, Y. J., Chien, K. Y., Fang, J. C., and Lyu, P. C. (2000)

Binding of a de novo designed peptide to specific glycosaminoglycans, FEBS letters 482,

154-158.

8. Ferran, D. S., Sobel, M., and Harris, R. B. (1992) Design and synthesis of a helix

heparin-binding peptide, Biochemistry 31, 5010-5016.

9. Margalit, H., Fischer, N., and Ben-Sasson, S. (1993) Comparative analysis of structurally

defined heparin binding sequences reveals a distinct spatial distribution of basic residues,

Journal of Biological Chemistry, 268, 19228-19231.

10. Sauceda, J. C., Duke, R. M., and Nitz, M. (2007) Designing Fluorescent Sensors of

Heparin, ChemBioChem 8, 391-394.

11. Jagt, R. B. C., Gómez-Biagi, R. F., and Nitz, M. (2009) Pattern-Based Recognition of

Heparin Contaminants by an Array of Self-Assembling Fluorescent Receptors,

Angewandte Chemie International Edition 48, 1995-1997.

48

12. Villanueva, G. B. (1984) Predictions of the secondary structure of antithrombin III and

the location of the heparin-binding site, Journal of Biological Chemistry 259, 2531-2536.

13. Vila, J. A., Ripoll, D. R., and Scheraga, H. A. (2001) Influence of lysine content and PH

on the stability of alanine-based copolypeptides, Biopolymers 58, 235-246.

14. Seyrek, E., Dubin, P. L., and Henriksen, J. (2007) Nonspecific electrostatic binding

characteristics of the heparin-antithrombin interaction, Biopolymers 86, 249-259.

15. Chakrabartty, A., Schellman, J. A., and Baldwin, R. L. (1991) Large differences in the

helix propensities of alanine and glycine, Nature 351, 586-588.

16. Sundaralingam, M., Drendel, W., and Greaser, M. (1985) Stabilization of the long central

helix of troponin C by intrahelical salt bridges between charged amino acid side chains,

Proceedings of the National Academy of Sciences 82, 7944-7947.

17. Stellwagen, E., Park, S.-H., Shalongo, W., and Jain, A. (1992) The contribution of

residue ion pairs to the helical stability of a model peptide, Biopolymers 32, 1193-1200.

18. García, A. E., and Sanbonmatsu, K. Y. (2002) α-Helical stabilization by side chain

shielding of backbone hydrogen bonds, Proceedings of the National Academy of Sciences

99, 2782-2787.

19. Vila, J. A., Ripoll, D. R., and Scheraga, H. A. (2000) Physical reasons for the unusual α-

helix stabilization afforded by charged or neutral polar residues in alanine-rich peptides,

Proceedings of the National Academy of Sciences 97, 13075-13079.

20. Ziegler, A., and Seelig, J. (2004) Interaction of the Protein Transduction Domain of HIV-

1 TAT with Heparan Sulfate: Binding Mechanism and Thermodynamic Parameters,

Biophysical Journal 86, 254-263.

21. Hoylaerts, M., Owen, W. G., and Collen, D. (1984) Involvement of heparin chain length

in the heparin-catalyzed inhibition of thrombin by antithrombin III, Journal of Biological

Chemistry 259, 5670-5677.

22. Rusnati, M., Tulipano, G., Spillmann, D., Tanghetti, E., Oreste, P., Zoppetti, G., Giacca,

M., and Presta, M. (1999) Multiple Interactions of HIV-I Tat Protein with Size-defined

Heparin Oligosaccharides, Journal of Biological Chemistry 274, 28198-28205.

23. Fromm, J. R., Hileman, R. E., Caldwell, E. E. O., Weiler, J. M., and Linhardt, R. J.

(1995) Differences in the Interaction of Heparin with Arginine and Lysine and the

Importance of these Basic Amino Acids in the Binding of Heparin to Acidic Fibroblast

Growth Factor, Archives of Biochemistry and Biophysics 323, 279-287.

49

24. Pimenta, D. C., Nantes, I. L., de Souza, E. S., Le Bonniec, B., Ito, A. S., Tersariol, I. L.

S., Oliveira, V., Juliano, M. A., and Juliano, L. (2002) Interaction of heparin with

internally quenched fluorogenic peptides derived from heparin-binding consensus

sequences, kallistatin and anti-thrombin III, Biochem. J. 366, 435-446.

25. Olson, S. T., Halvorson, H. R., and Björk, I. (1991) Quantitative characterization of the

thrombin-heparin interaction. Discrimination between specific and nonspecific binding

models, Journal of Biological Chemistry 266, 6342-6352.

26. Nitz, M., Rullo, A., and Ding, M. X. Y. (2008) Heparin Dependent Coiled-Coil

Formation, ChemBioChem 9, 1545-1548.

50

Chapter 3 Peptide-Glycosaminoglycan Cluster Formation Involving Cell

Penetrating Peptides2

3.1 Introduction

Cell penetrating peptides (CPPs) are generally known to be cationic peptides, typically less than

30 amino acids in length, which have the ability to penetrate cellular membranes within minutes

(1-4). The ability to penetrate cellular membranes has sparked widespread interest in the

application of CPPs as delivery vectors (5). Cellular delivery of various therapeutic agents has

been achieved though the synthesis of CPP-cargo conjugates, connected through covalent or non-

covalent interactions (2, 4, 6). In this study we focus on two common CPPs: Penetratin (Antp),

and TAT, as well as a third peptide previously synthesized, helical heparin binding peptide

HHBP (7), which has known glycosaminoglycan (GAG) affinity.

Antp is a well studied CPP. It is a 16 amino-acid peptide from the antennapedia protein

transduction domain and has attracted much attention for use as a delivery vector (8,9). Antp

potentially has seven positively charged residues at physiological pH and has been shown to bind

membranes composed of negatively charged lipids and to bind GAGs (9-11). Using circular

dichroism it was found that the Antp peptide undergoes a conformational transition from random

coil to a β-sheet upon binding to negatively charged DOPG lipids (12).

Another widely studied CPP is the TAT peptide which is derived from the HIV-1 transactivator

of transcription protein (13). TAT is 13 amino acids in length and is highly cationic due to the

presence of potentially eight positively charged residues at physiological pH. The TAT peptide

has been observed to facilitate cell uptake in both cell culture and in living animals (14).

2 Reproduced with permission from Rullo, A. Qian, J, Nitz, M (2011). Peptide-Glycosaminoglycan

Formation involving Cell Penetrating Peptides. Bioploymers ,95,722-731. Author Contributions: All

experiments performed and paper co-written by Rullo, A. DLS analysis carried out by Qian, J and Paper

co-written by Nitz, M. The introduction of the main text has been modified and figures have been added

from the supplementary information. The text from the materials and method sections in addition to the

results and discussion section remain unchanged.

51

Conformational studies using CD have demonstrated that TAT can bind negative DOPG lipids

and the GAG heparin in a random coil conformation (12, 14,15).

Dissecting the roles of energy dependent endocytotic pathways and energy independent direct

translocation pathways in CPP cell entry has been the focus of many studies (16). It is

indisputable that GAGs play a role in CPP cell uptake, as the CPPs are known to bind the GAG

heparin with nanomolar affinity, but how the GAGs affect the mechanism of cell uptake is not

clear (11, 15, 17-21). Experiments have shown the reduced internalization efficiency of CPPs

following the addition of exogenous heparin, in cells not expressing cell surface GAGs and in

cells treated with heparinase enzymes (10, 11, 18, 19, 22-25). The translocation of Antp and

TAT has been observed to be mediated by specific interactions with the heparan sulfate chains of

syndecan-4, belonging to the syndecan family of transmembrane proteoglycans (17). However,

in addition to these findings, experimental evidence has also been reported that indicates, in the

case of TAT, that uptake into the cell can occur even in the absence of cell surface GAGs (26-

28). It is probable that cell penetrating peptides differ in their degree of GAG dependence and

their mechanisms of uptake.

Jiao et al. have evaluated the internalization efficiencies of numerous cell penetrating peptides

employing a mass spectrometry based method capable of direct CPP quantification (29). The

cell penetrating peptides TAT and Antp, among others, were shown to differ significantly in the

involvement of translocation and endocytotic mechanisms, and in their dependence on GAGs for

uptake. When endocytosis was operative, at 37oC, and GAGs were present, an Antp analogue

showed cell uptake predominantly through an endocytotic mechanism (Fig3-1). In contrast the

TAT peptide was taken up mainly via an energy independent translocation pathway. Under these

conditions the Antp analog was also shown to have higher uptake efficiency than the TAT

peptide and it was suspected that GAG clustering at the cell surface may play a role (29).

52

Figure 3-1. A schematic representation of the different uptake mechanisms by which TAT

(translocation) and Antp (endocytosis) predominantly enter the cell highlighting a potential role

played by cell surface GAGs. (CPPs = cylinders, GAGs = branched structures)

The mechanism by which GAG binding may facilitate CPP uptake has been proposed to result

from the formation of clusters between surface GAGs and the cell penetrating peptides (11, 29-

31). In contrast to cationic peptide binding to monomeric GAG chains which is well known, the

ability of multiple peptides and multiple heparin chains to interact forming a cluster has not been

rigorously characterized. This is due to the analytical challenges that accompany the

characterization of peptide-GAG clustering due to the complex stoichiometry of binding within

the cluster, the heterogeneity of GAG structure, and the deconvolution of clustering from simple

peptide-monomeric GAG complexes. Peptide-GAG clusters have been observed previously in

solution with light scattering, but the rate of formation, the stability and the molecular

interactions leading to cluster formation have not been explored (11). These features of the CPP-

GAG clusters likely influence the mechanisms and efficiency of CPP cell entry. We set out to

characterize CPP-heparin clusters with respect to their rate of formation, peptide and heparin

dependence, size and stability.

53

3.2 Results

To study clustering that occurs between heparin and CPPs, three cationic peptides, a helical

heparin binding peptide (HHBP), TAT and Antp were synthesized (Table 3-1). HHBP is 30

amino acids in length, containing eight positively charged lysine residues separated by alanine

residues. HHBP has been shown to bind heparin in a helical conformation with a dissociation

constant of 640 nM at physiological ionic strength (7). This peptide was synthesized with either a

fluorescent dye, 7-methoxycoumarin-3-carboxylic acid, or a fluorescence quencher, DABCYL

acid, appended via amide formation to the ε-amino group of a C-terminal lysine side chain. The

dyes are attached at a position in the peptide’s first heptad repeat, directed away from the heparin

binding interface. The known cell penetrating peptides TAT and Antp, with dissociation

constants for binding heparin of 443 nM and 338 nM determined using ITC (11), were also

synthesized and conjugated to the same fluorophore or quencher on the side chain of a C-

terminal lysine residue (Table 3-1). The dissociation constant for the TAT peptide for heparin

was determined using quinolinium fluorescence with the method reported previously, and was

found to be in close agreement (KD = 517 nM (unpublished results)) with that determined by ITC

in literature (KD = 443 nM) (7,11).

Table 3-1. The sequences of the three cationic heparin binding peptides used in this study

Peptide KD Heparin

(nM)

Sequence

HHBP 640 AcGGKAAKAKmcoumAKAAKAAAKAAKAAAKAAKAAA

AcGGKAAKAKDABAKAAKAAAKAAKAAAKAAKAAA

TAT 443 AcGGRKKRRQRRRPPQKm-coum

AcGGRKKRRQRRRPPQKDAB

Antp 338 AcGGRQIKIWFQNRRNleKWKKKm-coum

AcGGRQIKIWFQNRRNleKWKKKDAB

54

Heparin-Sepharose Chromatography

Heparin-sepharose affinity chromatography was used to qualitatively characterize the relative

binding affinities of the three coumarin labeled cationic peptides for heparin. This technique has

been applied previously to obtain relative binding affinities for peptide-heparin interactions

(32,33). The three cationic peptides bound with high affinity to the immobilized heparin and

eluted from the column at NaCl concentrations between 1.8-2.0 M (Fig3-2). The elution profiles

varied between peptides with Antp eluting as an exceptionally broad peak. The same amount of

each coumarin labeled peptide was analyzed and the degree of peak broadening is responsible for

the differences in absorbance intensity observed. Given the affinities of the peptide-heparin

interactions determined in dilute solution it was expected that the HHBP peptide would elute

prior to the TAT peptide. However, the observed elution order was TAT-HHBP~Antp,

suggesting that TAT has the lowest affinity and that HHBP and Antp have a comparable heparin

binding affinity. To probe the possible origins of this discrepancy the chromatography was

repeated using a gradient of GdnHCl (0-3M) as the elutent. The peptide elution profiles became

sharper with greater separation between peaks. The peptides also eluted in the expected order

based on the previously determined dissociation constants (Fig3-2). The increase in peak

dispersion and the change in the order of elution with GdnHCl, a chaotropic agent, suggests the

presence of H-bonding and hydrophobic contributions to heparin binding that are disrupted by

GdnHCl as well as the expected ionic interactions (34-36). The presence of hydrophobic

interactions within the heparin-peptide complex was not expected, as interactions between

cationic peptides and heparin are primarily driven by the polyelectrolyte effect (37,38). A

possible explanation for the discrepancy in peak dispersion, the elution order and the apparent

binding affinities among the three cationic peptides is that these peptides are clustering with

multiple heparin chains on the heparin sepharose column and that there are hydrogen bonding

and hydrophobic contacts between peptides within these clusters.

Dynamic light scattering

To investigate the nature of the CPP-heparin clusters, dynamic light scattering (DLS) was used to

evaluate the hydrodynamic radii of the peptide-heparin complexes formed in the peptide

solutions (10 µM) with either an equimolar or a large excess (500x) of unfractionated heparin.

55

Under conditions of equimolar heparin, HHBP and Antp form complexes with heparin having a

broad distribution of hydrodynamic radii centered at approximately 80 nm (Fig3-3). Complexes

of this size must consist of multiple peptide and heparin chains (Rh Heparin alone approx. 0.45

nm) (39). Interestingly, the TAT peptide under the same conditions formed clusters with a

narrower size distribution centered at approximately 40 nm (Fig3-3). Previous DLS studies on

Antp and TAT under conditions of half saturation of the heparin (91 µM) reported the presence

of clusters having hydrodynamic radii of 94 and 129 nm respectively (11). It was rationalized

that the aggregate formed between TAT and heparin giving rise to the peak at 94 nm consisted of

up to 2.1 x 105 heparin molecules each complexed with 6 or 7 TAT peptides (11). In the case of

peptide-heparin complexes formed with Antp (10 µM) and HHBP (10 µM) under conditions of

excess heparin (5 mM) the distribution of aggregates formed broadens and the radii are

approximately half those observed at equimolar heparin concentrations. In the case of HHBP

under conditions of 5 mM heparin, a smaller peak is also present with a hydrodynamic radii of

4.5 nm possibly representing a smaller cluster which results from the disaggregation caused by

the large excess of heparin. TAT-heparin aggregates were not detected under conditions of

excess heparin, suggesting that the aggregates were too small in size to be detected or did not

form under these conditions.

Cluster stability by fluorescence spectroscopy

Given the presence of large peptide heparin complexes which differed significantly between the

CPPs, steady state fluorescence experiments were carried out to evaluate the thermodynamic and

kinetic stability of the peptide-heparin clusters. Under conditions where all of the peptide would

be bound to heparin (250 nM DABSYL labeled peptide, 250 nM coumarin labeled peptide, 5 µM

heparin) the fluorescence was monitored. Upon the addition of heparin to each of the peptide

solutions, a rapid drop in fluorescence intensity was observed with the majority of the

fluorescence quenching occurring too quickly to be resolved by steady state fluorescence.

Equilibrium was achieved after 10-30 seconds, giving the maximal fluorescence quench (Fig3-

4). The percentage of fluorescence quenching, with respect to the pre-heparin peptide solution,

observed with the TAT peptide (~43%) was less than that observed for HHBP (~63%) and Antp

(~73%) (Fig3-4). The reduced percentage of fluorescence quenching observed for the TAT

peptide is attributed to a lower degree of cluster formation as was observed with DLS, when

56

compared to the HHBP and Antp peptides. The lower degree of cluster formation potentially

represents the formation of smaller clusters sizes and/or the formation of lower numbers of

clusters. The rapid decrease in fluorescence observed for all three peptide solutions

accompanying the formation of peptide-heparin clusters is attributed to the close proximity of the

coumarin and DABSYL labeled peptides within the clusters.

Figure 3-2. Heparin sepharose affinity chromatogram of methoxycoumarin labeled HHBP, TAT,

and Antp. A) Elution with NaCl (0M, 0-35 min, 0-3 M, 35-65 min) B) elution with GdnHCl

gradient (0M, 0-35 min, 0-3M, 35-65 min). ■ HHBP , ● TAT , and ▲ Antp, all solutions

contain phosphate buffer (50 mM, pH 7) . The increasing salt gradient is shown on the right y-axis

for clarity.

57

Figure 3-3. The hydrodynamic radii of clusters present in peptide-heparin solutions. Coumarin

peptides (10 µM): ■ HHBP, ● TAT, ▲ Antp in phosphate buffer ( 50 mM, pH 7, 150 mM NaCl).

A) Heparin (10 µM) and B) Heparin (5 mM).

58

Figure 3-4. Fluorescence quenching of labeled peptide heparin solutions. Association experiments

(decreasing fluorescence) were initiated by adding heparin (5 µM) to DABSYL-peptide (250 nM),

coumarin peptide (250 nM) solution. Dissociation experiments (increasing fluorescence) initiated

by the addition of excess heparin (5 mM) to peptide-heparin solutions (Heparin 5 µM, dabsyl-

peptide 250 nM, coumarin peptide 250 nM). A solution of heparin alone gave no fluorescence

signal. All experiments were run in triplicate and were carried out in phosphate buffer (50 mM,

pH 7.0, 150 mM NaCl), Excitation 350nm, Emission 415nm, ▬ Antp , − − HHBP, ▬ TAT.

To confirm that peptide-peptide proximity was responsible for the observed fluorescence

quenching, the effects of diluting the donor labeled HHBP peptide with unlabelled HHBP

peptide to eliminate self-quenching (40) on fluorescence was determined, while maintaining a

fixed overall peptide and heparin concentration. The percentage of fluorescence quenching

decreased with increasing dilution of the labeled peptide confirming that fluorescence quenching

observed is due to the proximity between peptides and not a simple heparin binding event or a

conformational change in the labeled peptide (Fig3-5). The percentage of fluorescence

quenching (~15%) approached at high dilution of the labeled HHBP peptide by unlabelled

HHBP (1:50) peptide represents the fluorescence quenching that occurs in the absence of

fluorophore-fluorophore interactions, likely due to changes in solvation of the fluorophore upon

binding and clustering with heparin.

59

Figure 3-5. The quenching observed due to peptide-peptide interactions upon dilution of coumarin

–HHBP peptide with unlabeled HHBP peptide (total peptide concentration 10 µM) in the presence

of a fixed concentration of heparin (10 µM).

Following association experiments, the peptide-heparin clusters were treated with excess heparin

to determine if clusters could be dissociated. After the formation of the clusters (250 nM

DABSYL labeled peptide, 250 nM coumarin labeled peptide, 5 µM heparin), a large excess of

heparin (5 mM) was added and the fluorescence time course was acquired. An increase in

fluorescence approaching the intensity of the heparin free peptide solution was observed for all

peptide heparin solutions. The increase in fluorescence occurred on a time scale of minutes with

an estimated half life of approximately 100 sec for HHBP and Antp. Interestingly, the

dissociation of the TAT-heparin clusters occurred too quickly to be resolved (Fig3-4). These

experiments demonstrate that TAT-heparin clusters are less kinetically stable than clusters

involving HHBP and Antp peptides. Given that the TAT-heparin clusters completely dissociate

in the dead time of the experiment (< 5 s) this rate must be approximately two orders of

magnitude faster than the dissociation rate for clusters involving HHBP and Antp. As the

formation rates of the peptide-heparin clusters appear similar, (Fig3-4) the difference in

dissociation rates suggest that the clusters involving TAT are a least two orders of magnitude less

stable than clusters involving HHBP and Antp.

To further evaluate the stability of the CPP-heparin clusters, the fluorescence of the mixed

fluorophore-quencher peptide solutions was determined over a range of heparin concentrations.

0

10

20

30

40

50

60

70

0 2 4 6 8 10

% Q

uen

chin

g o

f Fl

uo

resc

en

ce

labelled peptide (µM)

60

The percent fluorescence quench of the peptide solutions (500 nM DABSYL labeled peptide,

500 nM coumarin labeled peptide) was quantified after equilibration with heparin. Fluorescence

quenching of the peptide solutions was observed to decrease with increasing concentrations of

heparin. This observation was attributed to disaggregation of the peptide-heparin cluster at

increasing concentrations of heparin into monomeric peptide-heparin complexes (Fig3-6). The

minimal fluorescence quenching observed upon disaggregation of TAT-heparin clusters occurred

in solutions containing > 100 µM heparin. The minimal fluorescence quenching observed upon

disaggregation of HHBP-heparin clusters occurred in solutions containing > 400 µM heparin and

half of the quenching of the Antp solution remained at 800 µM heparin. In agreement with the

kinetic data, clusters formed between heparin and the TAT peptide were markedly less stable

requiring significantly less heparin for disaggregation than clusters involving the HHBP and

Antp peptides. These observations also confirm that the heparin-peptide clusters can form at low

concentrations of peptide and low concentrations of heparin but can be disrupted with excess

heparin.

The dependence of cluster formation on peptide concentration (0.5 – 25 µM) was evaluated at

high heparin concentrations (5mM) (Fig3-7). The percentage of fluorescence quenching

increased with increasing concentrations of peptide in the solution. The maximal percentage of

quenching for the Antp was reached at 5 µM peptide and between 7-10 µM peptide for HHBP.

In the case of TAT, concentrations of 25 µM peptide were insufficient to achieve the saturation

of

61

Figure 3-6. Fluorescence of labeled peptide solutions in the presence of heparin. Peptides

(DABSYL-peptide (500 nM), coumarin peptide (500 nM)) All experiments were run in triplicate

and carried out in phosphate buffer (50 mM, pH 7.0, 150 mM NaCl), Excitation 350nm, Emission

415nm, ▲ Antp , ■ HHBP, ● TAT .

the fluorescence quenching. The data for Antp and HHBP could be fit to an isodesmic

aggregation model to obtain association constants for the addition of heparin-peptide monomers

to a growing peptide-heparin cluster. Isodesmic models have been applied previously to

characterize the stability of FGF1 and FGF2 oligomerization in the presence of GAGs and

solution oligomerization of the HIV-1 rev protein (41,42). The calculated dissociation constants

for Antp and HHBP were 0.68 ± 0.12 µM and 1.2 ± 0.2 µM respectively. An association constant

for the TAT-heparin cluster could not be determined accurately due to the significantly lower

stability of the cluster.

Figure 3-7. Fluorescence of peptide solutions in the presence of excess heparin. Solutions contain

heparin (5 mM) and an equimolar mixture of coumarin and DABSYL labeled peptides, the total

peptide concentration is given. The solid line represents nonlinear regression analysis using a

hyperbolic function assuming an isodesmic model. All experiments carried out in phosphate buffer

(50 mM, pH 7.0, 150 mM NaCl), Excitation 350 nm, Emission 415 nm, ▲ Antp , ■ HHBP, ●

TAT .

To confirm that clusters observed at low and high heparin:peptide ratios interconvert,

demonstrating the reversibility of the clustering process, clusters formed at peptide

concentrations ranging from 1 µM to 25 µM in the presence of heparin (200 µM) were

62

dissociated with the addition of excess heparin (5 mM). The final fluorescence observed was

similar to that found after a single addition of excess heparin to the corresponding concentration

of peptide indicating that the clusters readily equilibrate in minutes at room temperature (Fig3-7

vs Fig3-8).

Figure 3-8. Fluorescence change upon addition of excess heparin to preformed heparin-HHBP

aggregates. All clusters formed using 200 µM heparin at indicated peptide concentration and

dissociated with addition of heparin (5 mM): ♦ 1 µM , ■ 5 µM , ▲ 8 µM, ▬ 10 µM ●, 25 µM .

Data is normalized to the fluorescence of peptide solution in the absence of heparin.

3.3 Discussion and Conclusions

The experiments reported here were all conducted at concentrations well above the dissociation

constants for monomeric peptide-heparin interactions, determined in highly dilute solutions. The

micromolar peptide concentrations and buffers used also are comparable to conditions that would

be employed in a CPP cell uptake experiment. Under these conditions, DLS analysis of the CPP-

heparin solutions indicates large complexes are formed whose size can only be explained by the

interaction of multiple heparin and peptide chains (Fig3-3). These results are consistent with

previous DLS studies on Antp and TAT peptide-heparin solutions under conditions of 50%

saturation of the available heparin binding sites (91 µM peptide binding sites, 45 µM peptide)

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

0 100 200 300 400 500

No

rmal

ized

Flu

ore

sce

nce

41

5 n

m

time (s)

63

(11). The size of the clusters observed depends on the relative proportions of peptide and heparin

in solution, with smaller clusters formed in the presence of excess heparin.

The close peptide-peptide proximity within the peptide-heparin clusters is evident by the

fluorescence quenching observed upon the addition of heparin to a solution of the labeled CPP

peptides. Characterization of the thermodynamic and kinetic stability of the peptide-heparin

clusters indicates that clustering is a reversible process occurring at peptide concentrations as low

as 500 nM. Peptide-heparin clusters involving Antp and HHBP were more thermodynamically

and kinetically stable than TAT-heparin clusters (Fig3-4, 3-6, 3-7). CPP-heparin cluster

formation for Antp and HHBP could be described using an isodesmic model for the association

of monomeric peptide-heparin complexes with a growing peptide-heparin cluster (Fig 3-7). This

model assumes a single equilibrium constant for each addition of a monomeric peptide-heparin

complex to the growing cluster. The determined isodesmic dissociation constants for Antp and

HHBP are in the low micromolar range (Fig3-7). The higher thermodynamic and kinetic stability

of HHBP and Antp clusters compared to those involving the TAT peptide is in agreement with

the observation of smaller clusters formed between TAT and heparin observed by DLS.

Cluster formation under conditions of large excesses of heparin (Fig3-7) indicates that peptide-

peptide interactions are a major driving force for cluster formation. Under these conditions each

heparin chain would be expected to bind a single peptide. Thus, the clusters are likely formed

from monomeric peptide-heparin complexes associating to form the cluster. If the intracluster

contacts were due to peptides cross linking multiple strands of heparin, it should be possible to

dissociate these clusters with excess heparin. As it isn’t possible to dissociate the clusters formed

at high peptide concentrations (25 µM) (Fig3-7, 3-8), peptide-peptide interactions must be

responsible for the majority of the association energy associated with cluster formation at high

heparin-peptide ratios. At lower heparin-peptide ratios more than one peptide would be expected

to bind each heparin chain, bringing these multipeptide-heparin complexes together. This would

result in a multivalent interaction between the multipeptide-heparin complexes leading to cluster

formation at low peptide and low heparin concentrations.

These clusters formed at low heparin: peptide ratios can be dissociated with excess heparin to

form monomeric complexes with one peptide binding a single heparin chain (Fig3-4, 3-8).

64

When a sufficient concentration of monomeric peptide-heparin complexes are present these

single peptide bound monomeric heparin chains can associate to form clusters characterized in

Fig 3-7 using the isodesmic model. Clustering models have been proposed previously to explain

cationic dye quenching phenomena which predominately involve dye-dye contacts with each dye

on a separate heparin chain (43-46). While the structural features of the CPPs responsible for

formation of stable heparin clusters have not been investigated in detail from this study it appears

to stem from the ability of the peptide to form non-ionic, likely hydrophobic interactions with

other heparin-bound peptides. The presence of non-ionic contributions to the peptide-heparin

clusters was confirmed by the sensitivity of heparin affinity chromatography to the nature of the

eluting salt solution with changes in elution profile and a differing order of elution observed with

NaCl and GdnHCl (Fig 3-2).

The presence of hydrophobic dyes appended to the peptides studied potentially introduces a non-

natural hydrophobic element which may increase the degree of hydrophobic interactions and

therefore aggregation between the peptides. In this study however, all three peptides have the

same fluorophores attached thus the differences observed are most likely due to different

properties inherent in the peptide sequences. Previous detailed work in the Gellman laboratory

with coiled coil peptides have also shown the coumarin dyes to be minimally perturbing in

peptide-peptide interactions (40). Although the overall charges are similar in the CPPs studied,

Antp and HHBP consist of a higher proportion of non-ionic amino acids than the TAT peptide,

providing a greater hydrophobic surface for interactions within the cluster. The difference in

hydrophobic content of the CPPs may be the key to formation of stable heparin-peptide clusters.

Cell surface GAGs consist of mainly heparan sulfate and chondroitin sulfate. Although these

GAGs are less sulfated than heparin, heparin provides a useful mimic as to the relative binding

affinities of peptides for GAGs (38). Antp has been observed to enter cells through GAG-

dependant endocytotic pathways to a larger degree than TAT, with TAT being taken up by the

cell predominantly through direct translocation (29). The differing preferences for uptake

pathways in the presence of GAG may result from significant differences in the stability of

clusters with GAGs formed at the cell’s surface (Fig3-9). Our findings clearly illustrate the

increased thermodynamic and kinetic stability of clusters with heparin involving Antp compared

to clusters involving the TAT peptide. The higher stability of Antp-GAG clusters may sequester

65

the peptide in the glycocalyx allowing for energy dependent endocytosis to occur, while the TAT

peptide may form and dissociate from GAG clusters at the cell’s surface allowing the peptide to

access the cell membrane for direct translocation. In more complex systems where the CPP is

used as a delivery vector, the cargo attached, or even the type of fluorophore conjugated to the

CPP, may affect the stability of the GAG-peptide clusters formed, in turn affecting the

mechanism of cell uptake (40).

Figure 3-9. Our proposed model that employs GAG clustering to explain differences in CPP cell

uptake mechanisms

We have demonstrated that peptides with similar binding affinity for heparin can differ

significantly in their ability to cluster with heparin. At near equal molar heparin-peptide ratios

cluster formation can occur at very low concentrations (nM). The CPP Antp formed considerably

more stable clusters with heparin compared to clusters formed from the TAT peptide. The

stability of the clusters correlates with the number of hydrophobic amino acids in the peptides’

sequences. The differences in cluster stability observed between these two peptides provide a

possible explanation for their differing cell uptake routes.

66

3.4 Materials and Methods

Materials and peptide synthesis

All Fmoc protected amino acids and coupling reagents were purchased from Anaspec. Rink

amide MBHA resin (0.67 meq/g) was used as the solid support and was also purchased from

Anaspec. The three cationic peptides were synthesized on a 0.1 mmol scale by microwave

assisted solid phase peptide synthesis using a CEM liberty peptide synthesizer. The fluorescent

dye 7-methoxycoumarin-3-carboxylic acid was synthesized according to a known literature

procedure (47) and the fluorescent quencher [4-((4-(dimethylamino)phenyl)azo)benzoic acid]

(DABSYL acid) was commercially available from Anaspec. Unfractionated heparin was

purchased from Celsus laboratories isolated from pig intestinal mucosa and was obtained as the

sodium salt. The HiTrap HP heparin sepharose affinity column (1 ml) was purchased from GE

healthcare.

Fluorophore-peptide coupling

The amino acid Fmoc-N'-methyltrityl-L-lysine (MTT-lysine) was incorporated into the peptides

at the C-terminus during solid phase peptide synthesis (sequences in Table 3-1). After removal

of the MTT protecting group the methoxycoumarin fluorescent donor and DABSYL acid

quencher dyes were coupled to the peptides. Selective deprotection of the MTT-protected lysine

side chain was carried out on the solid supported peptide using a solution of 1% TFA in DCM.

The resin was suspended in this solution for 30 min and then filtered. This was repeated until no

further yellow color developed in the 1% TFA solution. The resin was subsequently washed

with 5% DIPEA/DMF prior to the dye coupling step. Coupling was carried out on the solid

support (0.1 mmol) with a solution of 0.4 mmol dye/ 0.4 mmol HATU /5% DIPEA /5 ml DMF

with vigorous shaking at room temperature for 4 hours. The efficiency of the coupling steps was

assessed by adding a sample of the resin to a solution of picryl sulfonic acid/DIPEA/DMF and

observing the emergence of red color indicating the presence of free amines.

67

Peptide deprotection and purification

The peptides were cleaved from the resin with a solution of trifluoroacetic acid TFA: TIS: H2O

in a 9.5: 0.25: 0.25 ratio for 4 h. The solid phase resin was removed by filtration, and the

resulting solution was concentrated and precipitated by the addition of cold diethyl ether. The

precipitate was collected by centrifugation. The resulting pellet was resuspended and washed

with diethyl ether. After drying, the pellet was dissolved in water and purified by preparatory

reverse-phase C18 HPLC using an acetonitrile/H2O (0.1% TFA) gradient of 0%–70%

acetonitrile over 50 min. The purified sample was lyophilized and dissolved in deionized water.

The concentrations of the peptide solutions were determined by measuring the A350 for

methoxycoumarin ( = 19000 M-1

cm-1

) and by measuring the A420 for the DABSYL substituted

peptides ( = 33000 M-1

cm-1

). MALDI-TOF MS analysis of the purified peptides gave the

expected masses in single protonated and sodiated forms.

Heparin sepharose affinity chromatography

The relative heparin binding affinity of the three cationic peptides was qualitatively assessed

using heparin sepharose affinity chromatography containing immobilized heparin from pig

intestinal mucosa at a density of 10 mg/ml. The column was adapted to a Gilson HPLC and

equilibrated with a solution of phosphate buffer (50 mM, pH 7.0) prior to injection for a period

of 30 min. Peptides labelled with methoxycoumarin were injected individually (150 µl of a 240

µM solution) at a flow rate of 0.5 ml/min. The elution consisted of an isocratic step from 0-35

min with phosphate buffer (50 mM, pH 7.0) followed by a gradient of 0-3 M NaCl (or 3M

GdnHCl) in phosphate buffer (50 mM, pH 7.0) from 35-65 min followed by a 45 min wash with

3 M NaCl or 3 M GdnHCl. Peptide elution was monitored using absorbance at 350 nm.

Dynamic light scattering

Right-angle dynamic light scattering (DLS) measurements were carried out at room temperature

23.0 ± 0.5℃ using an instrument from ALV described previously (48). Size information was

obtained by analysis of the autocorrelation function both by CONTIN and second cumulants

(49), using software included with the ALV instrument. Experiments were conducted on

68

solutions of each coumarin labelled peptide (10 µM) and combined with equimolar or excess

unfractionated heparin (5 mM) in phosphate buffer (50 mM, 150 mM NaCl, pH 7). Prior to

analysis, each solution was filtered individually using a 0.22 µm MILLEX-GP syringe filter to

remove dust and larger particulates.

Fluorescence Spectroscopy

Measurements were performed on a Perkin-Elmer LS-50B Luminescence Spectrophotometer

using a 45 mm x 12.5 mm x 12.5 mm quartz fluorescence cuvette pretreated with silicote. For all

fluorescence experiments the excitation and emission monochrometers were set to 350 nm and

415 nm respectively. Analyte concentrations were maintained within the linear response range

of the instrument. All spectra were recorded in phosphate buffer (50 mM, 150 mM NaCl, pH

7.0) using individually prepared solutions. The solutions were prepared for analysis by mixing

aliquots of aqueous sodium chloride (3 M), deionized water, and phosphate buffer (0.5 M) to

afford the desired final concentrations. Fluorescence spectra were corrected by subtraction of

buffer-only spectra.

Fluorescence time course studies

A solution of peptide consisting of equimolar coumarin and DABSYL labelled peptides was

prepared as described above a heparin was then added from a heparin stock solution (50 mg/mL,

164 units/mg) in deionized water stored at -200C and the fluorescence time course was recorded.

The heparin stock concentration was defined according to the number of peptide binding sites

using an equivalence point titration described previously (7). The molecular weights of the

heparin binding sites for Antp, TAT and HHBP were determined to be approximately equal and

a molecular weight of 4000 Da was used in subsequent heparin concentrations. Dissociation

experiments were conducted by adding a large excess of heparin (5 mM) to an equilibrated

solution of peptide and heparin. After mixing, the fluorescence intensity was followed collecting

fluorescence intensity readings at 1 sec intervals.

69

Determination of isodesmic association constants

Individual solutions of equimolar DABSYL and coumarin peptides were prepared in the

presence of heparin (5 mM). After equilibration (30 s) the fluorescence emission spectra were

recorded from 390 to 460 nm. The percentage of quenching of fluorescence, defined as final

fluorescence/fluorescence before heparin addition, was determined. The equilibrium binding

data was then fit to a hyperbolic function assuming an unlimited isodesmic association model.

Nonlinear least squares fitting analysis of the binding data using Origin graphing software

employed the following equations to determine the isodesmic association constant K, M-1

(50) :

(1)

CT = total concentration of peptide, C = concentration of free peptide, K = isodesmic association

constant

(2)

where Qobs = the observed percentage of fluorescence quenching, Qmax = the maximum

percentage of fluorescence quenching, Qmin = the minimum percentage of fluorescence

quenching. Eq 2 is then substituted into Eq 3 and the quadratic equation is applied

(3)

Rearrangement of the quadratic and substitution with equation 2 results in equating the

dependant variable y, to the observed percentage of fluorescence quenching (y = Qobs ) with the

total concentration of peptide set as the independent variable (x = CT )

CT = C

1 KC2

70

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75

Chapter 4 The synthesis of thioester carbohydrate conjugates as selective

lectin labelling agents3

4.1 Introduction

Carbohydrates play diverse roles in biological processes via selective binding interactions to

their cognate carbohydrate binding protein. This important class of biomolecular interactions is

exemplified by the interactions of Siglecs, Galectins and Selectins which control many aspects of

the immune response (1,2). The development of selective probes for carbohydrate binding

proteins would provide useful tools to further understand their complex biological roles. Early

labelling strategies for lectins focused on photoaffinity techniques (3-6). More recently elegant

examples using catalytic affinity based labelling approaches have been developed. These

approaches use a combination of a N,N-dialkyl-4-aminopyridine catalyst and a thioester

derivatized fluorophore to facilitate the selective acylation of the lectin of interest (7). Here, we

demonstrate that readily synthesized carbohydrate thioester conjugates can be used as selective

lectin labelling agents both in vitro and in E. coli.

Nature uses thioester chemistry extensively for selective acylation reactions. Perhaps the most

well known example being from intein-mediated protein splicing, which inspired the

development of native chemical ligation, now extensively used in protein synthesis (8). Other

examples include acetylation, ubiquitinylation and the C3/C4 proteins of the complement system

(9-11). The use of thioesters to perform these transformations may be due to thioesters being

more reactive towards nitrogen and sulphur nucleophiles while having similar reactivity to

hydroxide when compared to equivalent oxo-esters (12).

Thioesters have shown promise as ligand directed protein labelling agents. Glutathione

transferase has been site specifically modified at tyrosine and lysine residues using glutathione

based thioesters containing aryl substituents (13,14). Antiviral compounds targeting the HIV

3 Reproduced with permission from Rullo, A., Beharry, A. A.Gomez-Biagi, R., Zhao, X., Nitz,

M. 2012. Site-Selective affinity labeling of maltose binding protein in bacterial cells.

ChemBiochem, In Press.

76

nucleocapsid protein have included p-mercaptobenzamide thioesters which have been shown to

selectively acylate the protein (15). Recently, the amyloidogenic transthretin protein was

selectively labelled with fluorescent stillbene aryl thioesters at a pKa perturbed lysine residue

(16). The Flag-Tag peptide has been labeled with a thioester derived nickel chelator (17).

The labeling construct developed here is composed of a carbohydrate ligand and a fluorescent

thioester acyl donor. Maltose binding protein (MBP) was chosen as the lectin target for

evaluation of the labeling reaction. MBP is an E. coli periplasmic protein which binds

maltodextrins with micromolar affinity (1-80 µM). MBP is commonly used as a fusion probe for

the expression of poorly behaved proteins as it aids in protein solubilisation (18-20). MBP also

acts as a convenient purification tag as it displays high affinity for amylose resin (21).

4.2 Results and Discussion

Synthesis of thioester labeling constructs

Two sequential reactions allowed for the rapid synthesis of the thioester functionalized

carbohydrate ligands (Scheme 4-1). The free hemiacetals were first condensed with hydrazide

derived thiols (50-85% yield). Glycosyl hydrazides are readily formed under concentrated

reaction conditions with mild acid catalysis (22). The glucosylhydrazides are known to be stable

at neutral or basic pH for extended periods of time (months) and form preferentially the β-

glucosides (23). After isolation of the glucosyl hydrazides, the desired thioesters were accessed

by a thioester exchange reaction with a water soluble diethylaminocoumarin thioester to give the

desired labeling conjugates (10-40% yield). Due to the equilibrium conditions in each step, the

yields are highly dependent on the reaction conditions and the solubility of reaction components

(24,25). Aryl thioesters are less stable than alkyl thioesters thus a larger excess of reagent was

required and lower yields were obtained for the aryl thioester conjugates (24,25). The simplicity

of the synthesis should facilitate the generation of other labelled carbohydrates that may only be

available in small quantities from natural sources.

77

Scheme 4-1. Synthesis of the thioester labeling conjugates

MBP labeling in vitro

Maltose and lactose conjugates were synthesized to probe the selectivity of the MBP labeling

reaction. The thioesters, an alkyl and two aryl species, have different reactivities and geometric

preferences (Fig4-1). The diethylaminocoumarin (DAC) acylating agent was selected as it

provides a useful fluorescence handle as well as distinctive UV-spectral changes to monitor the

labeling reactions (26). Following synthesis of the constructs, labelling reactions were carried out

with purified MBP and the reaction progress was monitored with UV spectroscopy. The DAC

thioesters absorb maximally at 455 nm while the hydrolysis products, DAC carboxylates, absorb

maximally at 409 nm and amide or ester products absorb maximally at approximately 430 nm.

Incubation of MBP (5 µM) with the conjugates 2M or 3M (10 µM) in HEPES buffer (10 mM,

pH 8, 22 °C), resulted in a UV maxima which blueshifted with time, while little change was

observed with conjugate 1M (Fig4-2).

78

Figure 4-1. Structures of thioester constructs

These results suggested that the aryl thioesters were reactive under the labelling conditions. The

lack of reactivity displayed by conjugate 1M is likely due to the alkyl thioester being a less

reactive acylating agent as has been observed previously in the application of aryl and alkyl thiol

catalysts during native chemical ligation reactions (24). A greater rate and magnitude of UV

maxima shift was observed with conjugate 2M over conjugate 3M correlating well with the

expected reactivities of these thioesters. Construct 2M is expected to be more reactive, due to the

electron withdrawing hydrazide which reduces the pKa of the corresponding thiol by

approximately 1 pKa unit (25,27). Incubation of MBP with the lactose construct 3L gave only a

minimal blueshift in absorbance with time suggesting a selective acylation of MBP with 3M.

Incubation of MBP with the more reactive lactose construct 2L gave a similar rate of absorbance

shift to construct 3M, suggesting that hydrolysis or non-specific labelling may be occurring (Fig

4-2). Decomposition with an approximate half life of 7 h under the reaction conditions was

confirmed for construct 2M while construct 3M was stable for over 20 h (Fig4-3, 4-4).

79

Figure 4-2. Change in UV maxima (nm) during labeling. Thioester (10 µM), MBP (5 µM), HEPES

buffer (10 mM, pH 8, 22 °C) ● 2M ■ 3M ♦ 2L ▼ 3L ▲ 1M. To aid visualization, a line is drawn

through the data.

Figure 4-3. Stability of construct 3M: 3M (10 µM) in HEPES buffer (10mM, pH 8) at 25 °C. No

blue shift in UV maxima with time. Decrease in signal intensity is due to partial precipitation of

3M.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

350 400 450 500

Ab

sorb

ance

wavelength (nm)

T = 2hr

T = 4hr

T = 8hr

T = 12 hr

T = 16hr

T = 20 hr

80

Figure 4-4. Stability of construct 2M: construct 2M (10 µM) in HEPES buffer (10 mM, pH 8). Blue

shift in UV maxima with time is evident.

To confirm that specific labelling occurred with construct 3M, labelled MBP was purified from

the reaction mixture by size exclusion chromatography. Absorbance spectra of the purified

protein showed an absorbance band centered at 428 nm suggesting an amide or ester linked

MBP-DAC conjugate (Fig4-5). The covalent attachment of DAC to MBP was confirmed by

monitoring the labelling reaction with 3M using ESI-MS which showed a decrease in unlabelled

MBP (MW = 46,360 Da) with time and a corresponding increase in MBP labelled with one

molecule of DAC (found: 46,603 Da Fig4-6) with no indication of multiply labelled proteins

present. Isolation of MBP following incubation with the lactose construct 3L confirmed minimal

MBP labelling in comparison to that observed with construct 3M (Fig4-5). In the presence of

excess maltose (50 mM) no labelling of MBP was observed confirming the necessity of

recognition of construct 3M at the maltose binding site (Fig4-5).

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

350 400 450

Ab

sorb

ance

wavelength (nm)

T = 2hr

T = 4hr

T = 8hr

T = 12 hr

T = 16hr

T = 20hr

81

Figure 4-5. UV spectra of isolated DAC labelled MBP. MBP (5 µM) and thioester (10 µM), HEPES

buffer (10 mM, pH 8, 20 hr, 22 °C), 3M (▬), 3L (▬), or 3M + maltose (50 mM) (▬).

Unexpectedly, given the hydrolysis or nonspecific reaction observed by UV spectroscopy with

constructs 2M and 2L, the MBP isolated from labelling with these constructs showed similar

results to those observed with constructs 3M and 3L (Fig4-7). Constructs 2M and 2L gave

preferential labelling with maltose construct (2M) and labelling inhibition with free maltose was

again observed. As expected based on the absorbance studies no labelled MBP was isolated

when the alkyl thioesters 1M or 1L were used in the labelling reaction.

The yields of labelling MBP with constructs 2M and 3M were calculated by UV and the

selectivity of the labelling was assessed by comparing the labelling yields of reactions conducted

with the maltose and lactose constructs 2 and 3 (Fig4-8). Constructs 2M and 3M gave an average

MBP labelling yield of approximately 80% based on protein concentration and a selectivity of

13:1 when comparing to the isomeric lactose conjugate. Even greater selectivity was observed in

the presence of excess free maltose, suggesting some affinity for the lactose conjugate by MBP.

82

Figure 4-6. ESI-MS timecourse of MBP labelling reaction with 3M. Thioester (10 µM), MBP (5

µM), HEPES buffer (10 mM, pH 8, 22 °C). Labelled MBP indicated by asterix*

The point of attachment of the DAC fluorophore to MBP by reaction with construct 3M was

identified through analysis of a tryptic digest with ESI-MS/MS characterization. The HPLC

purification of the DAC-labelled-MPB tryptic digest performed gave one fraction with strong

UV absorbance at 428 nm (Fig4-9). ESI-MS/MS analysis of this fraction confirmed the covalent

addition of DAC to lysine 43 (Fig4-10). In the crystal structure of MBP this residue is found at

the periphery of the MBP binding cleft well positioned for reaction with the thioester constructs

(Fig4-11).

83

Figure 4-7. UV spectra of isolated DAC labelled MBP. MBP (5 µM) and thioester (10 µM), HEPES

buffer (10 mM, pH 8, 20 hr, 22 °C), 2M (▬), 2L (▬), or 2M + maltose (50 mM) (▬).

0

0.05

0.1

0.15

0.2

0.25

250 300 350 400 450 500

Ab

sorb

ance

wavelength (nm)

84

Figure 4-8. A chart of the tabulated MBP labeling yield and selectivity achieved using constructs

1M 2M and 3M. All reactions were done in triplicate and isolated using a PD-10 column as

described.

0

10

20

30

40

50

60

70

80

90

100

3M 1M 2M

Ave

rgae

lab

elli

ng

yie

ld (

con

cen

trat

ion

of

la

be

lled

MB

P/

tota

l MB

P)

conjugate

0

2

4

6

8

10

12

14

16

18

20

3M 1M 2M

Ave

rgae

Se

lect

ivit

y (r

atio

of

lab

elli

ng

yie

ld/l

acto

se-c

on

juga

te la

be

llin

g yi

eld

conjugate

85

Figure 4-9. C-18 HPLC chromatogram of DAC labelled MBP trypsin digest detected by dual UV

absorbance at 215nm and 428 nm. The major fraction indicated by a red arrow was subject to ESI-

MS/MS analysis to decipher the amino acid residue of DAC attachment.

(VTVEHPDKLEEK)-DACA

U

0.00

0.02

0.04

0.06

0.08

0.10

Minutes

10.00 15.00 20.00 25.00 30.00 35.00 40.00 45.00 50.00 55.00

AU

0.00

0.20

0.40

0.60

0.80

1.00

Minutes

10.00 15.00 20.00 25.00 30.00 35.00 40.00 45.00 50.00 55.00

428nm

215nm

DAC

M13+

(M-DAC)2+

B2

Y1

(Y10)2+

(Y9)2+

V T V E H P D K L E E K

DACM1 =1665.9

M-DAC

B2 Y1

Y10

Y9 Y7

Y7

86

Figure 4-10. Top: raw ESI-MS/MS spectrum of the single DAC labelled MBP tryptic peptide

collected by HPLC. Bottom: Deconvoluted ESI-MS/MS spectrum of multiply charged fragment

ions observed in the raw ESI-MS/MS spectrum.

M1

M1-DAC Y9

Y10

V T V E H P D K L E E K

DACM1 =1665.9

M1-DACY10

Y9

87

Figure 4-11. Structure of maltose bound MBP (PDB: 1MBD). Maltose (green (carbon), red

(oxygen) atoms), lys43 (yellow (carbon) ε-amine (blue)

MBP labelling in cellular extracts and in live cells

Given the specificity of the labelling reactions, the ability of the ligand-tethered aryl thioesters to

specifically label MBP in a complex protein mixture was evaluated. Labelling experiments with

3M and 3L were evaluated in crude bacterial cell extracts spiked with defined quantities of

MBP. Following incubation, the extracts were separated by SDS-PAGE and imaged for

fluorescence (Ex 330-385 nm Em 480-520 nm) A single labelled band corresponding to the

molecular weight of MBP was visible at concentrations of MBP as low as 1 µM (total MBP in

sample ~200 ng) with no observable labelling of other proteins in the extract (Fig4-12, lanes 4

and 5). Labeling with 3L at equivalent concentrations showed minimal labeling of MBP.

Comparing the fluorescence intensities of the labelled MBP bands with 3M and 3L (5 µM)

revealed a 17:1 selectivity in agreement with solution studies.

88

Given the selectivity of 3M for labeling in complex mixtures, the ability of 3M to label

intracellular MBP during expression in E. coli was evaluated. Incubation of 3M (10 µM) with E.

coli showed a significant degree of cell uptake of the construct as observed by confocal

microscopy and flow cytometry (Fig4-13 A-D,F, Fig4-14). Washing of induced and non-

induced cells with PBS/0.2% TWEEN following uptake of the labeling construct, cell fixation

and membrane permeablization showed partial removal of unreacted labeling construct in cells

not induced to express MBP. This suggested intracellular labeling of MBP in induced cells had

indeed taken place to some extent (Fig 4-15). SDS-PAGE analysis of the cellular extracts

following the intracellular labeling reactions confirmed that selective MBP labeling had taken

place using 3M with a single fluorescent band corresponding to labelled MBP (Fig 4-13E lane

1). A fluorescent band corresponding to labelled MBP was observed at intensities well above any

background labeling (Fig 4-13E lane 1 and 3) with the labeling also shown to be glycan specific,

as only weak labeling was observed with the lactose construct (3L)(Figure 4-13E lane 2).

Figure 4-12. SDS-PAGE analysis of MBP labeling in crude bacterial cell extracts. Left: Coomassie

blue stain, Right: Fluorescence image Ex 330-385 nm Em 480-520 nm (lane M: MW markers 1-6

crude cell extract with, 1: no MBP, 2: 100 nM MBP, 3: 500 nM MBP, 4: 1 µM MBP, 5: 5 µM MBP),

3M (10 µM) added to lanes 1-5, lane 6: MBP (5 µM), 3L (10 µM).

89

Figure 4-13. (A-D) Confocal microscopy of bacterial cells. (A) Bright field image of induced cells

without 3M. (B) Fluorescence imaging (Ex 408 nm, Em 460-500). (C) Bright field image of induced

cells with conjugate 3M (10 µM) for 3hr. (D) Fluorescence image of these induced cells using (Ex

408 nm, Em 460-500). (E) SDS-PAGE analysis of cellular extracts derived from E. coli. Lane M:

MW markers, 1: cells induced to express MBP incubated with 3M (10 µM, 3 hr), 2: cells induced to

express MBP incubated with 3L (10 µM, 3 hr), 3: non-induced cells incubated with 3M (10 µM, 3

hr), Left: Coomassie blue stain, Right: Fluorescence image (Ex 330-385 nm Em 480-520 nm). (F)

Flow cytometry analysis of E. coli. ▬ induced cells with 3M (10 µM, 3 hr). ▬ non-induced cells

with 3M (10 µM, 3 hr). (H) ▬ non-induced cells

E

F

A B

C D

Lane: M 1 2 3 1 2 3

170

130

705540

35

25

Construct: 3M 3L 3M 3M 3L 3MInduction: + + - + + -

C

90

Fig 4-14. Confocal images based on z-stacking analysis of an E.coli cell following incubation with

3M for 3hrs and PBS washing steps illustrating the cellular uptake of 3M. A) left: DAC

fluorescence image at z = 0.17 µm, right: brightfield image at z= 0.17 µm B) left: DAC fluorescence

image at z = 0.34 µm, right: brightfield image at z = 0.34 µm

91

Figure 4-15. Flow cytometry histograms generated from fixed bacterial cells following incubation

with 3M. Live bacterial cells were incubated with 10uM 3M for 3hr followed by PBS washing,

formaldehyde crosslinking/fixation, and washing (0.2% TWEEN-20, PBS) to remove unreacted 3M.

A. Cells with induced MBP expression. B. Cells not induced to express MBP. Cells overexpressing

MBP (A) retain more fluorescence following fixation and washing. In the absence of fixation and

washing steps, induced and non-induced bacterial cells both take up the labelling constructs with

the same efficiency according to flow cytometry as shown in fig 4-13f.

4.3 Concluding Remarks

In conclusion, we have developed a novel method to achieve the selective and high yielding

fluorescent labelling of the sugar binding protein, MBP, using ligand directed thioester based

acylation. Labelling was observed in real time using UV spectroscopy which reported on the acyl

transfer of DAC from the protein bound thioester constructs to a lysine residue proximal to the

maltose binding site (Scheme 4-2). Selective labelling was observed to occur in cell extracts in

the presence of an abundance of other proteins demonstrating the potential application of these

constructs for imaging applications in the biological milieu or in lectin profiling experiments.

Given the wide use of MBP as a protein fusion construct this approach may find use as a

A

B

92

selective method for recombinant protein labelling. The synthetic ease of generating the

carbohydrate thioesters provides the opportunity to produce labelling agents from small

quantities or even isolated carbohydrates to target other carbohydrate binding proteins in a time

efficient manner. The use of a single labelling construct simplifies the labelling reaction in

complex mixtures or in intracellular environments.

Scheme 4-2. The labelling of MBP using ligand directed thioester conjugates.

4.4 Materials and Methods

Scheme 4-3. Synthesis of glycosyl hydrazide thiols

Maltose bindingprotein

Site selectivelabelling

Thioesterrecognition

93

Synthesis of S1M and S1L:

3-Mercaptopropionic acid hydrazide (Fig A.1) (0.024g, 0.2mmol) and maltose or lactose (0.2 g,

0.6 mmol) were added to deionized water (1 ml) containing glacial acetic acid (20 mM) and the

resulting suspension was incubated at 37 oC for 24 h. Following the reaction, the product was

purified using C-18 HPLC with an ammonium acetate buffer (50 mM, pH 7): acetonitrile

gradient and monitored by UV at 215 nm. The desired product peak was collected and

lyophilized giving rise to a colourless solid corresponding to the conjugate or S1L (FigA.2, Fig

A.3) or S1M (FigA.4, FigA.5). After 3 successive lyophillization steps to remove residual

ammonium acetate the product was obtained in approximately 50 % yield (0.04-0.05 g).

Synthesis of S2M and S2L

p-Mercaptobenzohydrazide:

4-mercaptobenzoic acid (0.5g, 3mmol) was dissolved in methanol (50 ml) and left stirring on ice

until completely dissolved. Acetyl chloride (5 ml) was subsequently added drop wise and the

reaction was brought to room temperature and left stirring for 8 hrs. The mixture was

concentrated dissolved in ethyl acetate and extracted three times with a solution of sodium

bicarbonate (5 %). The organic layers were combined and concentrated under reduced pressure

to obtain the 4-mercaptobenzoate methyl ester product as a yellow-green powder in 90-95 %

yield (0.46g, 2.76 mmol, FigA.8)

Methyl 4-mercaptobenzoate (0.2 g, 1.2 mmol) was dissolved in dry methanol (5 ml) and stirred

at 50 oC until fully dissolved. Excess hydrazine hydrate (2 ml) was subsequently added and the

reaction was stirred at room temperature for 12 hr. The solvent was removed under diminished

pressure and the product was precipitated out of cold ethanol. The product was collected by

vacuum filtration and washed with cold ethanol to obtain 4-mercaptobenzohydrazide as a white

solid. Yield 88% (0.18 g, 1.06 mmol, (FigA.9))

4-mercaptobenzohydrazide (0.010 g, 0.06 mmol) and maltose or lactose (0.2 g, 0.6 mmol) were

added to deionized water (1 ml) containing glacial acetic acid (30 mM) and the resulting

94

suspension was incubated at 37oC for 24 hr. The suspension was centrifuged to remove the

undissolved starting material and the supernatant was purified by semiprep C-18 HPLC using an

ammonium acetate (50 mM pH 7 : acetonitrile gradient and monitoring by UV at 310 nm. The

product was collected and lyophilized giving rise to an amorphous solid corresponding to the

product S2L (FigA.10, FigA.11) or S2M (FigA.12, FigA.13). Weighing the product following 3

successive lyophilization steps resulted in a 80-85% yield (approximately 0.018g, 0.036 mmol)

Synthesis of S3M and S3L

4-Mercaptophenylacetic acid hydrazide:

4-mercaptophenylacetic acid (0.5 g, 2.75 mmol) was esterified as described above with 4-

mercaptobenzoic acid, to yield the methyl ester in 88 % yield (0.44 g 2.42 mmol FigA.16) The

methyl ester (0.2 g, 1.1 mmol) was subjected to hydrazinolysis as described above and the

mercaptophenylacetic acid hydrazide was obtained in 92% yield (0.183 g ,1.01 mmol FigA.17).

4-Mercaptophenylacetic acid hydrazide (0.010 g, 0.055 mmol) and maltose or lactose (0.2 g, 0.6

mmol) were added to deionized water (1 ml) containing glacial acetic acid (25 mM) and the

resulting suspension was incubated at 37oC for 24 hr. After 24 hours, the suspension was

centrifuged to remove undissolved starting material and the supernatant was purified by

semiprep C-18 HPLC using an ammonium acetate (50 mM pH 7): acetonitrile gradient

monitoring by UV at 270 nm. The product was collected and lyophilized giving rise to an

amorphous solid corresponding to S3L (FigA.18, FigA.19) and S3M (FigA.20, FigA.21).

Weighing the product following three successive lyophillizations allowed for an estimated yield

(62-76 %).

95

Scheme 4-4. Synthesis of glycosyl thioester conjugates

Synthesis of 1M and 1L

The maltose and lactose linked thiols S1M and S1L (~0.0025g, 0.0056 mmol) and 7-

diethylaminocoumarin thioglycolic acid thioester (2 mg, 0.0056 mmol) were dissolved in HEPES

(0.5 ml, 100 mM, pH 8) buffer containing 5mM tris(2-carboxyethyl)phosphine (TCEP) prepared

from a TCEP (0.5 M, pH 7) solution. The reaction was incubated for 4 h at room temperature,

diluted 10 x with HEPES (100 mM, pH 8) buffer and purified by C-18 HPLC with an

ammonium acetate (50 mM, pH 7): acetonitrile gradient monitoring absorbance 430 nm.

Collection and lyophilization of the desired peak gave an orange solid ~ 40 % yield (0.0022-

0.0025 mmol by UV) for 1L (FigA.6) / 1M (FigA.7).

Synthesis of 2M and 2L

Compound S2M or S2L (0.018 g, 0.036 mmol) and DAC thioglycolic acid thioester (2 mg, 5.66

µmol) were dissolved in HEPES (0.5 ml, 100 mM, pH 8) buffer containing TCEP (36 mM)

prepared from a TCEP (0.5 M, pH 7) solution. The reaction was incubated for 4 h at room

temperature, diluted 10 x with HEPES (100 mM pH 8.0) buffer and purified by C-18 HPLC

using a semiprep column with an ammonium acetate (50 mM, pH 7): acetonitrile gradient

monitoring absorbance at 430 nm. Collection and lyophilization of the desired peak gave rise to

an orange fluffy solid ~ 8-10 % yield for 2L (Fig A.14) / 2M (Fig A.15) (0.57 µmol by UV)

96

Synthesis of 3M and 3L

S3M/S3L (0.0044g, 0.008 mmol) and DAC-thioglycolic acid thioester (2 mg, 5.66 µmol) were

dissolved in HEPES (0.5 ml, 100 mM, pH 8) buffer containing TCEP (8mM) prepared from a

TCEP (0.5 M, pH 7) solution. The reaction was incubated for 4 h at room temperature, diluted

10 x with HEPES (100 mM pH 8) buffer and purified by C-18 HPLC using a semiprep column

with an ammonium acetate (50 mM, pH 7): acetonitrile gradient monitoring absorbance at 430

nm. Collection and lyophilization of the desired peak gave an orange fluffy solid ~ 15-18 %

yield for 3L (FigA.22) / 3M (FigA.23) (1.02-1.08 µmol by UV).

Characterization of MBP labeling in vitro

a) Preparation of Cell Culture, protein expression and purification

Maltose binding protein was expressed using a pMAL-c4X MBP protein fusion vector (6645 bp)

that was purchased from New England BioLabs Inc. The fusion vector (2 ng) was transformed

into BL21(DE3) competent cells and plated onto agar plates containing 100 μg/mL ampicilin. On

the following day, a single colony was used to inoculate Luria–Bertani (LB) broth (25 mL) that

had been supplemented with ampicilin (100 μg/mL). The 25 ml overnight culture was used to

inoculate 1 L of LB broth supplemented with 100 μg/mL ampicilin. Cells were grown at 37 °C

and induced with IPTG (0.3 mM) when an OD600 of 0.5 had been reached. The cells were kept at

37 °C and grown for a further 2 hrs before centrifugation to separate the media from the protein-

containing cell pellet. The pellet was resuspended in column buffer containing Tris-

HCl (20 mM, pH 7.5), sodium chloride (200 mM), and ethylenediaminetetraacetic acid (1 mM)

1 tablet of protease inhibitor cocktail and frozen at − 20 °C until purification. To obtain live cells

for the labelling, aliquots of cells were taken 1h after IPTG induction. The resuspended cell

pellet was sonicated in pulses on ice for 5 min and then centrifuged at 4000g for 1 hr to separate

the supernatant from the pellet. Aliquots of these cell extracts were used for the labelling

(described below). The protein was purified on an amylose resin column equilibrated with the

column buffer. The sample was loaded onto the column and the resin was washed with 3 column

volumes (CV) of the column buffer. The resin was subsequently washed with 2 CV of high-salt

buffer (i.e., column buffer supplemented with 1 M NaCl), followed by a further 3 CV of column

97

buffer. The protein was eluted with maltose (10 mM) in HEPES buffer (50mM, pH 8.0). The

eluted protein was dialyzed extensively against Tris–OAc (40 mM, pH 7.5) with

NaCl (100 mM ). The dialyzed protein was concentrated to ∼ 1.5 mL using an Amicon

ultracentrifugal device (nominal molecular weight limit, 10,000 ; Millipore) and subject to FPLC

purification using a MonoQ 10/100 GL (GE Healthcare) column equilibrated with Tris (20 mM

pH 7.5) and eluted with a NaCl gradient. The protein eluted at ~150 mM NaCl and the purity of

the samples was confirmed using 12.5% SDS-PAGE and electrospray ionization mass

spectrometry.

b) UV and ESI-MS analysis of MBP labeling reactions

Stock solutions of MBP were prepared at a concentration of 500 µM by UV (A280 ε 86,360) in

HEPES (10 mM, pH 8) and stored at 4 0C. Stock solutions of the thioester conjugates were

prepared at a concentration of 40 µM by UV (A455 ε 56,000) in HEPES (10 mM, pH 8, 10%

DMSO) and stored at -80 0C. MBP labeling reactions were initiated by adding MBP and the

desired conjugate thioester from pre-prepared stock solutions into a solution of HEPES (10 mM,

pH 8) affording a final MBP concentration of 5 µM and thioester conjugate concentration of 10

µM in 1 mL of solution. The reactions were monitored in a sealed quartz cuvette (150 µl) by

absorbance in spectrum acquisition mode with a temperature controlled sample chamber

automatically acquiring full spectrum with 1 nm increments at 1 hr intervals. Following a 20 hr

reaction time, the protein/conjugate solutions were immediately added to a PD-10 size column

pre-equilibrated with HEPES buffer (10 mM, pH 8) and MBP was eluted with the equilibration

buffer in the first 4 ml. The protein is visible to the naked eye when labeling is carried out with

constructs 3M and 2M. A clear orange band corresponding to unreacted conjugate is clearly

retained near the top of the column. Ethanol 20%/water washes are required to remove the

unreacted construct from the column and regenerate the PD-10 for further use. The MBP fraction

was lyophilized or concentrated using 3K spin filters and analyzed by UV in full spectrum

acquisition mode following buffer subtraction scanning from 250-600 nm. The ratio of A280/A428

(MBP ε = 86,360, DAC ε = 52,000) was used to calculate the yield of labeled MBP and the

determination of the extent of selective labeling occurring as described above. Solutions of DAC

labeled MBP were than subjected to modification site analysis. All absorption spectra were

measured on a Schimadzu UV-2401PC spectrophotometer at T = 296 K.

98

The labeling reaction was also monitored by ESI‐MS where 100 μl aliquots of the reaction

mixture (5 μM MBP, 10 μM 3M , 10 mM HEPES pH 8) were diluted 10 times in 50 mM

ammonium acetate pH 6 and subjected to ESI‐MS analysis.

c) Determination of the modification site on MBP

DAC labeled MBP (189 μg) was dissolved in a 90 μl solution of 2M UREA/50 mM Tris buffer

pH 7.5 and 10 μl of a trypsin stock solution (1 μg/μl in 50 mM acetic acid stored at ‐80oC) was

added and the solution was incubated at 37 °C for 24 h. The trypsin digest was purified by

analytical C‐18 HPLC using an H20 (0.1% TFA): acetonitrile (0.1% TFA ) gradient monitoring

by dual absorbance detection at 215 nm and 428 nm. The DAC containing fraction was collected

and desalted using a 3k spin filter and subject to ESI‐MS/MS analysis.

Characterization of MBP labeling in E. coli

a) Labeling reaction in bacterial cell extracts

E. coli BL21(DE3) pMAL-c4X extracts were prepared as described above, but excluding the

IPTG induction step. To remove MBP that may result from leaky plasmid expression, the extract

was passed through an amylose column as described above. The extracts were than dialyzed

against HEPES (10 mM, pH 8) for 2 h using millipore dialysis membrane with a 3.5 kDa

molecular weight cutoff. Labeling reactions were carried out in 50 µl of cell extract with known

amounts of purified MBP titrated into the extract in the presence or absence of thioester

conjugate 3M or 3L (10 µM) for a period of 20 hrs. The samples were then boiled for 5 min in

SDS denaturing buffer (10 µl) (6x Tris HCl/SDS buffer pH 6.8, 3 ml glycerol, 1g SDS, 0.93g

DTT) and analyzed by 12 % PAGE. The resulting gels were first imaged for DAC fluorescence

(Ex 330-385 nm Em 480-520 nm) and then stained with coomassie blue.

b) Labeling reaction in E. coli

The labeling reactions on live cells were conducted using MBP induced (1 h post-induction) and

non-induced cells immediately following cell growth with an aliquot (1 ml) of the culture having

99

an OD600 = 1.2. The cells collected by centrifugation for 5 min following which the supernatant

containing the media was removed and replaced with 290 ul of HEPES (pH 8, 10 mM)

containing 10 µM of the desired thioester conjugate. The cells were resuspended and the labeling

reactions were incubated at 37 oC for 3 h. The cells were collected by centrifugation and washing

with PBS to remove thioester construct not taken up by the cells. The cells were than prepared

for imaging by confocal microscopy, flow cytometry analysis, or were lysed and the cellular

extract was analyzed by SDS-PAGE as described above. The cells were lysed in SDS denaturing

buffer at 100oC for 5 min and then sonication for 5min. The suspension was clarified by

centrifugation to remove insoluble cellular components prior to SDS-PAGE analysis.

c) Confocal Microscopy of Live Cells Incubated with Thioester Conjugates

Samples of live bacteria cells with a density of ~1 x 108 cells/ml (OD = 1 at 600 nm) were plated

on 15 mm cover slips (Fischer) coated with poly‐L‐lysine for 20 min and were examined with a

Leica TCS SP5 confocal microscope using a 405 nm diode laser with a DAPI filter (460‐500nm)

and differential interference contrast (DIC) configuration. Z‐stacked plots were also generated

from a single bacterial cell of approximately 1 x 3.16 μm dimensions containing DAC

fluorescence. The images were acquired with a 0.17 μm step size thickness, scan speed of 40Hz

and a 67.9 um pinhole.

d) Flow Cytometry of Live Cells Incubated with Thioester Conjugates

Analysis of cells following labeling experiments by flow cytometry was carried out on a

DakoCytomation MoFlo cell sorter (Fort Collins, CO) equipped with a 407 nm laser. Solutions

of live cells (300 ul) in PBS were analyzed at a cell density of ~ 1 x 108

cells/ml. Analysis of the

forward and side scatter suggested that the cells were still intact. Between 20,000 and 50,000

events were gathered in each experiment. The data was analyzed using Summit Software

(DakoCytomation, Ft. Collins, CO). Attempts to wash out free unreacted conjugate from live

cells following labelling were carried out by pelleting the cells following 3hr incubation with 3M

and washing the cells with PBS to remove extracellular unreacted conjugate. The cells were then

fixed in a 3.7% formaldehyde/water solution for 30 min rt and washed with 0.2% TWEEN

20/PBS to remove intracellular unreacted free conjugate. The cells were then resuspended in

100

PBS and analyzed by flow cytometry. Completely removing free unreacted intracellular

conjugate is difficult due to the hydrophobicity of the DAC and will be the focus of future

efforts.

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Chapter 5 Summary and Perspective

Complex carbohydrates and their diverse structural and regulatory roles have historically

received less attention than the more popular amino acid and nucleic acid based biomolecules.

Proteins and DNA are directly linked through transcription and translation where the complex

carbohydrates of the cell are not genetically encoded, often structurally complex, and are

produced in many instances by poorly understood processes involving an ensemble of enzymes.

The difficulties associated with studying these complex carbohydrates in comparison to

biomolecules such as proteins has further limited our understanding of this class of biomolecules.

Proteins crystallize more easily than carbohydrates allowing for detailed structural analysis by X-

ray crystallography. Proteins generally ionize more efficiently by ESI than carbohydrates due to

the wealth of basic residues while their linear polymeric backbone simplifies analysis by tandem

mass spectrometry compared to complex carbohydrates which are often branched.

Analysis of the structure of GAGs is especially challenging due to the difficulty obtaining

homogenous material necessary for NMR and crystallographic analysis. Analysis can be

simplified by using synthetically pure heparin/HS which has only recently been achieved using

chemoenzymatic approaches (1) or by using oligosaccharides to serve as model structures (2)

Heparin derived oligosaccharides may be prepared synthetically or by enzymatic degradation of

full length heparin. The multi-functionality of GAG oligosaccharides makes organic synthesis

extremely difficult, low yielding and time consuming due to the orthogonal protecting group

chemistry required. Enzymatic synthesis of HS/heparin oligosaccharides is also non-trivial

requiring careful control of the enzymatic steps to avoid heterogeneity in the oligosaccharide of

interest.

Analysis of GAGs by mass spectrometry is also challenging in part due to the high degree of salt

adduct formation common to GAGs in addition to the labile sulfate and carboxylate groups

present on GAG chains such as heparin and HS. As a result, ion-pairing reagents are often

employed to reduce salt adducts which simplify the spectrum, and key mass spectrometric

parameters must be carefully optimized to reduce the loss of small neutral molecules and chain

fragmentation in the gas phase. In the case of ESI-MS, which is often used to analyze GAG

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structure, ion optics, and voltage gradients must be carefully tuned to lower the collisional

energy of GAG ions in the gas phase (3).

The ability to analyze GAGs in complex biological settings presents further challenges. It is

generally difficult to generate antibodies against glycosaminoglycans and only recently have a

limited number of phage display antibodies become available. Unfortunately the binding

specificity of these phage display antibodies are poorly defined. Through further understanding

of the binding interactions between peptides and proteins with GAGs it may be possible to

develop fluorescent probes selective for GAGs, capable of operating in a biologically relevant

context. These tools would allow the details of GAG expression patterns, degradation and

protein binding to be understood in a cellular context.

5.1 The importance of the spatial display of charge in peptide/protein interactions with

GAGs

The synthesis of quinolinium-tethered peptide-based probes for heparin was presented in chapter

2. These constructs were applied to gain structural information regarding peptide-heparin

interactions. Peptides with positive charge presented on one face of an α-helix (Peptide 1) in a

continuous linear distribution bound to heparin with the highest affinity. Peptide 1 with this

arrangement of lysines also experienced a significant increase in helical content upon binding

heparin. Taken together, these results strongly suggest that heparin prefers to bind to helical

cationic peptides in an extended conformation. These findings could lead to the design of higher

affinity and more selective heparin probes. Current heparin binding peptides generally contain

cationic functional groups to promote electrostatic contacts with heparin but little attention has

been given to optimizing the spatial display of these cationic groups. Recently it was reported

that radiolabelled peptide probes could selectively bind amyloid laden organ tissue in live mice

via binding interactions with amyloid protein associated HS (4). One probe in particular (p5)

bound to the amyloid laden liver and spleen with the highest avidity of all the peptides and could

be imaged in diseased organ tissue for the longest period of time. Interestingly, this peptide had

an identical display of lysine residues to peptide 1 used in our investigations. The authors

referenced our findings and proposed the high avidity binding of p5 to amyloid associated HS

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was due to the spatial display of positive charge by this peptide upon taking up an α-helical

conformation.

In addition to having an impact on HS probe design, our findings help reveal how heparin/HS

structure is related to biological function. Ternary complexes involving thrombin, AT III and

heparin, which have been observed to be important intermediates in the inhibition of thrombin,

have been studied using X-ray crystallography with a heparin mimetic (5). These studies suggest

that heparin is in an extended conformation acting as scaffold on to which both proteins can bind

and engage in protein-protein interactions. Binding to heparin in its preferred extended

conformation as was observed in our peptide-heparin binding studies likely allows for high

affinity interactions to take place between ATIII, thrombin, and heparin favoring ternary

complex formation. The optimal spatial display of charge found in the peptide studies may also

serve as a predictive factor for a protein’s ability to interact with HS.

Another important implication of our results on HS biology pertains to the mechanism of cell

penetrating peptide cell uptake. It was shown through the design of small peptide based

foldamers that the rigidity and display of charge may serve as important parameters for the

efficiency and mechanism of CPP uptake (6). These authors observed that one peptide (1b)

which displayed positive charge clustered on one face of the helix and possessed the highest

degree of 14-helical structure, entered cells to the greatest extent and with the fastest kinetics

when compared to peptides lacking helicity or displaying charge along the entire periphery of the

helix (2b-5b). It was also shown that peptide 1b entered predominantly through energy

dependant endocytic mechanisms. Our results suggest a likely explanation for these observations

as peptide 1b with a high degree of helicity and linear display of charge likely binds to cell

surface HS with the highest affinity when compared to the other peptides with HS subsequently

facilitating endosomal uptake, a well known biological role of HS.

5.2 Peptide-Cluster formation involving cell penetrating peptides

The debated mechanisms of cell uptake of CPPs and their dependence on cell surface GAGs such

as HS and chondroitin sulfate were addressed in chapter 3. These studies describe a rapid, high

sensitivity and in real time method for the investigation of the ability of heparin to differentially

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cluster CPPs. It was found that clustering to the GAG heparin, serving as a model of cell surface

HS, was largely dependent on the peptide amino acid composition and independent of its

apparent binding affinity to a single heparin chain. Peptides, similar in their dissociation

constants for binding heparin and in their number of positive charges showed significant

differences in their ability to form multimeric clusters with heparin. Using fluorescent

aggregation probes to distinguish clustering from single peptide-single heparin complexation, it

was observed that clustering between the well known CPP Antp and heparin was much more

stable than heparin clustering involving the TAT peptide. These findings not only confirm the

existence of this interesting binding property of HS but also help to explain the different degrees

of GAG dependence and mechanism of uptake preferred by Antp and the TAT CPPs, making an

important contribution to the field of CPP-cargo and drug delivery technology.

Gellman et al. conducted experiments on the ability of guanidinium β peptides to enter HeLa

cells for applications as delivery vectors. The peptides were observed to enter via

macropinocytosis and induce actin reorganization which was dependent on both the cell surface

HS and on peptide structure (7). The authors proposed that specific interactions between

oligocations and HS proteoglycans may occur resulting in the propagation of a signal triggering

actin reorganization. The results of our studies on CPP-heparin clustering suggest that this

specific interaction likely involves clustering of multiple oligocations to extracellular HS

proteoglycan with the structure of the peptide or oligocation likely effecting the stability of the

resulting multimeric peptide-HS cluster. This in turn would have a direct impact on the ability of

the peptide, potentially with the cargo of interest attached, to enter the cell via endocytosis. The

stability of these clusters likely governs the degree to which and efficiency with which the CPP

is taken up via endocytotic energy dependant mechanisms however how the resulting cluster

promotes endocytosis is not entirely understood.

The occurrence of CPP-heparin clustering has been reported recently to occur with polyarginine

peptides using dynamic light scattering and interestingly the mechanism of cell uptake changed

from endocytosis to translocation as the concentration of peptide increased (8). Clustering

stability and dynamics were not assessed and were observed under concentrations significantly

higher than those used in CPP uptake studies. Our method of characterizing peptide-heparin

clusters may shed light into the origins of this mechanistic switch. It is highly possible that the

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change in mechanism of uptake at high peptide:heparin ratios occurs due to an increase in the

degree of peptide-HS clustering at the cell surface. As observed in our studies, heparin:CPP

cluster formation is driven largely by peptide-peptide interactions and increases with peptide

concentration. The polyarginine peptides studied may enter the cell predominantly via

translocation upon clustering cell surface HS in contrast to what has been observed with the TAT

peptide. Our aggregation probes could study the stability of clusters involving heparin and this

polyarginine peptide at concentrations relevant to CPP uptake studies. This could allow for

correlations to be made between cluster stability and the observed mechanism of uptake

preferred by this peptide.

Another important contribution of our work geared towards the characterization of peptide-

heparin clustering is the ability to distinguish monomeric peptide heparin complex formation

from clustering and quantitatively characterizing the stability of peptide-heparin clusters. The

existence of clustering brings into question the accuracy of reported dissociation constants in the

literature for peptide-heparin interactions as the affinities of these interactions are often

determined using ITC or surface plasmon resonance. The higher concentrations of peptide used

in ITC analysis and the assumption of 1:1 binding and equivalent binding sites on heparin for the

peptide measured, results in a calculated binding affinity that is likely obscured from

contributions due to clustering. In SPR the immobilization of heparin or peptide likely enhances

clustering and likely non-native aggregates again resulting in flawed dissociation constants being

calculated for monomeric peptide-heparin binding.

The ability of cell surface HS to form stable clusters with cell penetrating peptides, confirmed by

our aggregation assays, may be an important factor underlying the mechanism of action of cell

penetrating peptides, viral peptides, and various toxins such as those isolated from certain snake

venoms. Additionally, the ability of HS to cluster peptides may be applicable to the HS

dependant clustering of proteins on the cell surface known to play a role in growth factor

receptor clustering which is a crucial step in signal transduction and cell growth and

proliferation.

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5.3 The synthesis of thioester carbohydrate conjugates as selective lectin labelling agents

In efforts to develop a selective labeling strategy for carbohydrate binding proteins, thioester

carbohydrate conjugates were designed to target maltose binding protein (MBP) as a model

lectin. In chapter 4, the high yielding and selective labeling of MBP with diethylaminocoumarin

was presented and it was shown that MBP was site selectively labeled at a lysine proximal to the

maltose binding site. It was also shown that selective labeling could be carried out in bacterial

cell extracts with an unoptimized limit of detection in the range of 1 µM MBP. Intracellular

MBP over-expressed in live bacterial cells could also be labeled in as quickly as three hours.

This serves as a potential way to efficiently identify and/or isolate unknown carbohydrate

binding proteins or profile the expression levels of known carbohydrate binding proteins in

cellular extracts derived from various sources. In addition, this strategy also offers a potentially

general way to target intracellular proteins in living cells allowing for the imaging of

biomolecules.

Numerous protein and peptide labeling strategies exist in the literature each with their own

particular advantages and limitations. These strategies can largely be categorized into two

groups, small molecule approaches and chemical genetic approaches. An effective small

molecule approach has been demonstrated recently through the selective labeling of proteins

including carbohydrate binding proteins using affinity labeling techniques employing the use of

DMAP catalysis (9,10). However, this method is limited to targeting extracellular targets, is

synthetically demanding, and likely requires high binding affinity targets. Binding to high

affinity targets maximizes the time the activated acylating intermediate spends proximal to a

nucleophillic residue on the protein vs free in solution where it is highly susceptible to

hydrolysis. The affinity based thioester constructs described in chapter 4 represent an alternative

small molecule labeling approach and were shown to selectively and rapidly label an intracellular

carbohydrate binding protein at low µM concentrations. Our constructs are also stable for long

periods of time due to the inherent stability of thioesters and hydrazones to hydrolysis.

Another small molecule affinity-based labeling approach has been developed to specifically

probe serine hydrolases using a variety of small molecule acylating agents including

fluorophosphonates (FP), carbamates, and triazole urea derivatives (11). These acylating agents

109

have also been shown to have negligible cross reactivity with other nucleophillic enzymes such

as cysteine hydrolases. FPs are highly reactive providing complete coverage of mammalian

serine hydrolases and have found applications in the activity based protein profiling of serine

hydrolases (12,13). Their reactivity limits their application as selective probes for individual

serine hydrolases however. 1,2,3-triazole urea derivatives have recently been shown to be the

most effective serine hydrolase inhibitors to date with sub nanomolar activity in cells and high

potency in mice (11). These inhibitors have a high selectivity for peptidase and lipases and were

used to characterize the peptidase APEH and identify unknown N-acetylated protein substrates

for APEH in T-cells. Inhibition of APEH was accompanied by an accumulation of certain N-

acetylated protein substrates and an increase in T-cell proliferation (11). One potential

application of our protein acylation strategy is the profiling of unknown carbohydrate substrates

for known carbohydrate binding proteins. Site specific labeling proximal to the sugar binding

site of the protein with a carbohydrate responsive fluorophore results in the generation of a

protein-based biosensor capable of giving off a fluorescence response upon recognition of its

carbohydrate substrates. This could provide a direct, sensitive, and reversible method for

characterizing unknown substrates for a ligand binding protein. This is more reliable than the

indirect characterization of unknown protein binding substrates that accumulate following the

suicide inhibition of a ligand binding protein or enzyme. Potential protein substrates could

accumulate due to the non-selective inhibition of a protein other than or in addition to the desired

protein in the complex cellular environment resulting in the generation of false positives.

The affinity based thioester constructs discussed in chapter 4 may also be used to generate MBP

protein fusion tags allowing for the simultaneous imaging, isolation, and purification of

intracellular fusion proteins. Protein fusion tags may be generated by genetically fusing the

protein of interest to a genetically encoded fluorophore (GFP fusions) or by genetically fusing an

enzyme capable of self labeling to the protein of interest (Snap tags, Halo Tags). The Snap tag

involves fusion to O6-alkylguanine-DNA alkyltransferases (14) while the Halo tags (15) involve

fusion to the de-halogenase enzyme. However, GFP fusions, Snap tags, and Halo tags do not

allow for the simultaneous fluorescent labeling and purification of a fusion protein in a single

step, an advantage conferred by using MBP which can easily be purified using an amylose

affinity column.

110

Chemical genetic approaches such as the Snap tag and Halo tag described above afford selective

protein labeling through genetic engineering techniques. These genetic engineering steps often

involve the fusion of a peptide/protein tag to, or the binding site modification of, the protein of

interest. This allows for subsequent bioorthogonal labeling using small synthetic acylating

agents or selective non-covalent binding interactions with a fluorescent ligand.

A widely used and general chemical genetics approach to afford selective protein labeling has

been successfully achieved using the FlaSH reagent (16). This involves the attachment of a

peptide tag to the protein of interest which in turn is capable of selectively binding to the FlaSH

reagent. This method has been recently employed as a novel aggregation probe used to

mechanistically study amyloidogenesis (17). The FLaSH approach employs reversible thiol-

arsenic chemistry and requires the incorporation of a small FlaSH binding motif CCXXCC (C=

cysteine) on the peptide or protein target. This strategy however, requires high concentrations of

reducing agent, and potentially suffers from toxicity issues from nonspecific interactions with

arsenic complicating analysis in thick tissue samples and in vivo. Background labeling is also

problematic due to the number of naturally occurring cysteine containing motifs present in

biological proteins and the hydrophobicity of the FLaSH reagent. The maltose containing

conjugates we developed present advantages to the FLaSH method as it requires no additives, is

suitable for in vivo applications and has been observed to involve little background labeling

imparted by the affinity of the construct for only proteins that bind the ligand tethered to the

construct.

A novel chemical genetics approach targeting kinases has been developed and involves the

engineering of the kinase binding site to afford selective labeling (18). This has allowed for the

study of EGFR stimulation by EGF and its downstream outputs Akt, Erk1, and Erk2. Kinases

such as EGFR and Src-1 were engineered to contain an electrophile targeting cysteine in addition

to a glycine gate keeper residue which could allow for the labeling agent (6-acrylamido-4-

anilinoquinazoline tethered to an NBD fluorophore) to bind selectively to the target protein

followed by an irreversible reaction between the cysteine thiol on the protein and the Michael

acceptor on the probe. These probes were also observed to selectively label the engineered

kinases in the presence of the native form of the kinase in cells. This represents an effective

strategy to selectively target specific members within a related class of proteins and/or enzymes.

111

Our approach is limited with respect to such applications as all related members of a class of

proteins such as kinases and certain proteases would likely interact with our constructs due to a

high degree of ligand binding site homology.

A popular chemical genetic approach to label extracellular proteins requiring the fusion of a

peptide tag to the protein of interest employs the biotin ligase enzyme. Highly selective labeling

of the protein of interest is achieved by genetically fusing a biotin acceptor peptide tag to the

protein (19). However, this method requires the synthetically challenging chemical modification

of the biotin for imaging applications, tolerance of the modified biotin by biotin ligase, and the

addition of ATP along with time costly genetic engineering steps. The small molecule method

using affinity based-thioester constructs described in chapter 4 is a potentially rapid and efficient

alternative to such chemical genetic approaches and can potentially be applied to target and label

any ligand binding protein extracellularly and intracellularly in vivo or in cellular extracts. The

protein of interest can be targeted by modifying the acylating agent to contain the native ligand

for the protein of interest avoiding the time demanding genetic engineering of protein ligand

binding sites and the fusion of peptide tags. However, although highly selective for MBP and

site selective for a nucleophillic amino acid proximal to the sugar binding site, the ability to label

site specifically is dependent on the residues available on the target protein close to the ligand

binding site and cannot be specifically targeted as is the case in techniques such as native

chemical ligation (20) and unnatural amino acid incorporation (21).

5.4 Future directions

Investigations into the structural requirements for high affinity interactions between

peptides/proteins and the complex carbohydrates heparin and HS presented in this thesis can

likely serve as a foundation for generating new probes for HS that can function in the presence of

biologically relevant proteins present on the cell surface. Towards this end, an extension of our

peptide based heparin probe design efforts could involve the incorporation of β and cyclically

constrained amino acids which would allow for the generation of α/β helical peptides with

positive charge placed in optimal arrangements for specific interactions with HS while

conferring resistance to proteases. The incorporation of these un-natural amino acids could

allow for enhanced control over the positioning of cationic functional groups while maintaining

112

helical structure potentially binding HS with higher affinity. Accomplishing higher affinity

binding with a lower overall degree of positive charge would increase the selectivity of these

probes for HS over other proteins present in a mixture or on the cell surface.

The distribution of sulfates in HS into domains, in contrast to the more homogeneous distribution

found in heparin, could potentially be targeted to increase probe selectivity for HS. This could

be accomplished by placing positive charge in clusters on one face of the helical α/β peptide

spaced at intervals complementary to the distribution of sulfates on HS.

The immobilization of these HS binding peptides with a defined spacing of positive charge onto

sepharose beads could also be utilized as a purification technique for obtaining homogeneous

HS. The immobilized peptides could be designed to contain acidic amino acid regions that flank

the regions of positive charge ensuring that HS chains that do not contain the complementary

sulfation pattern bind with lower affinity and are eluted. Attention would need to be given to

ensure that the immobilized peptides are long and rigid enough to accommodate binding to HS

chains in their extended conformation. Rigidity of the immobilized peptides could possibly be

accomplished by a peptide stapling technique (22). Purified homogeneous HS may then be

subjected to fluorescent probe titrations with rationally designed probes to determine the binding

affinity before being applied to sensing HS in biological samples.

Our ability to probe reversible heparin/HS aggregation with CPPs, and characterize aggregate

stability may be applicable to studying the role of HS in amyloid aggregation. All amyloid is

associated with differentially expressed HS which is often oversulfated as observed with the HS

involved in primary amyloidosis (AL) (23). An aggregation probe for the dependence of these

processes on HS has not been developed. Appending HS or the amyloidogenic peptide/protein

with FRET pairs at the reducing terminus of HS or the N terminus of the peptide may allow for a

more rigorous study of the aggregation mechanism in vitro and represent a more general

approach to determining the role of HS structure on amyloidogenesis.

The work presented in this thesis also involved the development of a strategy to selectively

acylate lectins with the potential to generate selective protein based biosensors for complex

carbohydrates. Our strategy can be further developed to acylate an HS binding protein such as

113

fibroblast growth factor 1 and 2 which recognize specific motifs on HS (24), with a fluorophore

capable of probing HS such as the quinolinium ion or acridine orange. The resulting HS

biosensor could then be applied to the imaging of differential HS expression in vivo on the cell

surface of cancer cells. Another application of our acylation strategy could involve substitution

of the carbohydrate ligand with a peptide ligand to target intracellular proteins and enzymes or

incorporate sialic acid based ligands to target selectins and image leukocyte rolling and adhesion

during an immune response. Despite the potential advantages imparted by our approach to afford

selective carbohydrate binding protein acylation in cellular extracts and in living cells, it has not

yet been rigorously assessed in terms of its general applicability to targeting other ligand binding

proteins in different types of biological media. Studies in this direction could result in the final

development of a very powerful chemical tool to study a wide range of biological

macromolecules and their interactions under native conditions in a very time efficient, cost

effective manner. The work presented in this thesis has contributed to our current understanding

of the biomolecular blackbox known as complex carbohydrates with respect to their interactions,

their biological roles, and potential strategies to study carbohydrates under biologically relevant

settings.

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116

Appendix

NMR characterization of acylating constructs in Chapter 4

117

Figure A.1. 1H NMR(CD3OD) of 3-Mercaptopropionic acid hydrazide

118

Figure A.2. 1H NMR of S1L (300 MHz, D2O) δ 4.41 (d, J = 7.7 Hz, 1H), 4.12 (d, J = 9.0 Hz, 1H), 3.98 –

3.32 (m, 12H), 2.77 (t, J = 6.6 Hz, 2H), 2.52 (t, J = 6.6 Hz, 2H).

119

Figure A.3. 13

C NMR of S1L. (101 MHz, D2O) δ 173.49 , 102.92 , 89.89 , 78.41, 75.81, 75.47, 74.90,

72.54 , 70.96 , 70.31 , 68.57, 61.05, 60.29 , 37.61 , 19.45 . HRMS-ESI calcd (MH+) (C15H28N2O11S)

444.1413 Da; obsd 444.1417 Da.

120

Figure A.4. 1H NMR of S1M. NMR (300 MHz, d2o) δ 5.33 (d, J = 3.7 Hz, 1H), 4.07 (d, J = 9.0 Hz, 1H),

3.89 – 3.24 (m, 12H), 2.74 (t, J = 6.5 Hz, 2H), 2.47 (dd, J = 14.1, 7.5 Hz, 2H).

121

Figure A.5. 13

C NMR of S1M. (101 MHz, D2O) δ 175.47 , 172.08 (carbonate), 160.28 , 99.58 , 89.34 ,

76.90 , 76.64 , 75.43 , 72.76 , 72.60 , 71.61 , 70.33 , 69.26 , 60.84 , 60.43 , 37.42 , 32.88 , 19.84 HRMS-

ESI calcd (MH+) (C15H28N2O11S) 444.1413 Da; 444.1416 obsd Da.

122

Figure A.6. 1H NMR of 1L (300 MHz, cd3od) δ 8.53 (s, 1H), 7.58 (d, J = 9.1 Hz, 1H), 6.83 (dd, J = 9.1,

2.4 Hz, 1H), 6.57 (d, J = 2.1 Hz, 1H), 4.32 (d, J = 7.4 Hz, 1H), 3.93 (dt, J = 8.2, 7.0 Hz, 2H), 3.83 – 3.0

(m, 17H), 2.54 (t, J = 6.9 Hz, 2H), 1.24 (t, J = 7.1 Hz, 6H) HRMS-ESI calcd (MH+) (C29H42N3O14S)

687.2309 Da; obsd 687.2312 Da.

123

Figure A.7. 1H NMR of 1M .(300 MHz, cd3od) δ 8.54 (s, 1H), 7.60 (d, J = 9.1 Hz, 1H), 6.85 (dd, J = 9.1,

2.4 Hz, 1H), 6.59 (d, J = 2.2 Hz, 1H), 5.10 (d, J = 3.8 Hz, 1H), 3.94 (d, J = 8.9 Hz, 1H), 3.88 – 3.30 (m,

18H), 2.56 (t, J = 6.9 Hz, 2H), 1.26 (t, J = 7.1 Hz, 6H) HRMS-ESI calcd (MH+) (C29H42N3O14S)

687.2309 Da; obsd 687.2307 Da .

124

Figure A.8. 1HNMR of 4-mercaptobenzoate methyl ester (400 MHz, MeOD) δ 7.86 – 7.81 (m, 2H),

7.37 – 7.33 (m, 2H), 3.88 (s, 3H).

125

Figure A.9. 1H NMR of 4-mercaptobenzohydrazide (400 MHz, D2O) δ 7.40 – 7.22 (m, 1H).

126

Figure A.10. 1H NMR of S2L (399 MHz, D2O) δ 7.10 (d, J = 8.4 Hz, 2H), 7.04 (d, J = 8.4 Hz, 2H), 4.08

(d, J = 7.8 Hz, 1H), 3.86 (d, J = 8.7 Hz, 1H), 3.65 – 3.00 (m, 4H).

127

Figure A.11. 13

C NMR of S2L. (100 MHz, d2o) δ 170.40 , 151.51, 132.05, 126.93 , 125.03 , 102.93 ,

89.81, 78.37 , 75.81 , 75.33 , 74.91 , 72.49 , 70.92 , 70.46 , 68.55 , 60.99 , 60.17 . HRMS-ESI calcd

(MH+) (C19H28N2O11S) 493.1413 Da; obsd 493.1486 Da.

128

Figure A.12. 1H NMR of S2M. (400 MHz, D2O) δ 7.28 (s, 4H), 5.26 (d, J = 3.0 Hz, 1H), 4.07 (d, J = 9.0

Hz, 1H), 3.750-3.2 (m, 12H).

129

Figure A.13. 13

C NMR of S2M. (101 MHz, D2O) δ 170.21 , 131.44 , 126.96 , 125.44 , 99.59 , 89.74 ,

76.95 , 76.63 , 75.42 , 73.46 , 72.72 , 72.59 , 71.60 , 70.47 , 69.23 , 60.73 , 60.35 . HRMS-ESI calcd

(MH+) (C19H28N2O11S) 493.1413 Da; obsd 493.1483 Da.

130

Figure A.14. 1H NMR of 2L (399 MHz, CD3OD) δ 8.42 (s, 1H), 7.78 (d, J = 8.6 Hz, 2H), 7.52 – 7.45

(m, 3H), 6.77 – 6.73 (m, 1H), 6.51 (d, J = 2.3 Hz, 1H), 4.24 (d, J = 7.5 Hz, 1H), 3.98 (d, J = 8.9 Hz, 1H),

3.89 – 3.34 (m, 15H), 3.03 (dt, J = 3.3, 1.6 Hz, 1H), 1.16 (t, J = 6.5 Hz, 6H) HRMS-ESI calcd (MH+)

(C33H42N3O14S) Da; 736.2387 obsd 736.2380 Da.

131

Figure A.15. 1H NMR of 2M. (399 MHz, cd3od) δ 8.52 (s, 1H), 7.89 – 7.86 (m, 2H), 7.60 – 7.56 (m,

3H), 6.84 (dd, J = 9.1, 2.4 Hz, 1H), 6.60 (d, J = 2.3 Hz, 1H), 5.15 (d, J = 3.8 Hz, 1H), 4.06 (d, J = 8.8 Hz,

1H), 3.98 – 3.0 (m, 16H), 1.25 (t, J = 7.1 Hz, 6H) HRMS-ESI calc’d (MH ) (C33H42N3O14S) 736.2387

Da; obs’d 736.2382 Da.

132

Figure A.16. 1H NMR of 4-mercaptophenyl methylester (400 MHz, MeOD) δ 7.24 – 7.21 (m, 2H), 7.16

– 7.10 (m, 2H), 3.67 (s, 3H), 3.58 (s, 2H).

133

Figure A.17. 1H NMR of mercaptophenylhydrazide (400 MHz, D2O) δ 7.24 (d, J = 7.9 Hz, 2H), 6.90

(d, J = 7.9 Hz, 2H), 3.37 (s, 2H).

134

Figure A.18. 1H NMR of S3L. (300 MHz, D2O) δ 7.15 (d, J = 8.2 Hz, 2H), 6.92 (d, J = 8.2 Hz, 2H), 4.27

(d, J = 7.8 Hz, 1H), 3.92 (d, J = 9.0 Hz, 1H), 3.79 – 3.28 (m, 13H), 3.10 (t, J = 9.0 Hz, 1H).

135

Figure A.19. 13

C NMR of S3L (100 MHz, d2o) δ 174.12, 132.58 , 129.50, 129.08 , 128.79 , 103.09 ,

89.52 , 78.53 , 75.87 , 75.48 , 74.96 , 72.64 , 71.07, 70.36, 68.67 , 61.12 , 60.34 , 40.93. HRMS-ESI calcd

(MH+) (C20H30N2O11S) 507.1570 Da; obsd 507.1557 Da.

136

Figure A.20. 1H NMR of S3M (400 MHz, D2O) δ 7.19 (d, J = 8.0 Hz, 2H), 6.90 (d, J = 8.0 Hz, 2H), 5.26

(d, J = 3.8 Hz, 1H), 3.95 (d, J = 9.0 Hz, 1H), 3.79 – 3.25 (m, 13H), 3.12 (t, J = 9.2 Hz, 1H).

137

Figure A.21. 13

C NMR of S3M (101 MHz, D2O) δ 173.12 , 141.31, 132.47, 128.90, 128.51, 99.66, 89.41

, 76.94 , 76.64 , 75.46 , 72.79, 72.67 , 71.66 , 70.35 , 69.29, 60.82 , 60.42 , 39.92 . HRMS-ESI calcd

(MH+) (C20H30N2O11S) 507.1570 Da; obsd 507.1564 Da.

138

Figure A.22. 1H NMR of 3L (399 MHz, CD3OD) δ 8.51 (s, 1H), 7.57 (d, J = 9.1 Hz, 1H), 7.44 – 7.38

(m, 4H), 6.83 (dd, J = 9.1, 2.4 Hz, 1H), 6.59 (d, J = 2.2 Hz, 1H), 4.33 (d, J = 7.5 Hz, 1H), 3.96–3.90 (m,

2H), 3.84–3.37 (m, 11H), 3.25–3.18 (m, 2H), 1.25 (t, J = 7.1 Hz, 6H) HRMS-ESI calcd (MH+)

(C34H44N3O14S) 750.2544 Da; obsd 750.2538 Da.

139

Figure A.23. 1H NMR of 3M. (399 MHz, CD3OD) δ 8.51 (s, 1H), 7.57 (d, J = 9.1 Hz, 1H), 7.46 – 7.36

(m, 4H), 6.83 (dd, J = 9.1, 2.4 Hz, 1H) , 6.59 (d, J = 2.3 Hz, 1H), 5.14 (d, J = 3.8 Hz, 1H), 3.92 (d, J =

8.9 Hz, 2H), 3.84 – 3.37 (m, 11H), 3.27 – 3.15 (m, 2H), 1.24 (t, J = 7.4 Hz, 6H) HRMS-ESI calcd (MH+)

(C34H44N3O14S) 750.2544 Da; obsd 750.2539 Da.