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Dissecting the entry mechanism of targeting lentiviral vectors in living cells and developing quantum dot labeling of viruses for single virus tracking

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Dissecting the entry mechanism of targeting lentiviral vectors in living cells and developing quantum dot labeling of viruses for single virus tracking

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Content DISSECTING THE ENTRY MECHANISM OF TARGETING LENTIVIRAL
VECTORS IN LIVING CELLS AND DEVELOPING QUANTUM DOT
LABELING OF VIRUSES FOR SINGLE VIRUS TRACKING

By

Kye Il Joo

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMICAL ENGINEERING)

December 2009

Copyright 2009 Kye Il Joo
ii
Dedication

This thesis is dedicated to God Almighty for his sufficient grace and love. It is
dedicated to my wife Hee Moon, my parents Dae-Soo Joo and Sun-Hee Lee, my
children Joanna and Noah, and my sister and brother-in-law. It is also dedicated to my
family of God, Koinonia Community Baptist Church. Their endless support
throughout this process has been invaluable.

iii
Acknowledgements

First and foremost I would like to thank God for enabling me to achieve all
that I have achieved. In him I have found shelter, comfort, and strength.
I would like to thank my advisor Dr. Pin Wang who has guided and supported
me through this process. All my works would not have been possible without his
encouragement and patient. I would also like to thank the other members of my
dissertation committee Dr. Katherine Shing and Dr. Sarah F. Hamm-Alvarez. Dr.
Shing has been a role model for all of us in the chemical engineering department. I
had the privilege of having Dr. Hamm-Alvarez join my committee as the outside
member. She is an expert in intracellular trafficking of biomolecules and her
comments and suggestions have given me new insight and direction into my research.
I would also like to thank Dr. Lili Yang who has given valuable advice, suggestions,
and critical reviews into all my researches. I thank all the members of the Wang lab.
Most importantly, I would like to thank my parents who have provided me
with endless support. Without their love, care, and support, it would not have been
possible for me to achieve all my studies. I also thank my sister and brother-in-law for
their care and love.
I gratefully thank Pastor Frank Roh and Yuni SMN who led me to Christ, my
salvation with their countless patience, guidance, love, and prayers.
iv
I also thank Pastor Dr. You Shin Kim who has guided me to a closer walk with God,
for all his advice, prayers, and love. I would like to thank all my family of God in
Koinonia Community Baptist Church.
Finally but definitely not least, I thank my beloved wife, Hee Moon who has
taught me what the valuable and meaningful life is. I am so grateful to God that He
brought her into my life and allowed us to walk through the entire journey together
and go through all hard times together. I am truly thanking her for everything she has
done to me. I also share this unforgettable moments with my children Joanna and
Noah.
The research in this thesis was funded by a National Institute of Health Grant,
and the following reagents was obtained through the AIDS Research and Reference
Reagent Program, Division of AIDS, NIAID, NIH: Monoclonal Antibody to HIV-1
p24 (AG3.0) from Dr. Jonathan Allan; HeLa/CD4/CCR5 (TZM-bl) cell line from
Drs. John C. Kappes, Xiaoyun Wu and Tranzyme Inc.

v
Table of Contents
Dedication ii
Acknowledgements iii
List of Figures ix
Abstract xi
Chapter 1. INTRODUCTION 1
1.1 Gene therapy 1
1.1.1 Viral mediated gene delivery 1
1.1.2 Targeting lentiviral vectors to specific cell types 3
1.2 Understandings of Virus Entry Mechanism 6
1.2.1 Main routes of virus entry 6
1.2.2 Early stage of virus entry 8
1.2.3 Virus membrane fusion 11
1.2.4 Single-virus tracking in live cells 13
1.3 Quantum Dot Labeling of Viruses for Single Virus Tracking 16
1.3.1 Problems encountered in single-virus tracking 16
1.3.2 Inorganic semiconductor: quantum dots 17
1.3.3 Functionalized quantum dots conjugated with biomolecules 19

1.4 Summary and Thesis Work 21
1.4.1 Visualization of targeted transduction by engineered lentiviral vectors 21
1.4.2 Imaging multiple intermediates of single-virus membrane fusion
mediated by diverse fusion proteins 23
1.4.3 Developing quantum dot labeling of enveloped/non-enveloped
viruses for single virus tracking 25

Chapter 2. VISUALIZATION OF TARGETED TRANSDUCTION BY
ENGINEERED LENTIVIRAL VECTORS 28
2.1 Introduction 29
2.2 Results 32
2.2.1 Co-incorporation of antibody and fusogenic protein on
the engineered virion 32
vi
2.2.2 Antibody directs lentivirus to target cells 35
2.2.3 Incorporated fusogenic protein triggers virus-endosome fusion 38
2.2.4 Tracking of viral transport through endosomes 41
2.2.5 Microtubule-mediated virus transport 48
2.2.6 Actin-mediated virus transport 53
2.2.7 Virus release from endosomes 54
2.3 Discussions 57
2.4 Materials and Methods 61

Chapter 3. IMAGING MULTIPLE INTERMEDIATES OF SINGLE-VIRUS
MEMBRANE FUSION MEDIATED BY DIVERSE FUSION
PROTEINS 68
3.1 Introduction 69
3.2 Results 73
3.2.1 Engineered lentiviruses require low pH to trigger endosomal fusion 73
3.2.2 The entry of engineered lentiviruses is clathrin-dependent 74
3.2.3 Different fusion proteins modulate different kinetics of
virus-endosome fusion 78
3.2.4 Expression of dominant-negative Rab7 inhibits the transduction of
HAmu-bearing viruses 82
3.2.5 Lentivirus displaying the fusion protein HAmu, but not SINmu,
requires functional late endosome trafficking for viral fusion 85
3.2.6 Hemifusion mediated by the SINmu fusogen occurs faster
than that by the HAmu fusogen at low pH 89
3.2.7 Fusion pore formation mediated by SINmu, but not by HAmu, is
delayed after lipid mixing 92
3.3 Discussions 95
3.4 Materials and Methods 100

Chapter 4. DEVELOPING QUANTUMM DOT LABELING OF ENVELOPED
/ NON-ENVELOPED VIRUSES FOR SINGLE-VIRUS TRACKING 107
4.1 Introduction 109

4.1.1 Site specific labeling of enveloped viruses with quantum dots for
single virus tracking 109

vii
4.1.2 Enhanced monitoring of non-enveloped viruses with quantum dot
labeling 111
4.2 Results 114
4.2.1 Construction of AP-tag 114
4.2.2 Incorporation of AP-tag onto lentiviruses 116
4.2.3 Photostability of QD-labeled viral particles 119
4.2.4 Trafficking of QD-labeled lentiviruses 121
4.2.5 Clathrin/Caveolin-dependent entry of VSVG-pseudotyped
retroviruses 124
4.2.6 Intracellular trafficking of QD-labeled HIV 128
4.2.7 Covalent attachment of quantum dots on adeno-associated viruses 131
4.2.8 Photostability and detection sensitivity of QD-labeled AAV2 134
4.2.9 Clathrin/Caveolin-dependent entry of AAV2 136
4.2.10 Trafficking of AAV2 through endosomes 138
4.2.11 Cytoskeleton-mediated AAV2 transport 140
4.3 Discussions 142
4.4 Materials and Methods 145
References 153

viii
List of Figures
Figure 1.1: The Schematic representation of lentivirus-mediated
gene delivery 3
Figure 1.2: The schematic of proposed lentiviral transduction pathway
in target cells 5
Figure 1.3: Two main virus entry pathways 8
Figure 1.4: Different pathways of endocytosis 9
Figure 1.5: Transition States in Membrane Fusion 12
Figure 1.6: Properties of bioconjugatable quantum dots (QDs) 18
Figure 1.7: Schematic representation of a proposed entry mechanism
for engineered lentiviruses 24
Figure 1.8: General strategy for the site-specific labeling of
enveloped viruses with quantum dots 26
Figure 1.9: Covalent attachment of quantum dots on adeno-associated
virus serotype 2 (AAV2) 27
Figure 2.1: Co-incorporation of antibody and fusogenic protein on
the single lentivirus particle 34
Figure 2.2: Antibody-mediated targeting of virus to a CD20-expressing
cell line 37
Figure 2.3: Detection of virus-endosome fusion at different time points 43
Figure 2.4: Internalization and transport of viruses through endosomes 45
Figure 2.5: The trafficking of the viral particles through various endosomes 47
Figure 2.6: Microtubule-associated transport of viruses 51
Figure 2.7: The effects of inhibitory drugs or siRNA treatment on
viral fusion, infection, and endosome maturation 52
Figure 2.8: Actin-associated transport of viruses 54
ix
Figure 2.9: Time series images of the viral core release from an endosome 56
Figure 2.10: Time series images of the viral core release from an endosome 56
Figure 3.1: Engineered lentiviruses can enter target cells via endocytosis 75
Figure 3.2: Clathrin/caveolin-dependent entry of engineered lentiviruses 78
Figure 3.3: Visualization of virus-endosome fusion at different time points
for engineered lentiviruses displaying SINmu or HAmu 81
Figure 3.4: Inhibition of virus entry by Rab5 and Rab7 dominant-negative
mutants 84

Figure 3.5: Detection of viral fusion in DsRed-Rab5- and GFP-Rab7-
expressing cell 87
Figure 3.6: Selected frames from the monitoring of viral fusion in
DsRed-Rab5- and GFP-Rab7-expressing cells 88
Figure 3.7: Detection of hemifusion between viruses and target cells
with a pH sensor 91

Figure 3.8: Hemifusion and fusion pore formation between viruses
and target cells 94
Figure 4.1: General strategy for the site-specific labeling of enveloped
viruses with QDs 113
Figure 4.2: Incorporation of AP tag onto the surface of 293T cells 115
Figure 4.3: Incorporation of AP tag onto the surface of lentiviruses
for QD labeling 118
Figure 4.4: Photostability comparison between QD-labeled or
FITC-labeled viral particles 120
Figure 4.5: The trafficking of QD-labeled viral particles through endosomes 123
Figure 4.6: Clathrin/caveolin-dependent entry of VSVG-pseudotyped
retroviruses 127
Figure 4.7: Binding and intracellular trafficking of QD-labeled HIV 130

x
Figure 4.8: Covalent attachment of quantum dots on adeno-associated
virus serotype 2 132

Figure 4.9: Photostability and detection sensitivity of
QD-labeled AAV2 135
Figure 4.10: Clathrin/caveolin-dependent endocytosis of AAV2 137
Figure 4.11: Trafficking of AAV2 through endosomes 139
Figure 4.12: Cytoskeleton-mediated viral transport 141

xi
Abstract

A strategy to target lentiviral vectors to specific cell types holds great promise
for future clinical applications of gene therapy. We have previously developed an
efficient method to target lentivirus-mediated gene transduction by introducing a
targeting antibody and pH-dependent fusogenic protein as two distinct molecules on
the lentiviral surface. However, the molecular mechanism that controls the targeted
infection needs to be defined. To elucidate the endocytic pathway of the engineered
lentivirus, we have monitored intracellular trafficking of the individual lentiviruses in
the targeted cells by various direct visualization approaches. This study proposed that
the fusogen-mediated membrane fusion could be a rate-limiting step of targeting
lentivirus transduction. However, the specific features of the fusogen-associated
membrane fusion that control the targeted infection still remain largely unknown.
Therefore, we further demonstrated the intracellular behaviors of two engineered
lentiviruses displaying a class I fusogen derived from Sindbis virus glycoprotein or a
class II fusogen derived from influenza virus hemagglutinin by tracking the
individual viral particles in target cells. Our results suggest that both engineered
lentiviruses enter target cells through clathrin-dependent endocytosis. However, the
different kinetics of virus-endosome fusion as well as the distinct requirement of
endosomal traffic for viral fusion of two engineered lentiviruses was suggested by
imaging multiple sequential steps of fusion event in target cells. These imaging
xii
studies shed some light on the infection mechanism of the engineered lentivirus and is
beneficial to the design of more efficient gene delivery vectors.
Good photostability of the labeling fluorophores is always desirable for the
continuous tracking of individual viruses or any other biomolecules, and also, with
the development of new imaging techniques requiring rapid and continuous excitation
of fluorophores for z-stack image acquisition (3D reconstruction) and time-lapse
imaging, greater photostability is necessary for detailed trafficking studies.
Traditionally, organic fluorophores are used to detect and track biomolecules, such as
antibodies, peptides, and viruses. However, problems with metabolic degradation, or
photobleaching, have limited their applicability for long-term imaging of biological
processes. The use of quantum dots can potentially mitigate these concerns, as well as
allow for the development of novel detection techniques. Quantum dots are resistant
to metabolic degradation and posses a wide absorption spectrum and a narrow
emission spectrum. Here, we report general strategy to label enveloped and non-
enveloped viruses with quantum dots for single-virus tracking. The results indicated
that quantum dot labeling holds several advantages over conventional organic dyes,
such as greatly improved photostability against photobleaching and detection
sensitivity, which leads to the enhanced and detailed monitoring of viral behaviors in
cells. We believe that these labeling methods can take advantage of the excellent
fluorescence property of QDs and may represent an attractive tool for elucidating the
molecular details of entry and intracellular transport of many kinds of enveloped and
non-enveloped viruses.
1
Chapter 1. INTRODUCTION

1.1 Gene Therapy
1.1.1 Viral mediated gene delivery
Gene therapy is generally defined as the introduction of genes into a target
cell or tissue to provide a therapeutic advantage. This has been increasingly
considered as the most promising method to treat or eliminate the cause of disease by
insertion of functional gene into a target cell (Mountain, 2000; Somia, 2000; Verma,
1997). In order to achieve the major aim of gene transfer, many efforts have been
made to develop gene delivery vectors. Non-viral vector based approaches (e.g. naked
DNA, DNA complexes with cationic lipids, and particles comprising DNA condensed
with cationic polymers) has been developed as a carrier to delivery DNA into cells,
which have no insert-size limitation and less immunogenicity. However, it has the
notable disadvantage of low transfection efficiency (Mountain, 2000; Verma, 1997).
Viral vector-mediated gene delivery system attempts to utilize aspects of the
natural life cycle of viruses to achieve high gene delivery efficiency. Many different
viruses are being adapted as vectors, but the most advanced are retrovirus, adeno-
associated virus, and adenovirus. Among these vectors, gamma retroviral vectors are
currently the most commonly used gene delivery vehicles due to their ability to
2
permanently integrate a therapeutic transgene into a target cell chromosome (Daly,
2000; Verma, 1997). Furthermore, lentiviral vectors exhibit promising features for the
development of gene therapy since they have the capacity to produce long-term
transgene expression and transduce nondividing cells (Cronin, 2005; Waehler, 2007;
Weinberg, 1991; Yang, 2006). However, despite the high efficiency of viral vectors-
mediated gene transfer, in many cases the natural tropism of viruses does not satisfy
therapeutic needs (Verma, 1997; Waehler, 2007). Sometimes, in order to achieve a
desirable therapeutic effect with gene therapy, the viral vectors must be capable of
precisely delivering a gene of interest to specific target cells without influencing non-
target cells (Cronin, 2005; Waehler, 2007; Yang, 2006). Many efforts have been
made to develop such targeting virus systems mostly by altering the viral envelope
glycoprotein which determines cell tropism (Gollan, 2002; Lavillette, 2001; Maurice,
2002; Nguyen, 1998; Sandrin, 2003; Somia, 1995). However, although gamma-
retroviral and lentiviral vectors are highly permissive for incorporation of
heterologous viral proteins, this manipulation adversely affects their host range or
viral titers due to the delicate coupling interactions of binding and fusion domains of
glycoproteins (Desmaris, 2001; Mochizuki, 1998; Sandrin, 2003; Schnierle, 1997;
Waehler, 2007).

3

Figure 1.1: The schematic representation of lentivirus-mediated gene delivery. Lentiviral vectors have
the capacity to produce long-term transgene expression and transduce nondividing cells. (From Ref.
(Verma, 1997))

1.1.2 Targeting lentiviral vectors to specific cell types
Most of the clinical trials for curing genetic diseases involve engineering
specific cells types ex vivo and then transplanting back the modified cells. However,
this process is both time-consuming and expensive. Thus, targeting specific cell types
in vivo holds great potential for future clinical applications. Lentiviral vectors hold
great promise for the development of gene therapy due to their ability to produce
long-term transgene expression and transduce nondividing cells.
4
We have previously developed an efficient method to target lentivirus-
mediated gene transduction to a desired cell type (Yang, 2006). Our engineering
approach involved the incorporation of a targeting antibody and pH-dependent
fusogenic protein as two distinct molecules on the lentiviral surface. For recognition,
we use antibodies, and for fusion, we use viral glycoprotein (i.e. fusogenic protein)
that has mutated to inactivate its binding ability. Our hypothesis for targeted
transduction was that the antibody binding induces endocytosis, and then the virus is
brought into an endosomal compartment where the low pH environment causes the
fusogenic molecule to trigger membrane fusion and release the viral core into the
cytosol. After reverse transcription and migration of the product to the nucleus, the
genome of the vector should integrate into the target cell genome, incorporating the
vector’s transgene into the cell’s inheritance.
So far, many of virus entry mechanisms have been biochemically defined, but
still those processes such as endocytosis, virus fusion, endosome-mediated transport,
especially when and how they occurs, still remain unclear. Improved understanding
of virus-host cell interactions can provide crucial insights for enhancing the efficacy
of virus-mediated gene delivery as well as preventing virus-triggered diseases. Insight
into the dynamics of the trafficking of viral particles in living cells is fundamental to
understanding a variety of the viral infection mechanisms. In this study, we intended
to develop virus-trafficking and virus-labeling methods to understand variety the viral
infection mechanisms.

5

Figure 1.2: The schematic of proposed lentiviral transduction pathway in target cells.

6
1.2 Understandings of Virus Entry Mechanism
1.2.1 Main routes of virus entry
An efficient transport of genetic materials from the cell surface to the nucleus
is a key requirement for viral infection. During viral entry, the virus interacts with
various cellular structures and takes advantage of their environments to optimize the
delivery of the viral genome to the nucleus and promote efficient viral replication
(Marsh and Helenius, 2006). Understanding virus trafficking involves understanding
both the interactions between the virus and cell, including the cellular structures, and
the viral infection routes (Brandenburg and Zhuang, 2007). Improved understanding
of their interactions can provide crucial insights for preventing virus-triggered
diseases, as well as enhancing the efficacy of virus-mediated gene delivery.
There are many routes taken for infection and they vary by virus type.
Generally, viruses attach to the receptors on the cell surface, such as proteins,
carbohydrates, or lipids, and deliver their viral genome into the cellular cytoplasm
(Anderson and Hope, 2005; Dimitrov, 2004; Gruenberg, 2001; Klasse, 1998;
Pelkmans, 2003). Viruses can use either an endocytic or non-endocytic route to enter
cells (Fig. 1.3). Many viruses enter cells through the endocytic pathway, which
involves clathrin- or caveolin-dependent pathways, or a clathrin- and caveolin-
independent pathway (Brodsky FM, 2001; DeTulleo and Kirchhausen, 1998;
Kirchhausen, 2000; Marsh and Helenius, 2006; Nabi and Le, 2003; Nichols and
Lippincott-Schwartz, 2001; Sieczkarski and Whittaker, 2002). Viruses that use the
7
endocytic route undergo a conformational change of the viral entry proteins or target
cell receptors triggered by the low endosomal pH, which is subsequently followed by
endosomal fusion for enveloped viruses or endosomal escape for non-enveloped
viruses (Harrison, 2008; Kielian and Rey, 2006). Viruses that use the non-endocytic
route enter cells by directly crossing the plasma membrane of the cell (i.e. cell
membrane fusion) at neutral pH. Some examples of viruses that use the non-
endocytic route include the human immunodeficiency virus type-1 (HIV-1) and the
Herpes simplex virus 1 (HSV-1) (Dimitrov, 2004). The cell membrane components
(e.g. lipid raft) (Manes, 2003; Nayak, 2002; Rawat, 2003) and cytoskeletons, such as
microtubules and actin-filaments (Anderson and Hope, 2005; Apodaka, 2001; Mallik
and Gross., 2004), have also been suggested to have an essential role in viral entry.
The ability to track individual viruses enables the possible elucidation of previously
unknown but critical steps involved in the penetration of viruses into cells and
dissemination of viruses, revealing novel therapeutic opportunities for controlling
virus pandemics and pathogenesis (Brandenburg and Zhuang, 2007; Marsh and
Helenius, 2006).

8

Figure 1.3: Two main virus entry pathways. a. Clathrin-mediated endocytosis, for example,
adenovirus. b. fusion at the cell membrane, for example, HIV. (From Ref. Dimitrov, 2004)

1.2.2 Early stage of virus entry
Most viruses enter cells via receptor-mediated endocytosis. However, the
entry mechanisms used by many of them remain poorly understood (DeTulleo and
Kirchhausen, 1998; Doxsey et al., 1987; Martin, 1991; Matlin KS et al., 1982).
9
Largely unknown is the way in which viruses are targeted to cellular endocytic
machinery to access the entry to cells. There are different pathways of endocytosis for
internalization. Phagocytosis mostly occurs in macrophage cells, and the entry of
many of non-viral particles is involved in macropinocytosis, which are receptor-
independent endocytosis (Mayor and Pagano, 2007).

Figure 1.4: Different pathways of endocytosis. (From Ref. (Mayor and Pagano, 2007))

For clathrin-dependent pathway, viruses bound to receptors are targeted
clathrin-coated pits (CCPs), and then CCPs are matured to clathrin-coated vesicle

10
(CCVs) resulting in the internalization of viruses. After uncoating of CCVs, viruses
are further transported to early endosomal compartments. It has been reported that
Semliki Forest Virus (SFV), Vesicular Stomatitis Virus (VSV), influenza virus, and
Hepatitis C virus (HCV) use clathrin-dependent pathway for internalization
(Blanchard et al., 2006; DeTulleo and Kirchhausen, 1998; Meertens et al., 2006; Rust
et al., 2004). Caveolae, 50~80 nm flask-shaped plasma membrane invagination, is
also used as a pathway for Simian virus 40 (SV40) (Pelkmans et al., 2001). Recently,
clathrin- and caveolae-independent pathways have also been studied for entry
mechanisms (Mayor and Pagano, 2007).
Recent studies have shown that intracellular virus trafficking is critically
involved in the endosome-mediated sorting and transport of influenza virus, vesicular
stomatitis virus (VSV), and semliki forest virus (SFV) (Lakadamyali et al., 2006;
Sieczkarski and Whittaker, 2003; Vonderheit and Helenius, 2005). The endocytic
pathways used by some viruses have been explored, but some specific features of the
entry mechanisms of engineered recombinant lentiviruses remain largely unknown. In
order to understand the interactions between the engineered lentivirus and the
targeted cells and the underlying mechanisms of viral transduction at a molecular
level, we intended to develop single-virus tracking techniques to directly visualize the
intracellular behavior of the virus in living cells.

11
1.2.3 Virus membrane fusion
Successful infection of envelope viruses to target cells usually requires the
merger of their membranes so that the genetic materials of viruses can be transferred
to the cytosol of the host cells. This membrane integration process is mediated by
fusion proteins on the surface of viruses. Viral fusion proteins lower a huge energy
barrier of the fusion reaction by using the activation energy provided upon their
conformational change to reach an energetically stable state (Carr et al., 1997;
Dimitrov, 2004; Hogle, 2002; Mothes et al., 2000). It is generally suggested that virus
membrane fusion proceeds through sequential multi events (Chernomordik and
Kozlov, 2003; Harrison, 2008; Jahn et al., 2003; Kuhn et al., 2002). Initially,
receptor-binding and/or acidic pH environment induces the conformational change of
viral fusion proteins, which yields a contact between the viral and the target
membrane. The proximal leaflets of two distinct lipid membranes are then merged
while the distal leaflets remain unchanged, called hemifusion (lipid mixing), which is
subsequently followed by the fusion pore formation via merging of the distal leaflets
(content mixing) that establishes the first aqueous connection between two apposed
membranes (Fig. 1.5) (Jackson and Chapman, 2008; Lentz et al., 2000; Zimmerberg
et al., 1994). Conventional fusion assays (e.g. cell-cell and virus-liposome fusion
analysis) have been widely developed to understand the viral membrane fusion
process, but they are generally limited to study precise kinetics of the single viral
fusion events as well as monitor a series of transient fusion intermediates formed
during a single virus fusion with target cell membrane.
12
Viral fusion proteins are currently defined into two groups, based on
important structural features: class I fusion protein exemplified by influenza
hemagglutinin (HA) and HIV-1 (gp120/gp41) are trimeric structures in both pre-
fusion and post-fusion conformations and triggers fusion using α-helical coiled-coil
domains (Dimitrov, 2004; Harrison, 2008). Class II fusion proteins represented by the
E1 protein of alphaviruses such as Semliki Forest virus (SFV) and Sindbis virus
(SIN) convert dimeric to trimeric structures during fusion triggering fusion with β-
sheet (Kielian and Rey, 2006). The direct visualization of the viral fusion process can
potentially mitigate many of these obstacles in single-virus fusion monitoring. The
observation of the actual fusion event and its kinetic characterization at the single
virus level can provide a critical understanding of the underlying mechanisms of the
virus infection in greater detail.

Figure 1.5: Transition States in Membrane Fusion. (From Ref. (Reinhard Jahn, 2003)

13
1.2.4 Single-virus tracking in live cells
Improved understanding of virus-host cell interactions can provide crucial
insights for enhancing the efficacy of virus-mediated gene delivery as well as
preventing virus-triggered diseases. Insight into the dynamics of the trafficking of
viral particles in living cells is fundamental to understanding a variety of the viral
infection mechanisms. Single-virus tracking potentially allows us to monitor dynamic
interactions of individual viral particles or viral components with the cellular
endocytic machinery. Traditionally, to track single viruses in live cells, both the virus
and the relevant cellular structures must be fluorescently labeled, then fluorescence
microscopy is employed to detect and track the labeled particles. One key concern is
fluorescently labeling the virus and appropriate cellular structures in a way that they
are bright enough to see and track but not so much so that the functions of the cell and
virus are affected. Antibodies to viral proteins for virus labeling cannot be used for
this reason; they block the function of viral proteins after binding. There are two
methods to incorporate a fluorescent label onto the cell or virus. First, fluorescent
proteins may be used by encoding them in the viral genes, thereby incorporating them
with viral structures (Joo and Wang, 2008; McDonald et al., 2002; Melikyan et al.,
2005). Since several copies are required per virion, capsid or tegument proteins are
usually selected, thus, this method can only be used with certain types of viruses. The
second method is to use a chemical label, which can be either covalently or non-
covalently attached to proteins. For example, a virus has many NH2 groups on its
surface. Amino-reactive dyes can attach to them; although there is a concern that the
14
non-specific modification of lysine residues on the viral protein may disrupt the viral
functions and infectivity, this labeling method has been relatively well-characterized
for labeling the capsid proteins of non-enveloped viruses (Bartlett et al., 2000; Greber
et al., 1997). Lipophilic dyes, which can be spontaneously incorporated into the outer
membrane of enveloped viruses, such as 1,1'-dioctadecyl-3,3,3',3'-
tetramethylindodicarbocyanine (DiD), can be used to label the viral envelope
membrane (Joo and Wang, 2008; Lakadamyali et al., 2003; (Sakai, 2006). This
membrane labeling method is widely used for studies of virus-target membrane
fusion mechanisms, but the virus cannot be tracked after the membrane fusion
process occurs. Therefore, these dyes have limited applications for imaging.
After fluorescent labeling, the viruses can be tracked in live cells through
detection with fluorescence microscopy. There are three principle setups for these
microscopes (Brandenburg and Zhuang, 2007). Epi-fluorescence microscopy has the
simplest setup. It features low signal loss, large imaging depth, and rapid wide-field
detection. However, the auto-fluorescence inherent in cells results in high background
noise for this setup and makes it unfavorable for situations where a small number of
fluorescent molecules must be detected. The second type of fluorescent microscope is
the confocal. This can be either in the form of the laser scanning confocal, for typical
use, or the spinning-disc confocal, for when fast dynamics in live cells need to be
imaged. Confocal microscopes have much lower background noise and are able to
produce 3-dimensional images, but lose much of the fluorescence signal during the
imaging process. Total internal reflection fluorescence (TIRF) microscopes allow
15
wide-field imaging with less signal loss than confocal microscopes and less
background noise than epi-fluorescence microscopy. Although the limited imaging
depth provides superior resolution for cell surface imaging, the drawback of this
technology is also its imaging depth – TIRF microscopy is only able to image at a
depth of 100-200 nm. Thus, only events that occur near the cell surface can be
imaged, such as the detection of viral entry.

16
1.3 Quantum Dot Labeling of Viruses for Single Virus Tracking
1.3.1 Problems encountered in single-virus tracking
There are several difficulties faced in fluorescence labeling and imaging of
viral particles. First, the small size of viral particles limits the number of fluorescent
probes that can be attached without causing a self-quenching effect. Too many dye
molecules can also affect the infectivity of the virus. For example, with the adeno-
associated virus, more than a few dye molecules attached to the virus causes it to
become non-infectious (Seisenberger et al., 2001). Another problem lies in the size of
the fluorescent proteins. The large and bulky molecules could lead to the disruption of
viral structures and a marked loss of viral infectivity (Muller et al., 2004). The major
problem for virus labeling with either dyes or fluorescent proteins is that fluorophores
can be photochemically destroyed in an excitation-induced phenomenon called
photobleaching. This limits the long-term tracking of single viruses in living cells,
which is necessary in monitoring the dynamic interactions between the virus and
cellular structures. The small size of the virus also necessitates longer exposure times
for signal detection, which also leads to photobleaching.
The use of quantum dots, tiny light-emitting nanocrystals, can potentially
mitigate many of these obstacles in single-virus tracking. Quantum dots exhibit
remarkable photostability and brightness, while maintaining a broad absorption
spectrum and narrow emission spectrum. They are inorganic molecules, which gives
17
them an improved resistance to metabolic degradation, or photobleaching, over the
organic fluorophores. This enables longer tracking of viral particles in living cells.

1.3.2 Inorganic semiconductor: quantum dots
Quantum dots are tiny light-emitting crystals with sizes in the nanometer
scale. Their small size causes a confinement of the electron-hole pairs in the crystal
and, as a result, they display discrete energy levels (Michalet et al., 2005). These
energy levels can absorb and emit wavelengths from ultraviolet to infrared depending
on the size of the quantum dot (Bruchez et al., 1998). Since quantum dots have
several advantages over conventional organic dyes, they are emerging as a fluorescent
probe for biological imaging and medical diagnostics. Organic dyes generally have
trouble fluorescing continuously for long periods of time due to metabolic
degradation and it is difficult to use them for multi-color applications, since they can
only be excited by light of a narrow wavelength while they emit light with a broad
spectrum, which often causes the overlapping of signals for different dyes.
Meanwhile, quantum dots are highly resistant to metabolic degradation and exhibit
high quantum yield – the ratio of the amount of light emitted from a sample to the
amount of light absorbed by the sample (Dabbousi et al., 1997; Hines and Guyot-
Sionnest, 1996). They also have molar extinction coefficients 10 to 100 times larger
than those of most organic dyes (Ballou et al., 2004). These properties allow quantum
dots to fluoresce brighter and for longer periods of time than conventional organic
dyes.
18
Quantum dots also have an advantage over organic dyes in terms of their optical
spectra (Fig. 1.6). In addition to the narrow absorption and broad emission spectrums
of organic dyes, they also experience red-tail, a phenomenon caused by the
asymmetric shape of their emission spectra and the consequent leakage of signal
towards the red end of the spectrum (Alivisatos et al., 2005; Jaiswal and Simon,
2004). Quantum dots, on the other hand, have broad absorption spectra while having
narrow and symmetric emission spectra. Thus, a single wavelength is sufficient to
excite multicolor quantum dots of different sizes, resulting in discrete emission
wavelengths with low signal overlap.

Figure 1.6: Properties of bioconjugatable quantum dots (QDs). (A) QDs are inorganic fluorophores and
consist of a cadmium selenide (CdSe) core with several layers of a thick zinc sulfide (ZnS) shell to
improve quantum yield and photostability. (B) The excitation spectrum (broken lines) of a QD (green)
is very broad, whereas that of an organic dye (rhodamine, orange) is narrow. The emission spectrum
(unbroken lines) is narrower for a QD (green) than for organic dyes (rhodamine, orange). Values
indicate the full spectral width at half-maximum intensity (FWHM value). (C) The emission of the
QDs can be tuned by controlling the size of the CdSe core: an increase in the size of the core shifts the
emission to the red end of the spectrum. The combined size of the core and the shell of QDs emitting
in the visible region of spectra are in the size range of commonly used fluorescent proteins such as
green fluorescent protein (GFP) and DsRed. (From Ref. (Jaiswal and Simon, 2004))

19
1.3.3 Functionalized quantum dots conjugated with biomolecules
To use quantum dots in biological applications, such as probing live cells,
they must first be conjugated with biological molecules. This needs to done carefully
to avoid disrupting the functions of the cell or the biological molecules. Currently,
quantum dot conjugation with biological molecules has been successfully performed
in several ways, including electrostatic interaction, mercapto (-SH) exchange, and
covalent linkage. With the electrostatic exchange approach, the negatively charged
quantum dot interacts electrostatically with the positively charged domain of the
engineered protein of interest. The resulting protein-quantum dot conjugates were
very stable and remarkably, displayed a fluorescence yield even higher than that of
non-conjugated quantum dots (Mattoussi et al., 2000). A mercapto exchange process
can also be used to conjugate biological molecules containing thiol groups, which
serve as an anchor to the ZnS shell of quantum dots via ligand exchange (Akerman et
al., 2002; Mitchell et al., 1999; Rosenthal et al., 2002; Willard et al., 2001; Winter et
al., 2001; Zhang et al., 2000). However, since the bond between Zn and thiol is
dynamic and weak, the biological molecules easily detach and the quantum dots
precipitate out of the solution. Biological molecules can also be covalently linked to
functional groups on quantum dot surfaces (Alivisatos et al., 2005; Bruchez et al.,
1998; Chan et al., 2002; Chan and Nie, 1998; Gerion et al., 2002; Gerion et al., 2001;
Pathak et al., 2001). This is done by cross-linking certain functional groups, such as -
COOH, -SH, or -NH2 on the quantum dot surface to corresponding reaction groups
on the biological molecules.
20
The resulting quantum dot-biological molecule structures are much more stably
linked than those produced by the mercapto exchange process. One example of a
cross-linker is 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC), which is able
to covalently link -NH2 and -COOH groups. Another cross-linker, 4-(N-
maleimidomethyl)-cyclohexanecarboxylic acid N-hydroxysuccinimide ester (SMCC)
can join -SH and -NH2 groups (Gao et al., 2005). Avidin/streptavidin-conjugated
quantum dots have also been employed to tag biotinylated proteins of interest via the
affinity of avidin/streptavidin for biotin (Cambi et al., 2007; Howarth et al., 2005).

21
1.4 Summary and Thesis Work
1.4.1 Visualization of targeted transduction by engineered lentiviral vectors
Lentiviral vectors hold great promise for the development of gene therapy due
to their ability to produce long-term transgene expression and transduce nondividing
cells. Although many attempts have been made to develop more efficient gene
delivery systems by using lentivral vectors, the underlying mechanisms of the viral
entry still remain elusive. Intracellular trafficking of the viral vectors is essential to
understand a variety of virus- host cell interactions. Previously, we developed the
targeting lentiviral vector by incorporation of antibody and fusogenic protein as two
different molecules on the viral surface1. To elucidate the endocytic pathway of the
engineered lentivirus, we have tracked the individual lentiviruses in the targeted cells
by labeling the virus with the GFP-Vpr fusion protein. By combining the GFP-Vpr
tagging with other labeling techniques, we visualized the surface-displayed proteins
on a single virus and the antibody induced targeting to a desired cell type. We also
revealed the dynamics of the virus fusion with an endosome, and endosome-mediated
transport of the viruses in the infected cells. Our results suggest that the fusion
between the engineered lentivirus and endosomes takes place in the early endosomal
compartment, and the completion of the virus-endosome fusion to release the viral
core into the cytosol is correlated with the endosome-mediated maturing processes.
These visualization approaches can be extended to study the virus entry process of
other engineered lentiviral vectors in living cells. The current method with other
22
virus-labeling techniques can also be used for exploring the infection pathway of
HIV. In chapter 1.2, this topic will be discussed.

23
1.4.2 Imaging multiple intermediates of single-virus membrane fusion mediated by
diverse fusion proteins
Fusion of two lipid bilayers is a prerequisite step for infection by viruses
surrounded by a lipid envelope during their entry into target cells. Membrane fusion
is triggered by viral fusion proteins, which involves sequential multiple steps in the
fusion process. However, the fusion mechanism containing multiple transient
intermediates catalyzed by the two different class fusion proteins has not been well
determined. Understanding of the different fusion kinetics and dynamics driven by
various fusion proteins can provide a crucial understanding of the underlying
mechanisms of the virus infection and reveal certain previously uncovered fusion
details. One obstacle to compare the fusion reaction mediated by different viruses in
living cells is their different receptor binding, which may subsequently lead to
different intracellular viral trafficking involved in the fusion process. To this end, we
designed a synthetic virus platform that consists of lentiviral particles co-enveloped
with a surface antibody as the binding protein along with a fusion protein derived
either from influenza virus (HAmu, class I fusogen) or from Sindbis virus (SINmu,
class II fusogen) onto the lentiviral surface (Fig. 1.7), which allows for better
understating of differential fusion mechanisms driven by diverse fusion proteins. In
this report, the single virus tracking study of the early internalization indicates that
both HAmu- and SINmu-lentiviruses enter cells through clathrin-dependent
endocytosis. However, different requirements of endosomal trafficking for the
membrane fusion of these two lentiviruses were observed.
24
The direct visualization of single viral fusion events clearly showed that hemifusion
mediated by SINmu (class II fusion protein) upon exposure to acidic pH occurs faster
than that mediated by HAmu (class I fusion protein). Real-time monitoring of
sequential fusion processes with a dual labeling of outer and inner leaflets of viral
membrane also suggested that formation of fusion pore was remarkable delayed after
hemifusion mediated by SINmu (class II fusion protein) as compared with that
mediated by HAmu (class I fusion protein). By this approach, we have demonstrated
that the combination of this versatile platform and single virus tracking can be a
powerful tool for understanding molecular details of fusion mediated by various
fusion proteins. These results will be discussed in chapter 1.3.

Figure 1.7: Schematic representation of a proposed entry mechanism for engineered lentiviruses
enveloped with a CD20-specific surface antibody (αCD20) and a fusion protein (HAmu or SINmu).

25
1.4.3 Developing quantum dot labeling of enveloped/non-enveloped viruses for single
virus tracking
Enveloped viruses are enclosed by a lipid membrane, which is picked up from
the host cell membrane during the budding process, or when the virus moves through
the cell’s membrane system such as the endoplasmic reticulum or the Golgi
apparatus. Examples of enveloped viruses include human immunodeficiency virus
(HIV), influenza virus, severe acute respiratory syndrome (SARS) virus, and hepatitis
C virus. Since conditions that damage membranes will also damage the envelope for
these viruses, they are usually more fragile than non-enveloped viruses. Thus, more
gentle and non-disruptive labeling methods are needed.
One method of labeling enveloped virus with quantum dots is through a site-
specific scheme (Fig. 1.8) (Joo et al., 2008). The strategy was to first incorporate a
15-amino acid biotin acceptor peptide (AP) tag onto the surface of a virion (Beckett et
al., 1999; Howarth et al., 2005). Subsequently, biotin ligase (BirA) was used to
specifically modify the AP-tag to introduce the biotin moiety to the viral surface. Due
to the tight interaction between biotin and streptavidin (Kd = 10
-13
M) (Piran and
Riordan, 1990), the further addition of streptavidin-conjugated quantum dots allowed
the site-specific labeling of viral particles with photostable and fluorescent quantum
dots. The colocalization assay with an antibody specific for HIV capsid protein (p24)
confirmed that AP-tag could be incorporated onto the surface of lentivirus for
quantum dot labeling. This approach has been shown to have no effect on virus
infectivity for both the lentivirus and gamma-retrovirus.
26
Good photostability of the labeling fluorophores is desirable for the
continuous tracking of individual viruses because a high magnification objective has
to be used in order to detect these tiny viral particles (20-100 nm), generating high
excitation light intensity in the focal plane of the objective (Wu et al., 2003). Also,
with the development of new imaging techniques requiring rapid and continuous
excitation of fluorophores for z-stack image acquisition (3D reconstruction) and time-
lapse imaging (Levi et al., 2005), greater photostability is necessary for detailed
trafficking studies. Compared with traditionally labeled fluorophores, these quantum
dot-labeled viruses exhibited excellent photostability against photobleaching (Joo et
al., 2008).

Figure 1.8: General strategy for the site-specific labeling of enveloped viruses with quantum dots.

27
Adeno-associated virus (AAV), a member of the non-envelope parvoviridae
family, has attracted considerable interest because it shows great promise for use in
human gene therapy. However, the membrane penetration and uncoating mechanisms
are poorly understood for these viruses, even though they are relatively small and
simple in structure. Since the AAV virion is only about 20 nm in diameter, the
number of dye molecules that can be attached to a single virus without causing self-
quenching or affecting viral infectivity is very limited. Quantum dots are much
brighter than the conventional fluorophores, which can allow the detection of viruses
with much lower amounts of labeling molecules. This study also reports the general
strategy for linking AAV virion with quantum dots through a coupling reaction for a
single-virus tracking in target cells. Intracellular trafficking of AAV, including
endosomal tracking and cytoskeleton-dependent transport will be addressed in more
detail. These will be discussed in chapter 1.4.

Figure 1.9: Covalent attachment of quantum dots on adeno-associated virus serotype 2 (AAV2). QDs-
AAV2 networks are generated by an amide-bond formation between carboxylic source of QDs and
primary amines from lysine residues on the AAV capsid via carbodiimide chemistry.
28
Chapter 2. VISUALIZATION OF TARGETED TRANSDUCTION BY
ENGINEERED LENTIVIRAL VECTORS

Portions of this Chapter are adapted from:
Kye-Il Joo and Pin Wang, Gene Ther. (2008) 15, 1384–1396

We have reported a method to target lentiviral vectors to specific cell types.
This method requires the incorporation of two distinct molecules on the viral vector
surface: one is an antibody that renders the targeting specificity for the engineered
vector, and the other is a fusogenic protein that allows the engineered vector to enter
the target cell. However, the molecular mechanism that controls the targeted infection
needs to be defined. In this report, we tracked the individual lentiviral particles by
labeling the virus with the GFP-Vpr fusion protein. We were able to visualize the
surface-displayed proteins on a single virion as well as antibody-directed targeting to
a desired cell type. We also demonstrated the dynamics of virus fusion with
endosomes and monitored endosome-associated transport of viruses in target cells.
Our results suggest that the fusion between the engineered lentivirus and endosomes
takes place at the early endosome level, and that the release of the viral core into the
cytosol at the completion of the virus-endosome fusion is correlated with the
endosome maturation process. This imaging study sheds some light on the infection
mechanism of the engineered lentivirus and is beneficial to the design of more
efficient gene delivery vectors.
29
2.1 Introduction
Gamma-retroviral and lentiviral vectors are currently the most commonly
used gene delivery vehicles due to their ability to permanently integrate a therapeutic
transgene into a target cell chromosome (Daly, 2000; Mountain, 2000; Somia, 2000;
Verma, 1997). Lentiviral vectors have the unique feature of being able to transduce
nondividing cells, making it particularly attractive for certain gene therapy
applications (Bukrinsky, 1992; Lewis, 1992; Weinberg, 1991). Sometimes, in order to
achieve a desirable therapeutic effect, the viral vectors must be capable of precisely
delivering a gene of interest to specific cells without influencing non-target cells
(Cronin, 2005; Waehler, 2007; Yang, 2006). Many efforts have been made to develop
such targeting viral vector systems mostly by altering the viral envelope glycoprotein
(Gollan, 2002; Lavillette, 2001; Maurice, 2002; Nguyen, 1998; Sandrin, 2003; Somia,
1995). Although certain envelope glycoproteins are structurally plastic enough to
allow insertion of a new molecular recognition unit (such as peptide, single chain
antibody, growth factor, etc.) for targeting, this manipulation can adversely affect the
delicate coupling interactions of the binding and fusion domains of glycoproteins,
resulting in enveloped vectors with decreased infectivity to the target cells (Cronin,
2005; Desmaris, 2001; Mochizuki, 1998; Sandrin, 2003; Schnierle, 1997).
We have previously developed an efficient method to target lentivirus-
mediated gene transduction to a desired cell type (Yang, 2006). Our engineering
approach involved the incorporation of a targeting antibody and pH-dependent

30
fusogenic protein as two distinct molecules on the lentiviral surface. Our hypothesis
for targeted transduction was that the antibody binding induces endocytosis, and then
the virus is brought into an endosomal compartment where the low pH environment
causes the fusogenic molecule to trigger membrane fusion and release the viral core
into the cytosol. In order to understand the interactions between the engineered
lentivirus and the targeted cells and the underlying mechanisms of viral transduction
at a molecular level, we intended to develop assays to directly visualize the
intracellular behavior of the virus in living cells.
Improved understanding of virus-host cell interactions can provide crucial
insights for enhancing the efficacy of virus-mediated gene delivery as well as
preventing virus-triggered diseases. Insight into the dynamics of the trafficking of
viral particles in living cells is fundamental to understanding a variety of the viral
infection mechanisms. Many enveloped viruses enter their host cells via receptor-
mediated endocytosis. The endocytosed viruses are internalized through endocytic
compartments, and the viruses fuse with the endosomal membrane to release viral
genome into host cells (Anderson, 2005; Gruenberg, 2001; Klasse, 1998; Martin,
1991). During these processes, viruses utilize microtubule networks for movement
towards the perinuclear regions (Apodaka, 2001; Mallik, 2004). Recent studies have
shown that intracellular virus trafficking is critically involved in the endosome-
mediated sorting and transport of influenza virus, vesicular stomatitis virus (VSV),
and semliki forest virus (SFV) (Lakadamyali, 2006; Sieczkarski, 2003; Vonderheit,
2005). The endocytic pathways used by some viruses have been explored, but some
31
specific features of the entry mechanisms of engineered recombinant lentiviruses
remain largely unknown.
In this study, we analyzed the intracellular trafficking of the targeting
lentiviral vectors by utilizing dynamic imaging of single viruses within target cells.
We visualized the incorporated molecules on a single virion and the targeting of
antibody-displaying virus to a CD20-expressing cell line. We also imaged viral fusion
and detected the endosome-associated transport of the engineered lentivirus. Our
results suggest that virus-endosome fusion takes place at the early endosome stage,
and that viral fusion is independent of microtubule- or actin-associated transport. We
also observed the process of the dissociation of the viral core from the fused
endosome. Our results shed some light on the infection model of the targeting
lentiviral vector incorporated with two separated binding and fusion proteins on the
surface.

32
2.2 Results
2.2.1 Co-incorporation of antibody and fusogenic protein on the engineered virion
We sought out imaging methods to characterize a previously engineered
lentivirus reported to be able to selectively transduce human B cells via CD20 as the
viral receptor (Yang, 2006). Co-display of an anti-CD20 antibody and a fusogenic
protein on the same virion is thought to be essential for this engineered lentivirus to
infect the target cell. To image the virus, we constructed lentiviral particles harboring
GFP fused to the N-terminus of the HIV accessory protein Vpr (designated GFP-Vpr,
Figure 2.1A). GFP-Vpr-labeled lentivirus enveloped with both anti-CD20 antibody
and fusogenic protein were produced as described (Yang, 2006) except with the use
of lentiviral backbone plasmid FUW lacking the GFP transgene (Figure 2.1A) instead
of FUGW and co-transfection of an additional plasmid that expresses GFP-Vpr.
During virus synthesis, GFP-Vpr provided in trans can be incorporated into the virion
via the interaction between Vpr and the P6 region of the gag protein. (McDonald,
2002) To determine whether αCD20 (anti-CD20 antibody) and SINmu (fusogenic
protein) (Yang, 2006) were incorporated on the same virion, we indirectly
immunofluorescent-stained the GFP-Vpr-tagged virions by a triple labeling method
(Figure 2.1B). As controls, we also included the staining of the GFP-Vpr-labeled
lentiviral particles bearing various surface proteins (FUW-GFPVpr/αCD20, FUW-
GFPVpr/SINmu, or FUW-GFPVpr/VSVG); VSVG (vesicular stomatitis viral
glycoprotein) is a widely used envelope glycoprotein with broad tropism.
33
Confocal images of the individual FUW-GFPVpr/αCD20+SINmu particles showed
that ~70% of the GFP-Vpr-labeled virions colocalized with both αCD20 and SINmu
(Figure 2.1C). This indicated that both the antibody and the fusogenic protein were
indeed displayed on a single virus particle. The detection of a few GFP-negative and
dye-positive spots for FUW-GFPVpr/αCD20+SINmu suggested that some of the
intact virions lacked the GFP-Vpr protein, which is consistent with the previous
report by McDoland et al.; (McDonald, 2002) some spots that were positive for
SINmu only could be virions that lacked the incorporation of the GFP-Vpr protein
and αCD20. As expected, colocalizations of the GFP-labeled virions with only
αCD20 (FUW-GFPVpr/αCD20) or with only SINmu (FUW-GFPVpr/SINmu) were
observed, while no colocalization of the GFP-labeled virions with either protein was
detected for FUW-GFPVpr/VSVG.
To test whether the GFP-Vpr-labeling of lentiviruses could affect the viral
infectivity, we made viruses bearing both αCD20 and SINmu
(FUGW/αCD20+SINmu); FUGW is a lentiviral backbone that contains a human
ubiquitin-C promoter driving the expression of a GFP transgene (Figure 2.1A) (Lois
et al., 2002). The target 293T/CD20 cells were exposed to FUGW/αCD20+SINmu
with or without the incorporation of GFP-Vpr, and the percentage of GFP-expressing
cells was measured by FACS three days post-infection. As shown in Figure 2.1D, a
similar transduction efficiency was obtained, indicating that the GFP-Vpr-labeling did
not markedly affect viral infectivity.

34

Figure 2.1: Co-incorporation of antibody and fusogenic protein on the single lentivirus particle. (A)
The schematic representation of the labeling (GFP-Vpr) and viral (FUW and FUGW) constructs.
CMV: cytomegalovirus immediate-early gene promoter; GFP: green fluorescence protein; Vpr: viral
protein R; Ubi: human ubiqutin-C promoter; WPRE: woodchuck hepatitis virus posttranscriptional
regulatory element. (B) The schematic representation of the virus-staining method for visualizing
individual viruses. Two antibodies were used to detect the presence of αCD20 and fusogenic molecule
(SINmu). (C) GFP-Vpr-labeled viral particles are colocalized with the αCD20 antibody (red) and
fusogenic molecule SINmu (blue). GFP-Vpr-labeled viruses psuedotyped by either both αCD20 and
SINmu, or αCD20 antibody only, SINmu only, or VSVG protein were strained with anti-human IgG
and anti-HA tag antibodies against αCD20 and SINmu. Overlapping green, red, and blue signals
appears as white in a merged image. Scale bar represents 2 µm. (D) 293T/CD20 cells (2 × 105) were
transduced with 2 ml of fresh unconcentrated FUGW/αCD20+SINmu virus (~ 1 × 106 TU/ml) with or
without GFP-Vpr labeling. The resulting GFP expression was analyzed by FACS. Solid line, analysis
of the infected 293T/CD20; Shaded area, analysis of the non-infected 293T/CD20 (as a control).
35
2.2.2 Antibody directs lentivirus to target cells
To examine whether the engineered lentiviral particles could efficiently
recognize the desired cell type, we analyzed the virus-cell binding complex using a
confocal microscope. A 293T cell line stably expressing the CD20 protein
(293T/CD20) was used as the target cell line, and its parental cell line 293T was used
as a negative control (Figure 2.2A). Neither GFP nor the αCD20 signal was detected
in the control 293T cells lacking CD20 expression (Figure 2.2A, upper). In contrast,
significant GFP and αCD20 signals were detected on the surface of 293T/CD20 cells
(Figure 2.2A, lower). This result suggests that our engineered lentivirus can
specifically bind to a CD20-expressing cell line.
To further confirm that the virus-cell binding was induced by the viral αCD20, the
lentiviral particles bearing various surface proteins (FUW-GFPVpr/αCD20+SINmu,
FUW-GFPVpr/αCD20, or FUW-GFPVpr/SINmu) were incubated with 293T/CD20
cells, followed by extensive washing.
The imaging results showed that the lentiviral particles bearing the αCD20 antibody
(FUW-GFPVpr/αCD20+SINmu and FUW-GFPVpr/αCD20) were able to bind to the
target cells, but no GFP signal was detected in the cells incubated with the viral
particles bearing only the fusogenic protein SINmu (FUW-GFPVpr/SINmu, Figure
2.2B). These results demonstrate that the virus-cell binding is mediated by a specific
interaction between the CD20 antigen on the cell surface and the αCD20 antibody on
the viral surface. In addition, the imaging results of virus-cell binding also suggest
that the fusogenic molecule on the viral surface does not interfere with the binding of
36
the αCD20 antibody to CD20-expressing cells (Yang, 2006). We quantified the
number of viral particles bound to the target cells by examining more than 20 cells
and counting the bound viruses. The result showed that FUW-
GFPVpr/αCD20+SINmu (15.4 particles/cell) and FUW-GFPVpr/αCD20 (16.3
particles/cell) exhibited similar binding to 293T/CD20 cells, while FUW-
GFPVpr/SINmu (0.1 particles/cell) was rarely detected on the cell surface.

37

Figure 2.2: Antibody-mediated targeting of virus to a CD20-expressing cell line. (A) Immunostaining
for the CD20 antigen expressed on the cell surface (left). 293T and 293T/CD20 cells were fixed with
formaldehyde and immunostained with anti-human CD20 antibody (red) including DAPI nuclear
staining (blue). 293T and 293T/CD20 cells were incubated with FUW-GFPVpr/αCD20+SINmu at 4°C
for 1 h, fixed, and then immunostained for αCD20 bound to the cell surface. The colocalization of
GFP-Vpr-labeled viruses with αCD20 signals (red) appears as yellow in a merged image. The boxed
region is enlarged in the right panel. (B) 293T/CD20 cells were incubated with FUW-
GFPVpr/αCD20+SINmu, FUW-GFPVpr/αCD20 alone, or FUW-GFPVpr/SINmu alone at 4°C for 1 h,
fixed, and then immunostained against αCD20 (red) and SINmu (blue) bound to cells. The boxed
regions are magnified and shown in separated panels below. Scale bar represents 5 µm.

38
2.2.3 Incorporated fusogenic protein triggers virus-endosome fusion
One hypothesis for the mechanism of viral entry was that the fusogenic
protein displayed on the surface of the virion could induce fusion in the acidic
endosome (Yang, 2006). To visualize the actual fusion event of internalized viruses
within the endosomes, we used lipophilic dye (DiD) that can be spontaneously
incorporated into viral membrane for a double-labeling of GFP-Vpr-tagged
lentiviruses. The incorporation of DiD dye at a high concentration on the viral
membrane can result in self-quenching of DiD fluorescence (Lakadamyali, 2003;
Sakai, 2006). Viral fusion with the endosome membrane can then cause a de-
quenching that is due to the dispersion of DiD and can be seen by the increase of
fluorescence (Figure 2.3A). We labeled GFP-Vpr-tagged lentiviruses (FUW-
GFPVpr/αCD20+SINmu) with DiD and incubated them with 293T/CD20 cells at
37°C for various time periods. The acquired images of both the green and red
fluorescence signals are shown in Figure 2.3C (upper), and the boxed regions are
enlarged and represented as panels below. At 0 min, the GFP-Vpr-labeled viral
particles were detected on the cell surface. After 10 min of incubation, viral particles
appeared to be internalized inside the cell but remained detectable solely by the green
signal, suggesting that although viruses were endocytosed, fusion had not occurred.
The image after 30 min of incubation showed that many particles were fused with the
endosomes, as indicated by the appearance of the red signal. After 60 min, more
endosome-fused viral particles and brighter fusion signals were observed. These
imaging results suggest that the majority of viral fusion occurs between 30 min to 60
39
min after incubation. In addition, viral fusion was observed in the peripheral region at
earlier time points (Figure 2.3C, upper, 30min), but the fusion signals at later time
points were mostly distributed around the perinuclear region of the cell (Figure 2.3C,
upper, 60min).
We compared the kinetics of viral fusion of our engineered lentivirus with that
of conventional VSVG-pseudotyped lentivirus. It is known that VSV is endocytosed
to early endosomes, where the low pH triggers endosomal fusion (Marsh et al., 1983).
Using the same fluorescence de-quenching assay, we observed the viral fusion of
FUW-GFPVpr/VSVG. The acquired images showed that virus-endosome fusion
could be detected at 10 min after incubation (Figure 2.3C, lower), indicating that viral
fusion of VSVG-pseudotyped lentivirus occurs faster than that of the engineered
targeting lentivirus.
In order to compare the timing of acidic pH-activated and fusion-involved
penetration of the engineered and VSVG-pseudotyped lentiviruses, we used
bafilomycin A1, the specific inhibitor of vacuolar proton ATPases, to block low pH
endosomal fusion. (Bowman et al., 1988) Viruses were pre-bound to 293T/CD20
cells at 4°C for 1 h, and entry was initiated by shifting cells to 37°C. At different time
points thereafter, bafilomycin A1 (25 nM) was added to block further viral fusion.
The percentage of viral entry was normalized based on the signal at 4.5 h, when no
further effect of bafilomycin A1 was observed and fusion was therefore considered to
be unrestricted by the treatment.
40
These results suggest that acid-induced penetration of VSVG-pseudotyped lentivirus
occurred ~5 min after warming, and a half-maximal penetration (50%) was reached
by ~40 min, whereas the penetration of engineered lentivirus
(FUGW/αCD20+SINmu) took place ~25 min after warming and reached a half
maximal level by ~120 min (Figure 2.3B). Consistent with the viral fusion kinetics
determined by the de-quenching assay (Figure 2.3C), we found that the engineered
lentivirus has slower kinetics of viral penetration than that of VSVG-pseudotyped
lentivirus.
To further characterize viral fusion, we performed real-time imaging of GFP-
Vpr/DiD-labeled lentiviruses in 293T/CD20 cells. We first incubated the doubly
labeled viruses with 293T/CD20 cells at 37°C for 30 min to allow the internalization
of viral particles, and then began imaging. Confocal time-lapse images were collected
cautiously to make sure that the same viral particle was tracked at ~15-second (s)
intervals over a period of 10 min and selected images are shown in Figure 2.3D. The
first image (at 334.7 s) showed that before endosomal fusion, the viral particle was
identified only by green color. At 418.8 s, viral fusion with the endosome was
visualized by the dramatic increase of fluorescence signal as the result of DiD de-
quenching (Figures 2.3D and 2.3E). Approximately 80% of the viral particles we
observed (n=34) showed this pattern of de-quenching. Interestingly, the virus-
endosome fusion signal was initially detected around the peripheral region (418.8 s)
and then migrated towards the nucleus over ~3 min (Figure 2.3D). We observed the
movement of fusion signals towards nucleus in 60% of the videos recorded (n=14),
41
while in the other 40% of cases, it appeared that the fusion signals were either
stationary or bidirectional. By combining the results derived from the fixed images of
virus-endosome fusion (Figure 2.3C) and live cell imaging (Figure 2.3D), we
conclude that virus-endosome fusion could occur between the peripheral region and
the perinuclear region. As shown in Figure 2.3E, the quantification of fluorescence
intensity of the GFP and the DiD signals showed that although some fluctuations of
the GFP signal were detected during the imaging, only the DiD signal increased
dramatically at certain moments of the recording, supporting our assumption that the
observed fluorescence change is an indication of viral fusion.

2.2.4 Tracking of viral transport through endosomes
To visualize the distribution of endosomal compartments, we used antibodies
against EEA1 (Lakadamyali, 2006; Sieczkarski, 2003; Vonderheit, 2005) and CI-
MPR (Lakadamyali, 2006; Urayama, 2004) as the early and late endosomal markers,
respectively. We performed double-staining for the early and the late endosomes
along with DAPI counterstaining in 293T/CD20. As shown in Figure 2.4A, the early
and the late endosomal markers were largely separated. The early endosomal markers
were distributed in the peripheral region, whereas the late endosomal markers were
primarily found in the perinuclear region of cells. However, a close examination of
the images identified several endosomes that were both EEA1- and CI-MPR-positive
(Figure 2.4A, arrow), which have been interpreted as maturing intermediates towards
42
the late endosomes (Rink, 2005; Stoorvogel, 1991). These intermediate endosomes
containing both markers were mostly observed in the perinuclear region of the cells.

43

Figure 2.3: Detection of virus-endosome fusion at different time points. (A) Schematic representation
of the visualization assay for virus-endosome fusion by fluorescence dequenching. (B) The time course
study of virus penetration by using bafilomycin A1. VSVG or both αCD20 and SINmu displaying
lentiviruses were prebound to 293T/CD20 cells at 4°C for 1 h. Entry was initiated by shifting the
temperature to 37°C. At the indicated time points, cells were treated with bafilomycin A1 (25 nM) to
inhibit the endosome acidification. After 3 h, the drug was removed and replaced with D10 media.
Cells were further incubated for 72 h, and the percentage of GFP+ cells was measured by FACS. The
4.5-h time point was normalized to 100% entry. (C) GFP-Vpr-labeled VSVG or both αCD20 and
SINmu displaying viruses (green) were labeled with DiD (red) for 1 h at room temperature. Double-
labeled viruses were incubated with 293T/CD20 cells at 37°C for 0, 10, 30, or 60 min, fixed and
imaged. The boxed regions are magnified and shown in separated panels below. Yellow particles
indicate viral particles fused to endosomes. Scale bar represents 5 µm. (D & E) Tracking of endosomal
fusion of individual viruses. (D) Selected images obtained from a time series study starting 30 min
after incubation. GFP-Vpr/DiD-labeled viruses were incubated with 293T/CD20 cells at 37°C for 30
min for virus internalization, and time-series images were obtained at every ~15 seconds over a time
period of 10 min. The arrow indicates the viral particle monitored in the live cell. The GFP-Vpr+
(green) signal and GFP-Vpr+DiD+ (yellow as the merged color of green and red) signals mark the
viral particle before and after endosomal fusion, respectively. Scale bar represents 2 µm. (E) Kinetics
of the fluorescence intensity of the GFP-Vpr (green) and DiD (red) signals of the virion. The
fluorescence intensity was measured within the regions of interests around viral particle using the
software package for the Zeiss LSM 510.
44
We conducted a colocalization experiment to examine the endosomal
transport of the engineered lentivirus and to analyze the virus-fused endosomes. As
shown in Figure 3B, at 0 min, the viral particles did not colocalize with either of the
endosomal markers. After 30 min of incubation, particles were colocalized with
EEA1 in the early endosomes, and after 60 min, many viral particles were observed in
endosomes positive for both EEA1 and CI-MPR. At 120 min, most of the viral
particles did not colocalize with either marker, and a small fraction was seen in the
late endosomes that stained positive only for CI-MPR. We quantified this
colocalization by viewing more than 20 cells (~100 viruses) for each time point
(Figure 2.4C) and the results were consistent with the observation seen in Figure
2.4B. Coupled with the previous evidence that virus-endosome fusion signals were
detected between 30 and 60 min after incubation (Figure 2.3C), this study suggests
that the virus-endosome fusion started before the late endosome stage.

45

Figure 2.4: Internalization and transport of viruses through endosomes. (A) Distribution of endocytic
compartments in living cells. Early and late endosomes were detected by EEA1 and CI-MPR,
respectively. 293T/CD20 cells were immunostained with EEA1 (red) and CI-MPR (green) and
counterstained with DAPI (blue). Arrows indicate individual endosomes positive for both early and
late endosomal markers. (B) GFP-Vpr-labeled viruses (FUW-GFPVpr/αCD20+SINmu) were
incubated with 293T/CD20 cells (MOI ~30) at 37°C for various time points of 0, 30, 60, or 120 min.
Then these cells were fixed, permeabilized, and immunostained with EEA1 (red) and CI-MPR (blue)
and counterstained with DAPI (white). The boxed regions are magnified and shown in separated
panels below. The bottom panels show the localization of GFP-Vpr-labeled viral particles in the two
different endosomal stages. Scale bar represents 5 µm. (C) Quantification of GFP-Vpr-labeled
lentiviruses colocalized with EEA1+ (black), CI-MPR+ (white), or EEA1+CI-MPR+ (gray)
endosomes at different incubation times. The result shown is the collective data from three
experiments.
46
To confirm the trafficking of the viral particles through various endosomes,
we further examined the colocalization of engineered viral particles with red
fluorescent protein-tagged Rab protein, DsRed-Rab5 and DsRed-Rab7, as the early
and late endosomal markers, respectively (Chavrier et al., 1990; Rink, 2005; Zerial
and McBride, 2001b). GFP-Vpr-labeled viruses were incubated with DsRed-Rab5- or
DsRed-Rab7-transfected cells at 37°C and fixed at different time points (Figure
2.5A). The time course images showed that the majority of viral particles were first
located in organelles that were positive for DsRed-Rab5 (Figure 2.5A, 30min), and
were later observed in Rab7-positive and Rab5-positive organelles (Figure 2.5A,
60min). The quantification of colocalization suggested that at 30 min, 62% of viruses
were located in Rab5-positive organelles and 6% in Rab7-positive organelles, and at
60 min, 68% in Rab5-positive organelles and 61% in Rab7-positive organelles. The
colocalization of viral particles with Rab5 was rarely detected at 120 min, but 20%
was observed in Rab7-positive organelles.

47

Figure 2.5: The trafficking of the viral particles through various endosomes. (A) 293T/CD20 cells were
transfected with DsRed-Rab5 or DsRed-Rab7. After 48 h, transfected cells were seeded on the glass
bottom dish overnight. GFP-Vpr-labeled viruses (FUW-GFPVpr/αCD20+SINmu) were incubated with
293T/CD20 cells at 37°C for various time points of 0, 30, 60, or 120 min and then fixed. The boxed
regions are magnified and shown in separated panels below. (B) Colocalization of the viral particle
with transferrin. GFP-Vpr-labeled virus (green) and Alexa647-labeled transferrin (blue) were
incubated with cells at 37°C for 60 or 120 min. Then these cells were fixed, permeabilized, and
immunostained with EEA1 (red). The boxed regions are magnified and shown in separated panels
below. Scale bar represents 5 µm.

48

Next, we compared the trafficking of our engineered lentivirus with
transferrin, which is known to be trafficked from early to recycling endosomes
(Lakadamyali, 2006). Engineered virus (FUW-GFPVpr/αCD20+SINmu) and
Alexa647-conjugated transferrin were incubated with 293T/CD20 cells at 37
o
C. At 60
min, most of the viral particles were colocalized with both EEA1 and transferrin, and
a small fraction (9%) of the viruses were observed in transferrin-positive only
organelles, which could be considered as recycling endosomes (Figure 2.5B, 60 min).
However, at 120 min, 30% of the viruses were colocalized with transferrin-positive,
EEA1-negative organelles, indicating that some fractions of viruses were trafficked to
recycling endosomes (Figure 2.5B, 120 min).

2.2.5 Microtubule-mediated virus transport
We further characterized the endosomal compartment where the viral fusion
occurs by disrupting the microtubules. It has been reported that the microtubule
network can facilitate the migration of viruses to the nucleus and promote the
transport of endosomes in cells (Apodaka, 2001; Mallik, 2004; McDonald, 2002).
Our colocalization study using the tubulin-specific antibody suggested that the
engineered lentiviruses travel along microtubule networks (Figure 2.6A). In order to
examine whether virus-endosome fusion occurs before or after microtubule-
dependent transport, microtubules were damaged by the drug nocodazole.
49
The reduced tubulin-staining confirmed the disruption of microtubule networks
(Figure 2.6B). We next studied the effect of microtubule disruption on virus-
endosome fusion and the confocal image showed that the viral fusion signals could be
detected (Figure 2.6C), and quantification indicated that the viral fusion is
independent of microtubule assistance (Figure 2.7A), suggesting that the viral fusion
with endosomes occurs before microtubule-associated movement.
To further investigate the effects of the microtubule-disrupting drug on the
localization of virus in endosomal compartments, we pretreated cells with nocodazole
for 30 min and quantified the viruses colocalizing with the different endosomal
markers after 60 min of incubation. The results showed that the viral population in
EEA1- and CI-MPR-double positive endosomes in nocodazole-treated cells was
significantly lower than that of untreated cells, whereas the population positive only
for EEA1 was higher in the treated cells (Figure 2.6D and Figure 2.7C). This
indicated that viral particles were mostly located in the early endosomes devoid of
CI-MPR upon nocodazole treatment and that the treatment restricted the further
transport and maturation of early endosomes to EEA1- and CI-MPR-double positive
endosomes. This result suggests that the maturation process of the virus-containing
endosomes is dependent on microtubule-associated transport. The quantification of
the viral fusion (Figure 2.7A) and localization in endosomal compartments (Figure
2.7C) also implies that the late endosomal compartment is not necessary for virus-
endosome fusion and the engineered viral particles are able to initiate fusion at the
early endosome stage.
50
To investigate whether endosome maturation was relevant to productive infection of
our engineered lentiviruses, nocodazole-pretreated cells were transduced by
FUGW/αCD20+SINmu. The FACS analysis showed that the transduction rate
decreased by 31%, suggesting that productive infection was affected markedly by the
nocodazole inhibition of the endosome maturation (Figure 2.7B).
To confirm the role of microtubules in viral infection, we used siRNA to
down-regulate the expression of α-tubulin in cells. 293T/CD20 cells were transfected
with α-tubulin-specific siRNA or control siRNA, and the knockdown of α-tubulin
expression was validated by microtubule staining (Figure 2.6E). The virus-endosome
fusion experiment on the siRNA-treated cells showed that the down-regulation of α-
tubulin did not significantly affect the efficiency of viral fusion (Figure 2.6F and
Figure 2.7A). Similar to the result of the nocodazole treatment, we also observed that
the maturation of the lentivirus-containing endosomes was dependent on microtubule-
associated transport, as most of these endosomes were retained at the early endosome
stage after the siRNA-treatment (Figure 2.6G and Figure 2.7C). As compared to
treatment of the control siRNA, lower transduction was obtained for 293T/CD20 cells
transfected with α-tubulin-specific siRNA and exposed to FUGW/αCD20+SINmu
(Figure 2.7D).

51

Figure 2.6: Microtubule-associated transport of viruses. (A) GFP-Vpr-labeled lentiviruses (FUW-
GFPVpr/αCD20+SINmu) were colocalized with microtubule networks 1 h after infection.
Microtubules were immunostained with the monoclonal antibody to α-tubulin (red). The boxed region
is enlarged in right panel. Arrows indicate viruses on the microtubules. (B) Microtubule staining in a
nocodazole-treated cell. Cells were preincubated with nocodazole at 37°C for 30 min to disrupt
microtubules, then immunostained with the monoclonal antibody to α-tubulin (red). (C) The fixed
image of GFP-Vpr/DiD-labeled viral particles in a nocodazole-treated cell. Cells were preincubated
with nocodazole at 37°C for 30 min, then incubated with GFP-Vpr/DiD-labeled viruses (FUW-
GFPVpr/αCD20+SINmu) at 37°C for 60 min, and then fixed. Arrows indicate the viral particles fused
to endosomes. (D) Localization of GFP-Vpr-labeled viral particles (FUW-GFPVpr/αCD20+SINmu)
with the two endosomal markers after 60 min of incubation in a nocodazole pre-treated cell. (E)
Microtubule staining of siRNA-treated cell. 293T/CD20 cells were transfected with α-tubulin-specific
siRNA- or control siRNA. After 72 h, transfected cells were seeded and immunostained with anti- α-
tubulin antibody (green). (F) The virus fusion for siRNA-treated cells. α-tubulin siRNA or control
siRNA-treated cells were incubated with GFP-Vpr/DiD-labeled viruses at 37°C for 60 min and then
fixed. Yellow particles denoted by arrows indicate the virus particles fused to endosomes. (G)
Localization of GFP-Vpr-labeled viral particles with the two endosomal markers after 60 min of
incubation with α-tubulin siRNA-treated cells. The boxed regions are enlarged and shown in separated
panels. Scale bar represents 5 µm.
52

Figure 2.7: The effects of inhibitory drugs or siRNA treatment on viral fusion, infection, and endosome
maturation. (A) GFP-Vpr/DiD-labeled viruses were incubated with drug- or siRNA-treated cells at
37°C for 60 min, and then fixed. The viral particles with the fusion signal (black) or without the fusion
signal (gray) were quantified. The viral particles both GFP-Vpr
+
and DiD
+
were considered to be fused
with endosomes, while particles that were only GFP-Vpr
+
were considered to be unfused virus. For
quantification, 60 viral particles were examined for no drug-treatment, 64 particles for nocodazole
treatment, 72 particles for α-tubulin siRNA treatment, and 68 particles for cyto-D treatment. The
results were collected from three independent experiments. (B) The role of microtubules and actin
filaments in the virus infection. 293T/CD20 cells which were preincubated with nocodazole or
cytochalasin-D (cyto-D) were transduced with 2 ml of fresh unconcentrated FUGW/αCD20+SINmu
virus. The resulting GFP expression was analyzed by FACS. (C) Quantification of GFP-Vpr-labeled
viruses colocalized with EEA1
+
(black), CI-MPR
+
(white), or both EEA1
+
and CI-MPR
+
(gray)
endosomes at 60 min of incubation in drug- or siRNA-treated cells. (D) The effect of α-tubulin
knockdown on virus infection. Control siRNA or α-tubulin siRNA transfected cells were transduced
with 2 ml of fresh unconcentrated FUGW/αCD20+SINmu virus. The percentage of GFP
+
cells was
analyzed by FACS.

53
2.2.6 Actin-mediated virus transport
We further investigated the role of actin cytoskeleton in trafficking the
engineered viral particles. The colocalization study using rhodamine-conjugated
phalloidin suggested that our engineered lentiviruses travel along actin-filaments
(Figure 2.8A). To characterize actin-mediated virus trafficking, 293T/CD20 cells
were pretreated with 20 µM of cytochalasin D (cyto-D) at 37°C for 30 min to disrupt
actin filaments, and then we examined the viral fusion and virus localization in
endosomal compartments. First, we confirmed the disruption of actin in cyto-D-
treated cells by phalloidin labeling (Figure 2.8B). The viral fusion assay in the cyto-
D-treated cells showed that the majority of viral particles (denoted by arrows) were
fused with the endosomes (Figure 2.7A and Figure 2.8C). The viral population in
endocytic markers showed that viral particles were mostly located in the early
endosomes (Figure 2.7C and Figure 2.8D). Compared to the results of microtubule
disruption by either nocodozale or siRNA, actin-disruption using cyto-D resulted in
greater inhibition of the viral transduction (Figure 2.7B). This further confirmed that
the inhibition of the maturation process of lentivirus-containing endosomes could
affect productive infection.

54

Figure 2.8: Actin-associated transport of viruses. (A) GFP-Vpr-labeled lentiviruses (FUW-
GFPVpr/αCD20+SINmu) were colocalized with actin-filaments 1 h post-infection. Actin filaments
were labeled with rhodamine-conjugated phalloidin (red). The boxed region is enlarged in the right
panel. Arrows indicate viruses on the actin cytoskeleton. (B) Actin stainng in a cytochalasin-D treated
cell. Cells were preincubated with cytochalasin-D at 37°C for 30 min to disrupt actin filaments, and
then labeled with rhodamine-conjugated phalloidin (red). (C) The virus fusion in cytochalasin-D
treated cells. Cells were preincubated with cytochalasin-D at 37°C for 30 min, further incubated with
GFP-Vpr/DiD-labeled viruses at 37°C for 60 min, and then fixed. Arrows indicate the viral particles
fused to endosomes. (D) Localization of GFP-Vpr-labeled viral particles with the two endosomal
markers after 60 min of incubation with cytochalasin-D pre-treated cells. The boxed regions are
enlarged and shown in separated panels. Scale bar represents 5 µm.

2.2.7 Virus release from endosomes
It has been reported previously that Vpr remains largely associated with the
pre-integration complex after fusion (McDonald, 2002); thus, GFP-Vpr labeling can
be used for visualizing the release of the viral core from fused endosomes. GFP-
Vpr/DiD-labeled viruses (FUW-GFPVpr/αCD20+SINmu) were incubated at 37oC for
55
60 min to induce virus-endosome fusion; based on our study of fusion kinetics
(Figure 2.3C), most of the viruses should be retained in the endosomes at this stage.
Confocal time-lapse images were then acquired every ~10 s over a time period of 20
min. At 0 s, the viral particle appeared to be yellow (a merged color of green and red),
indicating that fusion had already taken place (Figure 2.9). At 461 s, the GFP-Vpr-
labeled viral core (green) moved away from the DiD-labeled endosome (red),
suggesting that the virus is being released into the cytosol. We recorded ~50 videos
and 12 of them showed the similar process of virus release illustrated in Figure 6.
This seemingly low yield (24%) could be partially due to photobleaching as a result
of long-time exposure to the laser source during the recording. No virus release signal
was detected for the control virus lacking SINmu (FUW-GFPVpr/αCD20) (Figure
2.10). This live cell imaging suggests that the completion of virus membrane fusion
to release the viral core, which is required for the delivery of viral genome to the cell
nucleus, is relatively long-term and possibly a sequential multi-step process. This
process can be possibly explained by the viral fusion mechanism of hemifusion
(indicated by the dispersion of DiD) and fusion-pore formation and enlargement
(indicated by the separation of the GFP-Vpr and DiD signals), (Jahn, 2003; Melikyan,
2005) although more direct evidence needs to be obtained in order to support this
speculation.

56

Figure 2.9: Time series images of the viral core release from an endosome starting 60 min after
incubation. GFP-Vpr/DiD-labeled viruses (FUW-GFPVpr/αCD20+SINmu) were incubated with
293T/CD20 cells at 37°C for 60 min to initiate virus fusion, and then time-series images were obtained
at every ~10 seconds over a time period of 20 min. Scale bar represents 2 µm.

Figure 2.10: Time series images of the viral core release from an endosome starting 60 min after
incubation. GFP-Vpr/DiD-labeled viruses (FUW-GFPVpr/αCD20) were incubated with 293T/CD20
cells at 37°C for 60 min to initiate virus fusion, and then time-series images were obtained at every
~10 seconds over a time period of 20 min. Scale bar represents 2 µm.

57
2.3 Discussion
In this study, we have visualized individual engineered lentiviruses and have
analyzed the intracellular behavior of viruses in the target cells using single viral
particle tracking via confocal microscope. GFP-Vpr labeling along with other
immunostaining methods has allowed us to confirm the co-incorporation of an
antibody and fusogenic molecule on a single virion, separating the functions of
binding and fusion, respectively. We have confirmed by the direct visualization of
virus-host cell interactions that the antibody displayed on the virion surface targets a
specific cell type and we have shown that viral particles were efficiently endocytosed
upon antibody binding. Subsequently, the surface-displayed fusogenic molecule,
SINmu, (Yang, 2006) which is an engineered glycoprotein derived from the Sindbis
virus that is binding-deficient and fusion-competent, mediated fusion in the acidic
endosomal environment. By observing the trafficking of single lentivirus
intracellularly, we have shed some light on the endocytic mechanisms of engineered
lentivirus such as virus-endosome fusion and endosomal transport of viral particles in
target cells.
By using a fluorescence de-quenching assay, we have detected the actual
fusion event of the GFP-Vpr/DiD double-labeled viral particle and monitored the
dynamics of fusion in living cells. Colocalization studies using endocytic markers
suggested that the endosomal fusion of the engineered lentivirus started at an early
stage of the endocytic pathway. Inhibitory drug treatments showed that the transport
58
and maturation steps of the virus-endosome complex from the early to the
intermediate endosome require microtubule- or actin-assisted transport. However,
virus-endosome fusion was not restricted by the inhibition of microtubule- or actin-
associated transport, suggesting that the majority of viral fusion begins at the early
endosome level.
We also studied the fusion kinetics of the engineered lentivirus and compared
it to VSVG-pseudotyped lentivirus, which is known to require low pH to mediate
fusion (Sun et al., 2005). It has been shown that the fusion of VSV occurs before the
late endosome stage (Sieczkarski, 2003). Our data suggests that it takes 70~80 min
more for the engineered lentivirus to reach a half-maximum of fusion-involved
penetration than it does for VSVG-pseudotyped virus. We suspect that one possible
reason for this delay could be due to the separation of the functions of binding and
fusion into two individual molecules on our targeting lentivirus system. The different
requirements for fusion of different viruses could be another cause for the slower
entry kinetics of the engineered lentivirus. For example, the Sindbis virus requires
cholesterol for the endosomal fusion, while the fusion of VSV is independent of
cholesterol (Lu et al., 1999). We are conducting experiments to investigate the effects
of such membrane components on the entry efficiency of the engineered lentivirus.
Recently, several attempts have been made to understand the role of
endosomes in the intracellular trafficking of virus. It was demonstrated by using
dominant-negative Rab5 and Rab7 that SFV and VSV are capable of fusing at the
early endosome level, whereas the influenza virus requires the involvement of the late
59
endosome for infection (Sieczkarski, 2003). A colocalization assay using endosomal
markers suggested an endocytic pathway of SFV in which the virus is transported via
early endosomes to late endosomes and lysosomes (Vonderheit, 2005). By combining
this colocalization with a neutralization assay using NH
4
Cl, it was found that
productive SFV fusion occurred when the virus was still in early endosomes
(Vonderheit, 2005). Additionally, it was proposed that Rab7, a late endosome marker,
is recruited to early endosomes to mediate the sorting of SFV to late endosomes
(Vonderheit, 2005). Our results for the engineered lentivirus displayed a similar
pattern of virus-endosome fusion. Moreover, our approach using the GFP-Vpr/DiD
double-labeling method has enabled us to directly visualize the fusion and viral
release of the engineered lentivirus.
Using the real-time imaging of single viruses, we were able to observe the
dynamics of virus-endosome fusion and transport. After the viral fusion signal was
seen, we observed that many virus-containing endosomes (60%) moved towards the
nucleus. Since the lipophilic membrane dye is initially incorporated onto the outer
membrane of the virus, the lipid mixing (i.e. hemifusion) between the viral outer
membrane and the endosomal inner membrane is initially visualized by the increase
of DiD-fluorescence. The fusion-pore formation and enlargement of the pore is
believed to be required for the completion of the virus membrane fusion in order to
release the viral core into the cytosol (Markosyan, 2005; Melikyan, 2005). Our live-
cell imaging of viral release seems to support this hypothesis. We observed that the
separation of the viral core (labeled with GFP-Vpr) from the endosome (labeled with
60
DiD) followed a slow and possibly multi-step process, as indicated the time interval
between the appearance of colocalized GFP-Vpr and DiD signals (fusion) and the
separation of these signals (virus release). From the experiments of the drug and
siRNA treatments (Figure 2.7A), we have learned that the hemifusion (i.e. lipid
mixing) takes place before the late endosome stage. The reduced infectivity upon the
drug and siRNA treatments (Figure 2.7B & 2.7D) indicates that the release of the
viral core, which is a necessary step for a productive infection, is associated with the
maturation process from the early to the intermediate endosomal compartments. This
suggests that the virus is at least transported to the intermediate endosome to
complete infection.
In summary, we demonstrated that the direct visualization of individual
viruses can allow us to determine the endocytic pathway of the engineered lentivirus
in living cells. The single-virus tracking techniques confirmed our hypothesis of the
mechanism of targeted transduction for the engineered lentivirus: the surface-
displayed antibody directs the virus to a specific cell type, and this binding efficiently
induces the endocytosis of the viral particle, bringing it into an endosome; the
fusogen then mediates viral membrane fusion at the low pH environment of
endosomes, allowing the viral core to enter the cytosol. Our observations, based on
the kinetics of entry of engineered lentivirus, indicate that the fusogen-mediated
membrane fusion is a long process, which is significantly slower than that of VSVG-
pseudotyped lentivirus, suggesting that this might be the rate limiting step of virus
transduction. Therefore, the development of fusogenic molecules which induce more
61
efficient and stable fusion could be a key step in enhancing the lentivirus-mediated
gene delivery efficacy.

2.4 Materials and Methods
2.4.1 Cell lines and antibodies
The 293T/CD20 cell line was generated previously. (Yang, 2006) 293T and
293T/CD20 cells were maintained in a 5% CO2 environment in Dulbecco’s modified
Eagle medium (Mediatech, Inc.) with 10% FBS (Sigma), and 2 mM L-glutamine
(Hyclone). Mouse monoclonal antibody specific to the human CD20 antigen were
purchased from Caltag Laboratories. Anti-HA-biotin to stain SINmu was obtained
from Miltenyi Biotec Inc. Mouse monoclonal antibody against the early endosome
antigen 1 (EEA1), rabbit polyclonal antibody specific to the Mannose 6 phosphate
receptor (CI-MPR), and Cy5-conjugated goat anti-rabbit IgG antibody were
purchased from Abcam. Cy5-conjugated streptavidin was purchased from Zymed
Laboratories. Texas red-labeled goat anti-mouse IgG and AlexaFluor 594-labeled
goat anti-human IgG antibodies were obtained from Molecular Probes. Taxol,
nocodazole, cytochalasin D, and bafilomycin A1 were purchased from Sigma.

62
2.4.2 Plasmids
Assembly PCR was employed to fuse the GFP to the N-terminus of viral
protein R (Vpr). The PCR product was then inserted into pcDNA3 (Invitrogen) to
form pcDNA3-GFPVpr. The cDNAs for Rab5 and Rab7 were PCR-amplified and
cloned into the pDsRed-monomer-C1 (Clontech) to form DsRed-Rab5 and DsRed-
Rab7, respectively.

2.4.3 Virus production
GFP-Vpr-labeled lentiviruses were produced by transfecting 293T cells by a
calcium phosphate precipitation method. 293T cells at 80% confluence in 6 cm
culture dishes were transfected with 5 µg of the lentiviral plasmid FUW, together
with 2.5 µg each of pcDNA3-GFPVpr, pαCD20 (encodes a mouse/human chimeric
anti-CD20 antibody), pIgαβ (encodes human Igα and Igβ, two immunoglobulin
associated proteins that are required for surface expression of antibodies), pSINmu,
and the packaging vector plasmids.(Yang, 2006) Cells were washed at 4 h
posttransfection, and then medium was replaced. The viral supernatant was collected
after 48 h posttransfection and filtered through a 0.45-µm pore size filter. For high-
titer lentivectors, the viral supernatant was concentrated by ultracentrifugation
(Optima L-90 K ultracentrifuge, Beckman Coulter) for 90 min at 82,700 × g and
resuspended in an appropriate volume of Hank’s balanced salt solution (HBSS,
Hyclone).
63
2.4.4 Viral transduction
For viral transduction, 293T/CD20 cells (0.2 × 106 per well) were plated in a
24-well culture dish and spin-infected with viral supernatants of
FUGW/αCD20+SINmu (2 ml per well) at 2,500 rpm and 30°C for 90 min by using a
Sorval Legend centrifuge. Then, the medium was removed and replaced with fresh
medium and cultured for 3 days further before FACS analysis of GFP+ cells. For
viral transduction with drug- or siRNA-treated cells, 293T/CD20 cells were
preincubated with drugs (nocodazole (60 µM), cytochalasin D (20 µM), and
bafilomycin A1 (25 nM)) or transfected with siRNAs, and then the cells (0.2 × 106
per well) were spin-infected with 2 ml of viral supernatants in a 24-well culture dish.

2.4.5 Time course study of fusion inhibition
293T/CD20 cells (0.4 × 105 per well) were seeded in a 96-well culture dish
overnight. Viruses were added to cells and incubated at 4°C for 1 h and the resulting
cells were washed with cold PBS to remove unbound viruses. The viral entry was
initiated by shifting cells to 37°C. At the indicated time points, D10 media containing
25 nM of bafilomycin A1 was added to inhibit the endosome acidification. The drug
was removed 3 h later and replaced with fresh D10 media. Cells were further
incubated for 72 h, and the percentage of GFP-positive cells was analyzed by FACS.

64
2.4.6 Fluorescent labeling
For the detection of individual viral particles, fresh viral supernatant was
overlaid upon poly-lysine-coated NO.1 coverslips in a six-well culture dish and
centrifuged at 3,700 × g and 4°C for 2 h in a Sorval Legend RT centrifuge. The
coverslips were rinsed with cold PBS twice, the adhered viruses were immunostained
by Alexa 594 anti-human IgG and anti-HA-biotin antibodies, and they were then
incubated with Cy5-streptavidin. The coverslips were mounted in Vectashield (Vector
Laboratories), which is an antifade mounting medium. For imaging virus-cell
binding, 5 × 105 cells were seeded onto a 35 mm glass-bottom culture dish (MatTek
Corporation) and grown at 37°C overnight. The seeded cells were rinsed with cold
PBS twice and incubated with the concentrated viruses for 1 h at 4°C to allow for
binding. Cells were washed with cold PBS to remove unbound viruses and then fixed
with 4% formaldehyde on ice for 10 min. To co-label the viral particles bound to the
cell surfaces, Alexa594 anti-human IgG antibody was used to stain against the αCD20
heavy chain. Fluorescent images were taken by a Zeiss LSM 510 laser scanning
confocal microscope equipped with filter sets for fluorescein, rhodamine or Texas
red, and Cy5. A plan-apochromat 63x/1.4 oil immersion objective was used for
imaging. Images were analyzed with the use of the Zeiss LSM 510 software version
3.2 SP2.

65
2.4.7 Imaging virus fusion and transport through endosomes
The concentrated viruses were incubated with 100 µM of 1,1'-dioctadecyl-
3,3,3',3'-tetramethylindodicarbocyanine (DiD) (Molecular Probes) for 1 h at room
temperature. For imaging virus-endosome fusion, double labeled viruses were
incubated with 293T/CD20 cells at 37°C for various time periods and then fixed with
4% formaldehyde. GFP-Vpr and DiD were excited simultaneously with a 488 nm
Argon and a 633 nm HeNe laser, respectively, and the emitted light was separated
through the corresponding emission filter sets. All samples were scanned under the
same conditions for magnification, laser intensity, brightness, gain, and pinhole size.
For live cell imaging of virus fusion, 293T/CD20 cells were preincubated on a glass
bottom culture dish at 37°C overnight. Viruses were incubated with the cells at 37°C
for 30 min to initiate virus internalization. Images were then collected using the
confocal microscope. Fluorescence intensity vs. time within the regions of interest
around the virus particles were measured by using the Zeiss LSM 510 software
package.
For observation of the colocalization of the virus with different endosomal
markers, GFP-Vpr-labeled viruses were incubated with 293T/CD20 cells at 37°C and
fixed after different time periods. Cells were then permeabilized with 0.1% Triton X-
100 and immunostained with EEA1 and CI-MPR for early endosome and late
endosome markers, respectively. Texas red-conjugated anti-mouse IgG and Cy5-
conjugated goat anti-rabbit IgG antibodies were used as the secondary antibodies. To
66
remove viral aggregates, virus-containing media was filtered by a 0.45 µm pore size
centrifuge tube filter (Costar, NY) just before the experiments were conducted.
For the viral trafficking studies using Rab5 and Rab7 constructs, 293T/CD20
cells were transfected with DsRed-Rab5 or DsRed-Ran7 plasmid. At 48 h
posttransfection, cells were seeded onto the glass-bottom culture dish and grown at
37°C overnight. GFP-Vpr-labeled viruses at MOI ~30 were added and incubated at
37°C for the different time points and then fixed.
For the colocalization study with transferrin, GFP-Vpr-labeled viruses and 2 µM of
Alexa647-conjugated transferrin (Molecular Probes) were mixed and then incubated
with 293T/CD20 cells at 37°C for 60 or 120 min. Cells were fixed, permeabilized,
and immunostained with EEA1 to compare trafficking of the engineered lentivirus
with transferring through early to recycling endosomes.

2.4.8 Microtubule- and actin-mediated transport
To observe viruses on microtubule networks, the cells were incubated with
GFP-Vpr-labeled viruses at 37°C for 1 h, fixed, and then permeabilized with 0.1%
Triton X-100 containing 20 µM of taxol. Microtubules were then immunostained
with anti-α-tubulin mAb (Sigma) and Texas red-conjugated anti-mouse secondary
antibody. For the microtubule-disrupting assay, cells were preincubated with D10
media containing 60 µM of nocodazole at 37°C for 30 min, and then viruses were
added and incubated for further studies.
67
For the inhibition assay using small interfering RNA (siRNA), we purchased
α-tubulin siRNA and the negative control siRNA from Santa Cruz Biotechnology,
Inc. The transfection of siRNA was performed as described by the manufacture’s
protocol. At 72 h posttransfection, equal numbers of α-tubulin siRNA and control
siRNA treated 293T/CD20 cells were used for further studies.
To visualize viruses on actin filaments, the cells were incubated with GFP-
Vpr-labeled viruses at 37°C for 1 h, fixed, and permeabilized. Actin filaments were
labeled with rhodamine-conjugated phalloidin (Molecular Probes). For the actin-
disrupting assay, cells were preincubated with D10 media containing 20 µM of
cytochalasin-D at 37°C for 30 min, and then viruses were added and incubated for
further studies.

2.4.9 Live cell imaging of viral release
For the real-time observation of the release of the virus from the endosome,
GFP/DiD-labeled viruses were incubated with cells at 37°C for 1 h to induce virus-
endosome fusion, and confocal time-lapse images were then recorded.

68
Chapter 3. IMAGING MULTIPLE INTERMEDIATES OF SINGLE-
VIRUS MEMBRANE FUSION MEDIATED BY DIVERSE FUSION
PROTEINS

Portions of this Chapter are adapted from:
Kye-Il Joo, April Tai, Chi-Lin Lee, Madhavi Srisha, Clement Wong, and Pin Wang.
(Submitted 2009)

The merging of viral and target cell membranes is catalyzed by viral fusion
proteins, which involves sequential multiple steps in the fusion process. However, the
fusion mechanisms mediated by different fusion proteins involve multiple transient
intermediates that have not been well characterized. Here we report a synthetic virus
platform that allows us to better understand the different fusion mechanisms driven
by the diverse types fusion proteins. The platform consists of lentiviral particles co-
enveloped with a surface antibody, which serves as the binding protein, along with a
fusion protein derived from either influenza virus (HAmu) or Sindbis virus (SINmu).
By using a single virus tracking technique, we demonstrated that both HAmu and
SINmu viruses enter cells through clathrin-dependent endocytosis, but they required
different endosomal trafficking routes to initiate viral fusion. Direct observation of
single viral fusion events clearly showed that hemifusion mediated by SINmu upon
exposure to low pH occurs faster than that mediated by HAmu. Monitoring sequential
fusion processes by dual labeling the outer and inner leaflets of viral membranes also
69
revealed that the SINmu-mediated hemifusion intermediate is relatively long-lived as
compared with that mediated by HAmu. Taken together, we have demonstrated that
the combination of this versatile viral platform with the techniques of single virus
tracking can be a powerful tool for revealing molecular details of fusion mediated by
various fusion proteins.

3.1 Introdution
Fusion of the two lipid bilayers is a key step for enveloped viruses to
successfully infect target cells. Although merging the two membranes is
thermodynamically favorable, the fusion reaction is hampered by a huge energy
barrier (Chernomordik and Kozlov, 2003; Chernomordik et al., 2006; Harrison,
2008). Viral fusion proteins can lower this barrier through the activation energy
provided by their conformational change to reach an energetically stable state (Carr et
al., 1997; Dimitrov, 2004; Hogle, 2002; Mothes et al., 2000). It is generally believed
that virus membrane fusion proceeds through sequential and multiple steps
(Chernomordik and Kozlov, 2003; Dimitrov, 2004; Harrison, 2008; Jahn et al., 2003;
Kielian and Rey, 2006; Kuhn et al., 2002). Initially, viral fusion proteins undergo a
conformational change induced by receptor-binding and/or a pH change, which yields
a point of contact between the viral and the target membranes. This induces the
process of hemifusion (or lipid mixing), in which the proximal leaflets of two distinct
70
lipid membranes are merged while the distal leaflets remain unchanged. The
subsequent merging of the distal leaflets causes fusion pore formation (or content
mixing), which establishes the first aqueous connection between the two apposing
membranes (Jackson and Chapman, 2008; Lentz et al., 2000; Zimmerberg et al.,
1994). This general fusion reaction scheme is relatively well documented, but the
detailed mechanisms and pathways underlying each fusion step, which may vary
significantly among the diverse types of viral fusion proteins, have been less well-
studied.
Viral fusion proteins are currently classified into two groups, based on their
structural features. Class I fusion proteins, such as the influenza hemagglutinin (HA)
protein and the HIV-1 fusion protein (gp120/gp41), are trimeric structures in both
pre-fusion and post-fusion conformations and trigger fusion using α-helical coiled-
coil domains (Dimitrov, 2004; Harrison, 2008). Class II fusion proteins, such as the
E1 protein of alphaviruses like the Semliki Forest virus (SFV) and Sindbis virus
(SIN), need to convert from dimeric to trimeric structures for fusion, induced by β-
barrel structures, to occur (Dimitrov, 2004; Kielian and Rey, 2006). The influenza
and Sindbis viruses have been commonly studied as models of class I and II fusion
proteins to understand the fusion mechanisms of the two different classes of proteins
using both cell-cell (Blumenthal et al., 1996; Chernomordik et al., 1998; Mittal et al.,
2003; Zaitseva et al., 2005) and virus-liposome fusion assays (Shangguan et al., 1996;
Smit et al., 1999; Smit et al., 2001; Smit et al., 2002; White et al., 1982).
71
These conventional fusion assays have been useful for understanding the viral
membrane fusion process, but they cannot be employed to study the kinetics of single
viral fusion events or monitor the series of transient fusion intermediates formed
during single virus fusion with target cell membranes. Direct visualization of the viral
fusion process can expand our ability to reveal molecular details of virus fusion. The
observation of the actual fusion event and its kinetic characterization at the single
virus level can provide crucial understanding of the underlying mechanisms of virus
infection. One complication of comparing the fusion reactions of different viruses in
living cells is the differences in their receptor bindings, which may subsequently
result in different intracellular viral trafficking mechanisms being involved in the
fusion process. For example, the influenza virus HA1 protein has receptor-binding
specificity to sialic acid (Skehel and Wiley, 2000), while the E2 protein of Sindbis
virus has binding recognition to heparan sulfate structures on the cell surface (Byrnes
and Griffin, 1998).
In this study, we test a virus platform for the study of molecular fusion
mediated by the different types of fusogens. This platform is based on a lentivirus
designed to incorporate a cell binding and a cell fusion protein as two separate
molecules on the same viral particle (Fig. 3. 1A). We have recently reported such a
virus system displaying a CD20-specific surface antibody (αCD20) as the binding
molecule and a binding-deficient fusion-competent glycoprotein as the fusogen (Yang
et al., 2006). This recombinant virus could achieve targeted transduction of CD20-
expressing cells in vitro and in vivo. We have also visualized the late stages of
72
intracellular tracking and fusion of this virus (Joo and Wang, 2008). Beyond the
application for targeted gene delivery, we hypothesize that this virus system can be
used for the comparative study of different fusogen-mediated viral fusion. We
envision that such a system would offer an opportunity to directly compare the fusion
processes of various fusogens by allowing the production of viruses with the same
binding proteins but different fusogens. Such resulted viruses would undergo the
same pathway of initial internalization induced by the interaction between the binding
protein and the target receptor. The present study is to test this hypothesis by
investigating the fusion properties of two fusion proteins: one is the class I fusogen
derived from influenza virus hemagglutinin (designated as HAmu) and the other is
the class II fusogen derived from Sindbis virus glycoprotein (designated as SINmu).
The single virus tracking study of the early internalization process indicates that both
HAmu- and SINmu-lentiviruses enter cells through clathrin-dependent endocytosis.
This study further identifies the different requirements of endosomal trafficking for
the membrane fusion of these two lentiviruses. The planar fusion assay utilizing dual
labeling of outer and inner leaflets of viral membranes allows us to reveal the
different kinetics of hemifusion and fusion pore formation triggered by these two
fusogens in living cells.

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3.2 Results
3.2.1 Engineered lentiviruses require low pH to trigger endosomal fusion
To image the intracellular trafficking of individual engineered lentiviruses
displaying different fusion proteins (HAmu or SINmu) within target cells, we
constructed a labeling protein GFP-Vpr by fusing green fluorescent protein (GFP)
with the HIV accessory protein Vpr (viral protein R). GFP-Vpr-labeled lentiviruses
enveloped with αCD20 along with HAmu (FUW-GFPVpr/αCD20+HAmu) or SINmu
(FUW-GFPVpr/αCD20+SINmu) were produced as described (Joo and Wang, 2008;
Yang et al., 2006). To examine whether both types of engineered lentiviruses entered
target cells via endocytosis, GFP-Vpr-labeled lentiviruses were incubated with
293T/CD20 cells on ice for 30 min to synchronize infection. Cells were then warmed
to 37ºC for 30 min and immunostained against early endosome antigen 1 (EEA1), an
early endosomal marker (Lakadamyali et al., 2006; Sieczkarski and Whittaker, 2003;
Vonderheit and Helenius, 2005). As shown in Figure 3.1B and 3.1C, most of viral
particles were colocalized with EEA1, as evidenced by the appearance of yellow
color in the merged images, suggesting that both viruses were located in the EEA1+
endosomes. To further demonstrate that a low pH compartment is required to allow
engineered lentiviruses to penetrate into cells, bafilomycin A1, a specific inhibitor of
vacuolar proton ATPases, was used to block low pH endosomal fusion (Bowman et
al., 1988). To assay the transduction, we replaced the lentiviral backbone plasmid
FUW with FUGW, which carries a GFP reporter gene under the control of the human
ubiquitin-C promoter (Lois et al., 2002), in our transfection protocol to prepare
74
viruses. 293T/CD20 cells were pretreated with bafilomycin A1 at different
concentrations and transduced with lentiviruses bearing αCD20 and a fusion protein
(FUGW/αCD20+SINmu or FUGW/αCD20+HAmu), VSVG-pseudotyped lentivirus
(FUGW/VSVG), or VSVG-pseudotyped gamma-retrovirus (MIG/VSVG). Viruses
pseudotyped with vesicular stomatitis virus glycoprotein (VSVG) are known to enter
cells through pH-dependent endocytosis (Sieczkarski and Whittaker, 2003; Sun et al.,
2005) and were therefore utilized as a positive control. The FACS analysis of GFP+
cells showed that the viral transduction was inhibited in a dose-dependent manner
(Figure 3.1D), indicating that both types of engineered lentiviruses require low-pH-
triggered endosomal fusion to complete transduction.

3.2.2 The entry of engineered lentiviruses is clathrin-dependent
Clathrin- and caveolin-mediated endocytosis have been observed in the
internalization of many viruses into cells (Blanchard et al., 2006; DeTulleo and
Kirchhausen, 1998; Doxsey et al., 1987; Kirchhausen, 2000; Nabi and Le, 2003;
Nichols and Lippincott-Schwartz, 2001; Rust et al., 2004). To investigate the role of
clathrin- or caveolin-mediated endocytosis in the entry of engineered lentiviruses
bearing HAmu or SINmu fusion proteins to target cells, we visualized the individual
lentiviral particles (FUW-GFPVpr/αCD20+HAmu or FUW-
GFPVpr/αCD20+SINmu) and endocytic structures (clathrin or caveolin) in target
cells after different incubation time periods (0, 10, 30 min).

75

Figure 3.1: Engineered lentiviruses can enter target cells via endocytosis. (A) Schematic representation
of a proposed entry mechanism for engineered lentiviruses enveloped with a CD20-specific surface
antibody (αCD20) and a fusion protein (HAmu or SINmu). (B and C) GFP-Vpr-labeled viruses (green)
enveloped by αCD20 and either SINmu (FUW-GFPVpr/αCD20+SINmu, B) or HAmu (FUW-
GFPVpr/αCD20+HAmu, C) were added to 293T/CD20 cells at 4ºC for 30 min to synchronize the
binding. The cells were then warmed to 37ºC for 30 min, fixed, permeabilized, and immunostained
against the early endosome antigen 1 (EEA1; red). The boxed regions are enlarged in the right panels.
Scale bar represents 5 µm. (D) The effect of the vacuolar proton ATPases inhibitor bafilomycin A1 on
viral infection. 293T/CD20 cells (2 × 10
5
) were pretreated for 30 min with 0, 25, and 50 nM of
bafilomycin A1. The cells were then spin-infected with supernatants of lentiviruses bearing αCD20
and fusogenic protein (SINmu: FUGW/αCD20+SINmu, or HAmu: FUGW/αCd20+HAmu), VSVG-
pseudotyped lentivirus (FUGW/VSVG), or VSVG-pseudotyped gamma-retrovirus (MIG/VSVG).
After 1.5 h of spin-infection and the additional 3 h of incubation in the presence of the drug, the cells
were washed with PBS and resupplied with fresh media. The percentage of GFP
+
cells was analyzed
by FACS.

76
As shown in Figure 2A and 2C, significant colocalization of SINmu-bearing viruses
(67%, n=59) or HAmu-bearing viruses (61%, n=64) with the discrete clathrin
structures was detected after 10 min of incubation. After incubation for 30 min, less
colocalization of SINmu-viruses (20%, n=71) or HAmu-viruses (19%, n=82) with the
clathrin structures was observed, suggesting that many of the viruses had already
been dissociated from uncoated clathrin vesicles and had likely been transported to
early endosomes. However, no significant colocalization of engineered lentiviruses
and caveolin was observed during these time periods (Figure 3.2B and 3.2D),
suggesting that caveolin might not be involved in the entry of these engineered
lentiviruses. Thus, these time-course images showed that both types of engineered
lentiviruses enter target cells through clathrin-mediated endocytosis.
To confirm the results observed from confocal imaging of the individual viral
particles with clathrin or caveolin, we examined the effect of inhibitory drugs on viral
entry (Figure 2E and 2F). Chlorpromazine is a drug known to prevent clathrin
polymerization and inhibit internalization mediated by clathrin-coated vesicles (CCV)
(Wang et al., 1993), while filipin is a cholesterol-binding reagent that blocks
caveolin-dependent internalization (Orlandi and Fishman, 1998). The FACS analysis
showed that the entry of both types of lentiviruses (FUGW/αCD20+HAmu or
FUGW/αCD20+SINmu) was markedly inhibited by the chlorpromazine treatment
(Figure 2E), whereas no inhibitory effect of filipin was observed (Figure 3.2F),
indicating that the entry of both types of viruses is dependent on clathrin.
77
We found that VSVG-pseudotyped gamma-retrovirus (MIG/VSVG) enters cells via
the clathrin-dependent route (Figure 3.2E), which is in agreement with previous
reports (Blanchard et al., 2006; Lee et al., 1999) and is consistent with the endocytic
nature of the vesicular stomatitis virus (VSV) (Sun et al., 2005). However, no
inhibitory effect by chlorpromazine was observed for the entry of the VSVG-
pseudotyped lentivirus (FUGW/VSVG) (Figure 3.2E), which is surprising but
consistent with previous biochemical studies by us and others (Daecke et al., 2005;
Joo et al., 2008). Overall, this set of drug inhibition studies confirmed the results of
the imaging study that the entry of both types of engineered lentiviruses relies on a
clathrin-dependent route.

78

Figure 3.2: Clathrin/caveolin-dependent entry of engineered lentiviruses. GFP-Vpr-labeled lentiviruses
(green) enveloped with αCD20 and SINmu (A and B) or HAmu (C and D) were incubated with
293T/CD20 cells at 4ºC for 30 min. The cells were warmed to 37ºC for various time periods (0, 10,
and 30 min), fixed, permeabilized, and immunostained with anti-clathrin (A and C; red) or anti-
caveolin-1 (B and D; red) antibodies. The boxed regions are magnified and shown below in individual
panels. Scale bar represents 5 µm. Inhibition of clathrin-dependent internalization by chlorpromazine
(E) or caveolin-dependent internalization by filipin (F). 293T/CD20 cells were preincubated with
chlorpromazine or filipin for 30 min at 37ºC. The cells (2 × 10
5
) were then spin-infected with
supernatants of lentiviruses (FUGW/αCD20+SINmu, FUGW/αCd20+HAmu, FUGW/VSVG, or
MIG/VSVG). Both drug concentrations were maintained during the spin-infection as well as for the
additional 3 h incubation, after which the drug was removed and replaced with fresh media. The
percentage of GFP
+
cells was analyzed by FACS.

79
3.2.3 Different fusion proteins modulate different kinetics of virus-endosome fusion
To compare the endosomal fusion kinetics of engineered lentiviruses bearing
different fusion proteins (SINmu or HAmu), we visualized the actual fusion event of
internalized viruses within endosomes by labeling viruses with a lipophilic dye, 1,1’-
dioctadecyl-3,3,3’,3’-tetramethylindodicarbocyanine (DiD). The spontaneous
incorporation of DiD into the viral membranes of GFP-Vpr-tagged lentiviral particles
offers a double-labeling scheme (Joo and Wang, 2008). Viral fusion with endosomal
membranes can be detected by fluorescent dequenching of DiD (Joo and Wang, 2008;
Lakadamyali et al., 2003; Sakai et al., 2006). GFP-Vpr/DiD-labeled viruses were
initially added to 293T/CD20 cells in the cold to synchronize binding. Cells were then
incubated at 37ºC for different time periods (0, 30, 60, and 90 min), and then fixed.
The acquired images with both green and red fluorescent signals are shown in Figure
3.3A. At 30 min of incubation, SINmu-bearing viruses (FUW-
GFPVpr/αCD20+SINmu) were fused with endosomes, as indicated by the appearance
of the red signal (Figure 3.3A, upper, 30 min). However, although HAmu-bearing
viruses (FUW-GFPVpr/αCD20+HAmu) also appeared to be endocytosed, the
majority of the viral particles were detected solely by the green signal, suggesting that
fusion had not yet occurred (Figure 3.3A, lower, 30 min). The images obtained after
60 min of incubation showed that many HAmu-bearing viral particles were fused
with endosomes (Figure 3.3A, lower, 60 min). After 90 min of incubation, pure green
signals were seen for some of the GFP-Vpr-carrying viral particles bearing SINmu,
suggesting that certain viral particles were presumably released into the cytosol after
80
endosomal fusion (Figure 3.3A, upper, 90 min), whereas significant colocalization of
HAmu-displaying viruses and fusion signals remained (Figure 3.3A, lower, 90 min).
We quantified the endosomal fusion of the engineered viruses by viewing
more than 100 viral particles for each time point and counting the GFP-Vpr-labeled
viruses colocalized with or without the DiD signal (Figure 3.3B). This quantification
method showed that the virus-endosome fusion of SINmu-bearing viruses peaked at
30 min after incubation. This fusion event was maintained for up to 60 min of
incubation, but decreased significantly between 90 and 120 min of incubation. The
lower percentage of colocalization might be attributed to the release of viral particles
from the endosomes. On the other hand, the fusion of HAmu-bearing viruses peaked
at 60 min after incubation, and the colocalization of GFP-Vpr with the fusion signal
(DiD) remained at the same level for up to 120 min of incubation. The results of this
direct visualization of viral fusion indicated that fusion of SINmu-bearing viruses
occurs faster than that of HAmu-bearing viruses.
This quantification study also showed that the fusion yield at peak time points for
SINmu-viruses (~ 80%) is higher than that of HAmu-viruses (~ 60%), and that the
release of SINmu viral cores from fused endosomes into the cytosol seems to occur
earlier than that of HAmu viral cores.

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Figure 3.3: Visualization of virus-endosome fusion at different time points for engineered lentiviruses
displaying SINmu or HAmu. (A) GFP-Vpr-labeled engineered lentiviruses (FUW-
GFPVpr/αCD20+SINmu or FUW-GFPVpr/αCD20+HAmu; green) were labeled with DiD (red) for 1 h
at room temperature. Double-labeled viruses were added to 293T/CD20 cells at 4ºC for 30 min to
synchronize the binding. The cells were then warmed to 37ºC for 0, 30, 60, or 90 min, then fixed and
imaged. The boxed regions are magnified and shown below in individual panels. Yellow signals
indicate viral particles fused with endosomes. Scale bar represents 5 µm. (B) Quantification of the
fused viral particles at different incubation time periods. GFP-Vpr+ viral particles with the fusion
signal were quantified by viewing more than 100 viral particles at each time point. The viral particles
positive for both GFP-Vpr and DiD were considered to be fused with endosomes, whereas particles
that were only GFP-Vpr+ were considered to be unfused viruses. The results were collected from three
independent experiments.
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3.2.4 Expression of dominant-negative Rab7 inhibits the transduction of HAmu-
bearing viruses
Viruses that enter cells in a pH-dependent manner are believed to fuse in
either early or late endosomes for a successful infection. To determine which of these
compartments are necessary for the productive transduction of engineered
lentiviruses, the dominant-negative mutants of Rab proteins were used to disable
either the early, Rab5, (Stenmark et al., 1994) or the late, Rab7, endosome function
(Press et al., 1998). 293T/CD20 cells transfected with either the wild-type or
dominant-negative form of Rab5 or Rab7 were incubated with
FUGW/αCD20+SINmu, FUGW/αCD20+HAmu, FUGW/VSVG, or MIG/VSVG.
Expression of the Rab5 dominant-negative mutant reduced the transduction rate of
SINmu- or HAmu-bearing viruses by 60~70% as compared to the transduction of
wild-type Rab5-expressing cells (Figure 3.4A), suggesting that the engineered
lentiviruses must be trafficked through early endosomes for a successful transduction.
The transduction by VSVG-pseudotyped gamma-retrovirus (MIG/VSVG) was
inhibited by the over-expression of dominant-negative Rab5 (Figure 3.4A), consistent
to the notion that VSVG-pseudotyped viruses are internalized into low-pH endosomes
to infect target cells. However, over-expression of the Rab5 mutant did not result in
significant inhibition to the transduction of 293T/CD20 cells by VSVG-pseudotyped
lentivirus (Figure 4A). This result is somewhat unexpected, although a similar
observation was reported previously (Vidricaire and Tremblay, 2005).
83
As shown in Figure 3.4B, expression of the dominant-negative Rab7 mutant
resulted in significant transduction inhibition of HAmu-bearing viruses, indicating
that the HAmu virus requires a functional late endosome for infection. Expression of
mutant Rab7 also blocked transduction by SINmu-bearing viruses, although to a
lesser degree than HAmu-bearing viruses, suggesting that infection by the SINmu
virus is also associated with late endosomal trafficking. We found that the entry of
VSVG-pseudotyped viruses (FUGW/VSVG or MIG/VSVG) was not affected by the
expression of the dominant-negative Rab7 mutant, which is not too surprising
because VSV is known to fuse in early endosomes (Sieczkarski and Whittaker, 2003).

84

Figure 3.4: Inhibition of virus entry by Rab5 and Rab7 dominant-negative mutants. (A) 293T/CD20
cells transiently transfected with wild-type or dominant-negative mutant Rab5 were spin-infected with
various lentiviruses (FUGW/αCD20+SINmu, FUGW/αCd20+HAmu, FUGW/VSVG, or MIG/VSVG).
(B) 293T/CD20 cells transiently transfected with wild-type or dominant-negative mutant Rab7 were
spin-infected with different viruses as indicated above. The percentage of GFP+ cells was analyzed by
FACS.

85
3.2.5 Lentivirus displaying the fusion protein HAmu, but not SINmu, requires
functional late endosome trafficking for viral fusion
To further characterize the endosomal compartment where fusion of the
engineered lentiviruses occurs, the individual fusion events were imaged with
fluorescent protein-tagged endocytic structures (DsRed-Rab5 and GFP-Rab7) in
target cells. We generated engineered lentiviruses lacking GFP-Vpr
(FUW/αCD20+SINmu or FUW/αCD20+HAmu) for this study. DiD-labeled
lentiviruses were incubated with 293T/CD20 cells transfected to express DsRed-Rab5
and GFP-Rab7 for 30 min at 4ºC. The cells were then moved to 37ºC for different
time periods and analyzed by confocal microscopy (Figure 3.5). The acquired images
showed that at 30 min, 62% of the viral fusion signals (n=98) for
FUW/αCD20+SINmu were detected in DsRed-Rab5+ organelles (Figure 3.5A, 30
min), and after 60 min, 78% of the fusion signals (n=93) were observed in both
DsRed-Rab5+ and GFP-Rab7+ organelles (Figure 3.5A, 60 min). On the other hand,
viral fusion of FUW/αCD20+HAmu was rarely detected at 30 min of incubation
(Figure 5B, 30 min) but after 60 min, 73% of the fusion signals (n=86) were observed
in endosomes positive for both DsRed-Rab5 and GFP-Rab7 (Figure 3.5B, 60 min).
Based on these observations, we concluded that fusion of SINmu-bearing viruses
takes place in early endosomes and the virus is further transported to intermediate
endosomes containing both early and late endosome markers, whereas fusion of
HAmu-bearing viruses does not occur until the viruses reach the intermediate
endosomes.
86
To further confirm our observations, viral fusion with Rab5- and Rab7-
endosomes was monitored in real-time. DiD-labeled lentiviruses bearing SINmu were
incubated with 293T/CD20 cells transfected with DsRed-Rab5 and GFP-Rab7 for 20
min at 37ºC to allow for the initial internalization of the viruses, then live cell
imaging was conducted using spinning-disk confocal microscopy. Selected images
obtained from a time-series are shown in Figure 3.6A. At 100 s, viral fusion with
endosomes was identified by the dramatic increase in signals as the result of DiD
dequenching. The images clearly show that the fusion signals (DiD) were colocalized
with DsRed-Rab5+ endosomes but not with GFP-Rab7+ endosomes. The
quantification of the fluorescent signals associated with fusion sites indicated that
viral fusion was detected in endosomes positive for DsRed-Rab5, but not GFP-Rab7
(Figure 3.6C), confirming that viral fusion of the SINmu-bearing virus occurs in early
endosomes. In contrast, analysis of the HAmu-bearing virus imaged after 50 min of
incubation at 37ºC showed that viral fusion was detected in endosomes where both
DsRed-Rab5 and GFP-Rab7 were present (Figure 3.6B and 3.6D), supporting the
conclusion that functional late endosomes are required for HAmu-bearing viral
fusion. In addition, to determine the accumulation of viral particles in the lysosomal
compartments, we monitored the colocalization of GFP-Vpr-labeled lentiviruses with
lysosome associated membrane protein 1 (Lamp-1), a marker for lysosomes
(Sieczkarski and Whittaker, 2003). After incubating the GFP-Vpr-labeled lentiviruses
(FUW-GFPVpr/αCD20+SINmu or FUW-GFPVpr/αCD20+HAmu) with 293T/CD20
cells at 37ºC for 4 h, the cells were immunostained against Lamp-1 (Figure 3.6E and
3.6F).
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Figure 3.5: Detection of viral fusion in DsRed-Rab5- and GFP-Rab7-expressing cells. DiD-labeled
lentiviruses displaying SINmu (A) or HAmu (B) were incubated with 293T/CD20 cells transiently
transfected with DsRed-Rab5 (red) and GFP-Rab7 (green) at 4ºC for 30 min to synchronize the
binding. The cells were then moved to 37ºC for 30 or 60 min, fixed, and analyzed for the
colocalization of the viral fusion signal (blue) with the different endosomal markers. The boxed
regions are enlarged in the right panels. Scale bar represents 5 µm.

The images showed that no significant colocalization (< 5%, n=97) of SINmu-bearing
viruses with lysosomes was observed (Figure 3.6E), whereas some viruses bearing
HAmu (16.8%, n=103) were detected in lysosomes.
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Figure 3.6: Selected frames from the monitoring of viral fusion in DsRed-Rab5- and GFP-Rab7-
expressing cells. DiD-labeled lentiviruses displaying SINmu (A) or HAmu (B) were added to
293T/CD20 cells transiently transfected with DsRed-Rab5 (red) and GFP-Rab7 (green) in the cold to
synchronize the infection. Time series imaging was acquired 20 or 50 min after incubation at 37ºC to
initiate the internalization of SINmu- or HAmu- bearing lentiviruses, respectively. Scale bar represents
2 µm. (C, D) Fluorescent time trajectories of DiD (blue), DsRed-Rab5 (red), and GFP-Rab7 (green)
signals associated with viral fusion (arrow) of SINmu-displaying lentivirus (FUW/αCD20+SINmu, C)
or HAmu-displaying lentivirus (FUW/αCD20+HAmu, D) shown in A or B, respectively. (E, F) Viral
degradation in the lysosomal compartments. GFP-Vpr-labeled viruses (green) enveloped by αCD20
and SINmu (E), or HAmu (F) were added to 293T/CD20 cells at 4ºC for 30 min to synchronize the
binding. The cells were then warmed to 37ºC for 4 h, fixed, permeabilized, and immunostained with
lysosome associated membrane protein 1 (Lamp-1; red). The boxed regions are enlarged in the right
panels. Scale bar represents 5 µm.

89
This seemingly low colocalization (16.8%) could be significant if one considers that
only portions of the degrading viruses in the lysosomes could be seen at a particular
time point. The imaging results suggest that HAmu-bearing lentiviruses are more
prone to degradation in the lysosomal compartments than SINmu-displaying
lentiviruses.

3.2.6 Hemifusion mediated by the SINmu fusogen occurs faster than that by the
HAmu fusogen at low pH
Membrane fusion proceeds through at least two steps: lipid mixing (i.e.
hemifusion) and content mixing (i.e. fusion pore formation and/or enlargement)
(Dimitrov, 2004; Harrison, 2008; Melikyan et al., 2005). Analysis of both lipid and
content mixings between a single virus and its target membrane is useful for
elucidating the molecular details of the membrane fusion process, but direct
observation of such processes within endosomes is difficult due to the constant
movement of the virus-endosome complex. To overcome this difficulty, we adhered
viral particles to a planar substrate and placed the target cells above the viruses, then
triggered virus-cell membrane fusion by lowering the pH environment. This setup
allowed us to monitor both lipid- and content-mixing processes within the same focal
plane (Markosyan et al., 2005; Melikyan et al., 2005).
90
With this experimental setup, we compared the kinetics of hemifusion mediated by
HAmu and SINmu fusion proteins after exposure to the optimal low pH at which
influenza (pH 5.0) and Sindbis viruses (pH 5.5) are known to be able to fuse with
cells (Chernomordik et al., 1998; Düzgüneş et al., 1992; Smit et al., 1999; White et
al., 1982; Zaitseva et al., 2005). The kinetics of hemifusion (lipid mixing) was
determined by observing the decrease of fluorescence signals of the lipophilic
carbocyanine dye (DiO) incorporated into the viral membrane at a low concentration
(below the self-quenching level). For the precise measurement of the time at which
the environmental pH drop occurred, we included virions labeled with a pH sensitive
dye, CypHer5, as a pH sensor. The mixture of DiO-labeled viruses and CypHer5-
labeled viruses (at the particle number ratio of 10:1) was adhered to the poly-lysine
coated coverslip and 293T/CD20 cells were overlaid on the viruses for 30 min. Virus-
cell fusion was triggered by adding the appropriate volume of 0.2 M acetic acid, pre-
titrated to provide the desired pH, and monitored by pH drop (CypHer5, red) and lipid
transfer (DiO, green) (Fig. 3.7A and 3.7B). There was no observable lipid transfer at
neutral pH (data not shown). Lipid mixing, indicated by the disappearance of the
green signal, for viruses bearing SINmu occurred quickly upon exposure to low pH,
as seen by the sudden increase in CypHer5 fluorescence (Fig 3.7A and 3.7C),
whereas HAmu-bearing viruses exhibited a longer lag time (~30 sec) between pH
drop and lipid mixing (Fig 3.7B and 3.7D). Thus, these results indicate that the
hemifusion of lentiviruses with SINmu is faster than that of HAmu-bearing virus.

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Figure 3.7: Detection of hemifusion between viruses and target cells with a pH sensor. DiO-labeled
(green) and CypHer5-labeled (red) viruses (mixing ratio 10:1) bearing SINmu (A) or HAmu (B) were
prebound to 293T/CD20 cells for 30 min at room temperature. During live-cell imaging, virus-cell
fusion was triggered by adding the appropriate volume of acidic buffer, pre-titrated to provide the
desired pH (HAmu: pH 5.0, SINmu: pH 5.5), and monitored by pH drop (asterisk) and lipid transfer
(arrow). Scale bar represents 2 µm. (C, D) The fluorescent intensity of the pH-sensitive CypHer5 (red)
and the hemifusion (green) signals associated with the viral particles bearing SINmu (C) or HAmu (D),
which are indicated by the asterisk and the arrow in A or B, respectively.

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3.2.7 Fusion pore formation mediated by SINmu, but not by HAmu, is delayed after
lipid mixing
For a more detailed analysis of hemifusion and fusion pore formation catalyzed by
Hamu and SINmu fusion proteins, viral particles were double-labeled with lipophilic
dye (DiD), for the detection of hemifusion, and DsRed-F (DeRed fused at the N-
terminus with the farnesylation sequence of p21 (Ras) (Harvey et al., 2001)), which
could be incorporated into the inner leaflet of the viral membrane, for the detection of
fusion pore formation. This labeling scheme allowed for the analysis of both lipid-
and content-mixing trajectories of single viruses within target cells (Campbell et al.,
2007). The kinetics between hemifusion and fusion pore formation were measured by
observing the loss in DiD and DsRed-F signals as the result of lipid- and content-
mixing, respectively. DiD/DsRed-F-labeled viruses were adhered to a poly-lysine-
coated coverslip and were then prebound to 293T/CD20 cells for 30 min.
Colocalization analysis of the viral particles showed that 40~60% of the DsRed-F-
labeled particles contained DiD. Virus-cell fusion was then triggered by low pH
(HAmu: pH 5.0, SINmu: pH 5.5) and monitored by lipid (DiD, green) and content
(DsRed-F, red) mixing (Fig. 3.8A and 3.8B). Interestingly, it was observed that the
decay of the DsRed-F signals on SINmu-bearing viruses was usually not
instantaneous after the DiD transfer (Fig. 3.8A and 3.8C), while the loss of the DiD
and the DsRed-F signals on HAmu-bearing viruses usually occurred simultaneously
(Fig. 3.8C and 3.8D).
93
Quantitative analysis of the delay times between lipid mixing (tL) and content mixing
(tC) for single viruses showed that ~60% of the DsRed-F transfer events mediated by
SINmu (n= 53) were delayed around 3 to 6 sec after the DiD transfer (resolution, 3
sec/frame), whereas ~85% of the DsRed-F transfer events mediated by HAmu (n=
52) were detected simultaneously (in the same frame) with the DiD transfer (Fig.
3.8E). These results suggest that the hemifusion intermediate (i.e. lipid stalk, before
fusion pore formation) mediated by SINmu is maintained relatively longer than that
induced by HAmu (Fig. 3.8F).

94

Figure 3.8: Hemifusion and fusion pore formation between viruses and target cells. DiD/DsRed-F-
labeled viruses displaying SINmu (A) or HAmu (B) were prebound to 293T/CD20 cells for 30 min at
room temperature, and then virus-cell fusion was triggered by adding the appropriate volume of acidic
buffer, pre-titrated to provide the desired pH (HAmu: pH 5.0, SINmu: pH 5.5). Lipid (DiD, green) and
content (DsRed-F, red) mixing was monitored, as indicated by the disappearance of signals. Yellow
signals indicate viral particles labeled by both DiD and DsRed-F. Viral particles that contain either
DiD or DsRed-F are shown in green or red, respectively. The viral particles that transferred the DiD
and DsRed-F signals are indicated by arrows in A and B. Scale bar represents 2 µm. (C, D) The
fluorescent intensity of DiD (green) and DsRed-F (red) signals associated with the viral particles
bearing SINmu (C) and HAmu (D), as indicated by the arrows in A or B, respectively. (E) Distribution
of the delay times between lipid mixing (tL) and content mixing (tC) of individual SINmu (black bar,
n=53) or HAmu (red bar, n=52) particles. The time intervals (tC-tL) between DiD and DsRed-F signal
disappearance were measured as shown in C and D. (F) A proposed sequence of events in membrane
fusion driven by HAmu and SINmu fusion proteins.

95
3.3 Discussion
Membrane fusion plays an essential role in the entry of enveloped viruses into
target cells. The pathways and underlying mechanisms of membrane fusion might be
different for different viral fusion proteins and this difference has not been well-
characterized. Our method of engineering targeting lentiviruses involves the
incorporation of a cell-binding protein and a fusion protein as two distinct molecules
on the lentiviral surface, which provides a platform for such a study. This platform
allows for the direct comparison of fusion processes mediated by various fusion
proteins without need for concern over the different effects of binding caused by the
different viruses. In this study, we used a real-time imaging technique to compare the
entry pathways and the membrane fusion processes of lentiviruses enveloped with
binding-deficient, fusion-competent proteins derived from either a class I fusion-
associated glycoprotein, influenza virus (HAmu), or a class II fusion-associated
glycoprotein, Sindbis virus (SINmu), along with αCD20. The viral entry mechanism
of this engineered virus is hypothesized to begin with the binding of αCD20 to its
cognate CD20 antigen, which expressed on the target cell surface, then the virus is
internalized into the acidic endosomes where the surface-displayed fusogen triggers
membrane fusion. This molecular process has been supported by our direct
visualization approach using confocal microscopy (Joo and Wang, 2008).
In present study, we investigated the early entry stage of these engineered
lentiviruses by visualizing the interactions of single viral particles with endocytic
structures in living cells. This study clearly showed that engineered lentiviruses
96
containing either SINmu or HAmu enter cells through pH-dependent and clathrin-
mediated endocytosis, a conclusion that was also confirmed by the drug inhibition
study. It was perhaps not surprising that we observed that both types of engineered
lentiviruses exploited the same entry route because they contained the same binding
molecule, αCD20, which likely determined the early stage of virus entry.
The actual fusion event of the engineered lentiviruses was detected in target
cells by a fluorescence-dequenching assay. This experiment revealed that viral fusion
of the SINmu-bearing virus occurred faster and exhibited relatively higher fusion
efficiency than that of the HAmu-bearing virus. In addition, viral transduction to cells
overexpressing the dominant-negative mutants of Rab proteins showed that the
HAmu-displaying virus required functional late endosome trafficking for productive
infection. This was further confirmed by real-time imaging of viral fusion, which
showed that the HAmu-bearing virus had to travel at least to intermediate endosomes
containing both early and late endosomal markers to initiate fusion. In contrast, viral
fusion of the SINmu-displaying virus started at the early endosome stage, which is
consistent with our previous observation (Joo and Wang, 2008). In addition, the
involvement of the late endosomal compartments in trafficking the SINmu virus
(Figure 3.4B and 3.5B, 60min) is in line with our previous observation, in which the
release of the core of the engineered SINmu lentivirus was associated with the
maturing process of the early to the intermediate endosomes (Joo and Wang, 2008).
The results from the current study is in good agreement with other studies,
demonstrating that the influenza virus requires late endosomal compartments for
97
successful infection (Sieczkarski and Whittaker, 2003; Stegmann T, 1987; Yoshimura
and Ohnishi, 1984), while the Semliki Forest virus (SFV), a member of the alphavirus
family that the Sindbis virus also belongs to, undergoes fusion at the early endosome
level (Sieczkarski and Whittaker, 2003; White and Helenius, 1980). Additionally, the
colocalization assay with lysosomal markers suggests that HAmu-bearing viruses
seem to be more prone to degradation in the lysosomes than SINmu-displaying
viruses.
Many studies have proposed that influenza and Sindbis virus glycoproteins
undergo a conformational change upon exposure to the low pH within endosomes,
which exposes the fusion peptide or loop(s) and allows it to interact with the target
membrane to cause hemfusion (Dimitrov, 2004; Harrison, 2008). By a real-time
imaging method along with a pH sensor, we were able to demonstrate the precise
kinetics of the hemifusion of single viruses upon exposure to the optimal low-pH
environment at which influenza and Sindbis viruses are known to be able to fuse with
cells (Chernomordik et al., 1998; Düzgüneş et al., 1992; Smit et al., 1999; White et
al., 1982; Zaitseva et al., 2005). Our results indicated that hemifusion mediated by
SINmu in response to low pH is faster than that induced by HAmu, which is
consistent with results from virus-liposome fusion assays (Shangguan et al., 1996;
Smit et al., 1999; Smit et al., 2002).
It has been proposed that membrane fusion by both class I and II fusion
proteins progresses through a transient hemifusion intermediate (Jahn et al., 2003),
but fluorometer-based conventional fusion assays are generally limited to monitoring
98
only the early fusion events of single viruses. A method of dual labeling the outer and
inner leaflets of single virus membranes enabled us to sequentially detect hemifusion
(lipid mixing) and fusion pore formation (content mixing) between individual viruses
bearing HAmu or SINmu fusion proteins and target cells. We observed that more than
60% of the fusion pore formation events mediated by SINmu were remarkably
delayed after hemifusion, while ~85% of the HAmu-mediated fusion pores were
formed almost simultaneously with hemifusion, indicating that SINmu-mediated
hemifusion intermediate is relatively long-lived compared with that of HAmu (Fig.
3.8F). This imaging result might provide an explanation for the fusion process
leakiness observed by an experimental model of liposome-based virus fusion, which
suggested that the influenza virus-mediated fusion process was quite leaky
(Shangguan et al., 1996), whereas the fusion process mediated by alphaviruses is non-
leaky (Smit et al., 2002). The long-lived hemifusion intermediate caused by SINmu,
in which the proximal (outer) leaflets of the interacting membranes have exchanged
their lipid components while the distal (inner) leaflets remain unopened, can yield a
non-leaky fusion process. On the other hand, the occurrence of the very transient
hemifusion intermediate induced by HAmu suggests that the interactions between the
outer and inner leaflets of the two distinct membranes are nearly simultaneous,
resulting in unstable contact of the two lipid bilayers, thereby creating a leaky fusion
process. One possible explanation for the marked delay between lipid mixing
(hemifusion) and content mixing (fusion pore formation) observed in SINmu-virus
fusion could be that there is a requirement for membrane components such as
cholesterol and sphigolipids in the target membrane for alphavirus fusion (Phalen and
99
Kielian, 1991; Smit et al., 1999), but not for influenza virus fusion (Sieczkarski and
Whittaker, 2002). Some studies have suggested that cholesterol and sphingolipids are
needed for the formation of fusion pores, which supports our explanation (Nieva et
al., 1994; Smit et al., 1999; White and Helenius, 1980). We are currently conducting
experiments to investigate the exact effects of these membrane lipid components on
the formation of the hemifusion intermediate upon virus fusion.
The recent study by Floyd et al. has developed a two-color fluorescent assay
to monitor kinetics of single influenza virus particles fusing with a target planar
bilayer and suggested a delay between hemifusion and pore formation for influenza
HA. The reason for this difference is likely due to a different labeling method
employed to detect fusion pore formation (Floyd et al., 2008); the pore formation was
monitored by the loss of sulforhodamine B signal (viral interior labeling), resulted
from the dye-diffusion into the fluid support of the bilayer after the fusion pore open,
that may be kinetically slower than the loss of the dye incorporated into the inner
leaflet of the viral membrane. The different experimental set-up for the viral fusion
assay (e.g. target membranes) could also be another cause for the difference in
kinetics of hemifusion and/or fusion pore formation.
To summarize, our findings suggest that engineered lentiviruses enveloped
with either HAmu or SINmu fusion proteins have different requirements of
endosomal trafficking for viral fusion. The results also indicate that SINmu (class II
fusion protein)-mediated hemifusion reacts faster upon exposure to low pH, but
delays the formation of the fusion pore as compared with the processes mediated by
100
HAmu (class I fusion protein). In addition, we have demonstrated that this imaging
approach can provide a useful technological platform to dissect the mechanisms of
viral membrane fusion.

3.4 Materials and Methods
3.4.1 Cell lines, antibodies, reagents
The 293T/CD20 cell line was generated previously (Yang et al., 2006). Cells
were maintained in a 5% CO2 environment in Dulbecco’s modified Eagle’s medium
(Mediatech, Inc., Manassas, VA, USA) with 10% FBS (Sigma, St Louis, MO, USA)
and 2 mM L-glutamine (Hyclone, Logan, UT, USA). Mouse monoclonal antibodies
against early endosomal antigen 1 (EEA1), clathrin, caveolin-1, and lysosome-
associated membrane protein 1 (Lamp-1) were purchased from Abcam (Cambridge,
MA, USA). Texas red-conjugated goat anti-mouse immunoglobulin G (IgG) antibody
was obtained from Molecular Probes (Carlsbad, CA, USA). Bafilomycin A1,
chlorpromazine, and filipin were purchased from Sigma.

3.4.2 Plasmids
Assembly PCR was employed to fuse GFP to the N-terminus of Vpr.
101
The PCR product was then inserted into the expression plasmid pcDNA3 (Invitrogen,
Carlsbad, CA, USA). The cDNAs for Rab5 and Rab7 were PCR-amplified and
cloned into pcDsRed-monomer-C1 (Clontech, Mountain View, CA, USA) as
described (Joo et al., 2008). The plasmid encoding the dominant-negative mutant of
DsRed-Rab7 (Rab7T22N) was generated by site-directed mutagenesis using the
forward primer (5’- GTCGGGAAGAACTCACTCATGAACC-3’) and the backward
primer (5’- GGTTCATGAGTGAGTTCTTCCCGAC-3’). The integrity of the DNA
sequence for this mutant was confirmed by DNA sequencing. The constructs for
GFP-Rab7 and the dominant-negative mutant of DsRed-Rab5 were obtained from
Addgene (Cambridge, MA, USA).

3.4.3 Virus production
GFP-Vpr-labeled lentivectors enveloped with αCD20 and fusogenic protein
(SINmu or HAmu) were produced by transfecting 293T cells by a calcium phosphate
precipitation method. 293T cells at 80% confluence in 6 cm culture dishes were
transfected with 5 µg of the lentiviral vector FUW, together with 2.5 µg each of
pcDNA3-GFPVpr, pαCD20 which encodes a mouse/human chimeric anti-CD20
antibody, pIgαβ which encodes human Igα and Igβ, two immunoglobulin associated
proteins that are required for the surface expression of antibodies, pSINmu or
pHAmu, and the packaging vector plasmids (pMDLg/pRRE and pRSV-REV) (Yang
et al., 2006). For the production of VSVG-pseudotyped lentiviruses or retroviruses,
102
293T cells were transfected with 5 µg of the lentiviral vector FUGW or the retroviral
vector MIG, along with 2.5 µg each of the packaging plasmids (pMDLg/pRRE,
pRSV-REV, and gag-pol for lentiviruses; pClEco for retroviruses), and the envelope
plasmid (VSVG). The cells were washed at 4 h post-transfection, and the medium
was replaced. The viral supernatant was collected after 48 h post-transfection and
filtered through a 0.45-µm pore size filter. The viral supernatant could be
concentrated by ultracentrifugation (Optima L-90 K ultracentrifuge, Beckman
Coulter) for 90 min at 82,700 × g and resuspended in an appropriate volume of PBS
containing 5 mM MgCl2.

3.4.4 Viral transduction
For the viral transduction of drug-treated cells, 293T/CD20 cells were
preincubated with drugs (bafilomycin A1: 25 and 50 nM; chlorpromazine: 10 and 25
µg/ml; filipin: 1 and 5 µg/ml) for 30 min at 37ºC and the cells (0.2 × 10
6
per well)
were then spin-infected with 2 ml of viral supernatants in a 24-well culture dish at
2,500 rpm and 30°C for 90 min with a Sorval Legend centrifuge. The drug
concentration was maintained during the spin-infection. The cells were further
incubated for 3 h at 37ºC. The drugs were then removed and replaced with fresh D10
media. For the viral transduction with Rab protein-treated cells, 293T/CD20 cells
were transfected with DsRed-Rab5 or -Rab7 (either wild type or dominant-negative
mutant), then seeded (0.2 × 10
6
per well) and spin-infected with 2 ml of viral
103
supernatants in a 24-well culture dish. The percentage of GFP+ cells was analyzed by
FACS at day 3 post-infection. All transduction assays were performed in triplicate
and the results are presented as mean values with ± standard deviations.

3.4.5 Confocal imaging
Fluorescent images were acquired on a Zeiss LSM 510 META laser scanning
confocal microscope equipped with Argon, red HeNe, and green HeNe lasers using a
Plan-apochromat 63×/1.4 oil immersion objective. A Coherent Chameleon Ti-
Sapphire laser was attached as well for multiphoton imaging. Some images were
collected by a Yokogawa spinning-disk confocal scanner system (Solamere
Technology Group, Salt Lake City, UT) using a Nikon eclipse Ti-E microscope
equipped with a 60×/1.49 Apo TIRF oil objective and a Cascade II: 512 EMCCD
camera (Photometrics, Tucson, AZ, USA). An AOTF (acousto-optical tunable filter)
controlled laser-merge system (Solamere Technology Group Inc.) was used to
provide illumination power at each of the following laser lines: 491 nm, 561 nm, and
640 nm solid state lasers (50mW for each laser).
For the viral trafficking studies with different endocytic markers, GFP-Vpr-
labeled viruses (FUW-GFPVpr/αCD20+SINmu or FUW-GFPVpr/αCD20+HAmu) at
a multiplicity of infection (MOI) ~ 20 were added to 293T/CD20 cells at 4ºC for 30
min. The cells were then kept at 37ºC for various incubation periods, fixed with 4%
formaldehyde, permeabilized with 0.1% Triton X-100, and immunostained with
104
antibodies against early endosomes, clathrin, or caveolin-1. Texas red-conjugated
goat anti-mouse IgG antibody was used as the secondary antibody. To remove viral
aggregates, virus-containing media were filtered by 0.45-µm pore size centrifuge tube
filters (Costar, NY, USA) before the experiments were conducted.
To image virus-endosome fusion, the concentrated GFP-Vpr-labeled viruses
were incubated with 50 µM of 1,1’-dioctadecyl-3,3,3’,3’-
tetramethylindodicarbocyanine (DiD) (Molecular Probes) for 1 h at room
temperature. Unbound dye was removed by Microcon filter units with a 50kDa cutoff
(Millipore, Billerica, MA) or by Zeba gel filtration columns (Thermo Scientific).
Double-labeled viruses were incubated with 293T/CD20 cells at 4ºC for 30 min to
synchronize infection, after which the cells were moved to 37ºC for various time
periods and fixed with 4% formaldehyde. All samples were scanned under the same
conditions for magnification, laser intensity, brightness, gain, and pinhole size. To
observe viral fusion in different endosomal compartments, 293T/CD20 cells were
transfected with DsRed-Rab5 and GFP-Rab7 plasmids for early endosome and late
endosome markers, respectively. At 48 h post-transfection, cells were seeded onto
glass-bottom culture dishes (MatTek Corporation, Ashland, MA, USA) and grown at
37ºC overnight. DiD-labeled viruses (FUW/αCD20+SINmu or FUW/αCD20+HAmu)
at MOI ~ 20 were then incubated with the cells at 4ºC for 30 min to synchronize
infection. The cells were then kept at 37ºC for 30 or 60 min and fixed. Images were
analyzed with the use of the Zeiss LSM 510 software version 3.2 SP2 or the Nikon
NIS-Elements software.
105
3.4.6 Live cell imaging of virus-endosome fusion
For the real-time detection of viral fusion in different endosomes, 293T/CD20
cells were transfected with DsRed-Rab5 and GFP-Rab7 plasmids. DiD-labeled
viruses (FUW/αCD20+SINmu or FUW/αCD20+HAmu) at MOI ~ 20 were then
incubated with the cells at 4ºC for 30 min to allow for virus binding. The cells were
then warmed to 37ºC for 20 or 50 min to induce the viral internalization of SINmu- or
HAmu-displaying lentiviruses. Confocal time-lapse images were recorded by
spinning-disk confocal microscopy.
Imaging of virus-cell fusion. For the real-time detection of virus-cell fusion, 293T
cells were transfected with plasmids for virus production as described above. To label
the inner leaflets of the virus membrane, cells were cotransfected with a plasmid
encoding DsRed-monomer-F (Clontech, Mountain View, CA, USA). The transfected
cells were washed at 4 h post-transfection and incubated with 2.5 µM of DiO (3,3′-
dioctadecyloxacarbocyanine perchlorate) or DiD in serum-free medium for 3 h at
37ºC to label the virus at levels below self-quenching concentrations. The cells were
washed again and regular D10 medium was added. To label the virus with pH-
sensitive CypHer5E mono NHS ester (GE Healthcare), the concentrated viruses were
incubated with the dye in 0.1 M sodium bicarbonate buffer (pH 9.3) for 1 h at room
temperature. Unbound dye was removed via buffer exchange into PBS (pH 7.4) using
a gel filtration column.
To detect lipid mixing upon exposure to low pH, the DiO-labeled and
CypHer5-labeled viral particles (total ~ 106 infectious units) were mixed at a particle
106
number ratio of 10 to 1 and adhered to poly-lysine-coated no.1 coverslips for 1 h at
37ºC. To monitor the sequential steps of lipid and content mixing, DiD-/DsRed-F-
labeled viruses (~10
6
infectious units) were immobilized on poly-lysine-coated
coveslips. After removing unbound viruses by PBS washing, 293T/CD20 cells were
overlaid on the viruses and incubated for 30 min at room temperature. Virus-cell
fusion was triggered by adding the appropriate volume of 0.2 M acetic acid,
pretitrated to achieve the desired final pH. Time-lapse images were captured at ~ 3 s
intervals. Fluorescent intensity versus time within the regions of interest was analyzed
using the Nikon NIS-Elements software.

107
Chapter 4. DEVELOPING QUANTUMM DOT LABELING OF
ENVELOPED / NON-ENVELOPED VIRUSES FOR SINGLE-VIRUS
TRACKING

Portions of this Chapter are adapted from:
Kye-Il Joo, Yuning Lei, Chi-Lin Lee, Jonathon Lo, Jiansong Xie, Sarah F. Hamm-
Alvarez and Pin Wang, ACS Nano. (2008) 2, 1553–1562

We report here a general method of labeling enveloped / non-enveloped
viruses with semiconductor quantum dots (QDs) for use in single virus trafficking
studies. Retroviruses, including human immunodeficiency virus (HIV), could be
successfully tagged with QDs through the membrane incorporation of a short
acceptor peptide (AP) that is susceptible to site-specific biotinylation and attachment
of streptavidin-conjugated QDs. We showed that this AP tag-based QD labeling had
little effect on the viral infectivity and allowed for the study of the kinetics of the
internalization of the recombinant lentivirus enveloped with vesicular stomatitis virus
glycoprotein (VSVG) into the early endosomes. It also allows for the live cell
imaging of the trafficking of labeled virus to the Rab5+ endosomal compartments.
We further demonstrated by direct visualization of QD-labeled virus that VSVG-
pseudotyped lentivirus enters cells independent of clatherin- and caveolin-pathways,
while the entry of VSVG-pseudotyped retrovirus occurs via the clathrin pathway. Our
studies monitoring HIV particles using QD-labeling showed that we could detect
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single virions on the surface of target cells expressing either CD4/CCR5 or DC-
SIGN. Further internalization studies of QD-HIV evidently showed that the clathrin
pathway is the major route for DC-SIGN-mediated uptake of viruses. Taken together,
our data demonstrates the potential of this QD-labeling for visualizing the dynamic
interactions between viruses and target cell structures.
Non-enveloped viruses, for example, the poliovirus, rotavirus, parvovirus
including the adeno-associated virus, and the adenovirus, are small and stable
particles lacking a lipid bilayer membrane. Although they are relatively simple in
structure, the intracellular trafficking of viruses is poorly understood. Due to robust
nature of non-enveloped viruses, the capsids of purified viruses can be targeted for
virus labeling by the covalent of dyes without a significant loss of infectiviety. Here
we report the general strategy for linking adeno-associated virus serotype 2 (AAV2),
which is only about 20 nm in diameter, with quantum dots through a coupling
reaction.

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4.1 Introduction
4.1.1 Site specific labeling of enveloped viruses with quantum dots for single virus
tracking
The ability to track individual viruses is a powerful tool for investigating viral
infection routes and characterizing the dynamic interactions between viruses and
target cells (Brandenburg and Zhuang, 2007). It enables the possible elucidation of
previously unknown but critical steps involved in the penetration of viruses into cells
and dissemination of viruses, revealing novel therapeutic opportunities for controlling
virus pandemics and pathogenesis (Brandenburg and Zhuang, 2007; Marsh and
Helenius, 2006). The first step towards the realization of single virus tracking in live
cells is to label the external and/or internal constituents of viruses with fluorophores,
allowing imaging using fluorescence microscopy. While fluorescent proteins (such as
green fluorescent protein, GFP) can be genetically engineered to be incorporated into
the interior of viruses during viral assembly to accomplish the internal labeling of
viruses (McDonald et al., 2002; Sherer et al., 2003), labeling the external components
of viruses is usually achieved by using organic dyes through either chemical labeling
of capsid proteins of non-enveloped viruses (Seisenberger et al., 2001) or physical
incorporation into the membrane of enveloped viruses (Brandenburg and Zhuang,
2007; Lakadamyali et al., 2003; Rust et al., 2004; Vonderheit and Helenius, 2005).
Good photostability of the labeling fluorophores is desirable for the continuous
tracking of individual viruses because a high magnification objective has to be used
in order to detect these tiny viral particles (20-100 nm), generating high excitation
110
light intensity in the focal plane of the objective (Wu et al., 2003). In this context, the
recently characterized inorganic nanoparticle semiconductor quantum dots (QDs)
exhibit many promising features (such as remarkable photostability and brightness)
for these types of viral imaging applications (Bruchez et al., 1998; Chan and Nie,
1998; Michalet et al., 2005). Successful examples of tagging viruses with QDs in the
literature include: 1) the use of two or three antibody layers (a virus-specific primary
antibody, followed by a secondary antibody and streptavidin-QD) (Agrawal et al.,
2005; Bentzen et al., 2005); 2) encapsulation of QDs in viral capsids (Dixit et al.,
2006); 3) covalent linkage of QDs to viral capsids (Medintz et al., 2005; Portney et
al., 2007). However, these labeling schemes likely affect the normal properties of
viruses including their host interactions. We have yet to see a trafficking study of live
viruses using these labeling methods to determine whether their perturbations can be
tolerated during the process of viral infection. On the other hand, development of
methods that can direct the QDs to selected positions of the viruses to ensure minimal
disturbance of virus-host interactions can open up new opportunities for single virus
tracking in live cells.
The goal of this study was to develop and characterize a general method to
site-specifically label live and membrane-enveloped viruses using QDs. Our strategy
was to first incorporate a 15-amino acid biotin acceptor peptide (AP) tag onto the
surface of a virion (Figure 4.1) (Beckett et al., 1999; Howarth et al., 2005).
Subsequently, biotin ligase (BirA) was used to specifically modify the AP-tag to
introduce the biotin moiety to the viral surface.
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Due to the tight interaction between biotin and streptavidin (Kd = 10-13 M) (Piran
and Riordan, 1990), further addition of streptavidin-conjugated QDs allows the site-
specific labeling of viral particles with photostable and fluorescent QDs. Such a
labeling method has been successfully applied to targeting QDs to surface proteins in
living cells (Howarth et al., 2005). We focused here on retroviruses to demonstrate
the viability and utility of this method for virus labeling, but this method could easily
be applied to other types of membrane-enveloped viruses.

4.1.2 Enhanced monitoring of non-enveloped viruses with quantum dot labeling
Non-enveloped viruses are small and stable particles lacking a lipid bilayer
membrane. Since they can form crystals that diffract to good resolutions, the
structures of a relatively large number of representatives from different virus families
have been determined by X-ray crystallography to high resolutions (Dimitrov, 2004).
However, the membrane penetration and uncoating mechanisms are poorly
understood for these viruses, even though they are relatively small and simple in
structure (Hogle, 2002). The membrane penetration process requires significant
reorganization of the viral nucleoprotein complex compared with enveloped viruses
(Dimitrov, 2004; Hogle, 2002). Some examples of non-enveloped viruses include the
poliovirus, rotavirus, parvovirus, including the adeno-associated virus (AAV), and the
adenovirus. Due to the robust nature of non-enveloped viruses, the capsids of purified
viruses can be labeled by the covalent attachment of dyes without a significant loss of
virus infectivity (Bartlett et al., 2000; Greber et al., 1997).
112
AAV, in particular, has attracted considerable interest because it shows great
promise for use in human gene therapy (Mueller and Flotte, 2008). Since the AAV
virion is only about 20 nm in diameter, the number of dye molecules that can be
attached to a single virus without causing self-quenching or affecting viral infectivity
is very limited (Seisenberger et al., 2001). Quantum dots are much brighter than the
conventional fluorophores, which can allow the detection of viruses with much lower
amounts of labeling molecules. It is believed that AAV virions are transported into
the nucleus before viral uncoating occurs (Ding et al., 2005; Seisenberger et al.,
2001), thus, the viral capsid can be targeted for labeling by quantum dots to track the
viral particle movements in the cell.
The general strategy for linking adeno-associated virus serotype 2 (AAV2)
with quantum dots through a coupling reaction is illustrated in Fig. 4.8. Quantum dot-
AAV2 networks were produced by covalent amide bonds formed by carbodiimide
chemistry between carboxylic moieties on the quantum dots and the primary amines
from the lysine residues on the virus capsid protein.

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Figure 4.1: General strategy for the site-specific labeling of enveloped viruses with QDs. Biotin
acceptor peptide (AP) tag is incorporated onto the surface of viruses through the natural budding from
cells expressing surface AP tag. The concentrated AP-tagged viruses resuspended in PBS-MgCl
2

buffer are biotinylated by adding biotin ligase (BirA), ATP, and biotin. Further incubation with
streptavidin-conjugated QDs allows the site-specific labeling of QDs to the biotinylated surface of
viruses.

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4.2 Results
4.2.1 Construction of AP-tag
We first designed a construct to allow the efficient incorporation of AP-tag to
the surface of virus-producing cells and subsequently to the surface of virions. The
construct contained a signal peptide derived from the human CD5 protein fused with
an AP-tag sequence and the transmembrane (TM) domain of the human CD7 protein,
designated AP-TM (Figure 4.2A). We tested this construct for its ability to display
AP-tag by transfecting virus-producing 293T cells with a plasmid encoding AP-TM.
After two days post-transfection, cells were washed with the buffer solution (PBS
containing MgCl2), biotinylated by adding BirA, biotin and ATP, and further labeled
with streptavidin-conjugated dye (R-Phycoerythrin) for flow cytometry analysis, or
QD (QD525) for confocal microscopy analysis. Fluorescent signals were detected for
cells transfected with AP-TM and treated with biotinylation (Figure 4.2B and 4.2C);
approximately 40% of cells displayed the AP-tag, as analyzed by flow cytometry. No
signal was detected for control cells without AP-TM transfection. Some background
signals were obtained for transfected cells without exogenous biotinylation (no BirA
treatment, Figure 4.2B and 4.2C); this was likely caused by the expression of
endogenous biotin ligase in 293T cells, which can endogenously biotinylate AP-tag
before it is transported to the cell surface (Howarth et al., 2005). Nevertheless, we
confirmed that the AP-TM construct could successfully display functional AP-tag on
the cell surface for site-specific biotinylation and labeling. It is noteworthy that both
115
BirA and streptavidin are membrane-impermeable (Howarth et al., 2005), thus no
intracellular protein can be tagged under our labeling condition.

Figure 4.2: Incorporation of AP tag onto the surface of 293T cells. (A) The schematic representation of
the AP-TM construct. 293T cells were transiently transfected with a plasmid encoding AP-TM. After
48 h post-transfection, the cells were incubated with biotin and ATP in the presence (B and C, right) or
absence of BirA (B and C, middle) and labeled with streptavidin-PE (PE-Savi) for analysis by flow
cytometry (B) or streptavidin-QDs (QD525-Savi) for confocal microscopy analysis (C). Cells without
AP-TM transfection were included as controls (B and C, left).
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4.2.2 Incorporation of AP-tag onto lentiviruses
We next determined whether AP-tag could be incorporated onto the surface of
lentivirus for QD labeling. HIV-1-based lentivirus is one type of retrovirus that
possesses the unique feature of transducing non-dividing cells and has been
recognized as one of the most efficient and potent systems for the development of
gene delivery vectors (Kohn, 2007; Somia and Verma, 2000). QD labeling of
lentivirus can potentially enable us to visualize the process of viral entry and
transport, which in turn can facilitate our understanding of the virus lifecycle and the
design of more efficient vectors for gene therapy. To produce AP-tag-bearing
lentivirus, we transiently transfected 293T cells with a lentiviral backbone plasmid
FUW (Lois et al., 2002), a plasmid encoding envelope glycoprotein derived from
vesicular stomatitis virus (VSVG), a plasmid encoding AP-tag (AP-TM), and other
necessary packaging plasmids (gag, pol, rev). Concentrated viral particles were
resuspended in MgCl2-containing PBS and biotinylated in the presence of BirA,
biotin and ATP, followed by incubation with QD525-streptavidin. We employed a
previously reported method to detect lentiviral particles using confocal imaging
(McDonald et al., 2002). The particle solutions were overlaid onto the coverslips, and
adhered viral particles were immunostained with an antibody specific for HIV capsid
protein (p24). Clearly the AP-tag was incorporated onto the viral surface, as the
fluorescence signal of QDs was readily detected on biotinylated virions, and most of
the QD signals (> 70%) were co-localized with p24 (Figure 4.3A, upper). No QD
signal was observed for viral particles lacking the AP-tag (Figure 4.3A, bottom).
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To test whether the labeled viruses remained infectious, we made lentiviruses
carrying a GFP reporter gene (FUGW (Lois et al., 2002), instead of FUW, was used
as the lentiviral backbone plasmid for preparation of the virus). We labeled half of the
viruses with QDs and used them to infect 293T cells; the other half was used as the
control for unlabeled viruses. It was found that a similar transduction efficiency was
obtained for the labeled (FUGW/VSVG+AP+QD525) and unlabeled lentiviruses
(FUGW/VSVG+AP) (Figure 4.3B), suggesting that this strategy of labeling
lentiviruses with QDs could allow us to retain viral infectivity.

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Figure 4.3: Incorporation of AP tag onto the surface of lentiviruses for QD labeling. (A) Vesicular
stomatitis virus glycoprotein (VSVG)-pseudotyped lentiviruses produced by cotransfection either with
AP-TM (FUW/VSVG+AP) or without AP-TM (FUW/VSVG) were incubated with BirA, ATP, and
biotin, followed by labeling with streptavidin-QD525. The viruses (green) were overlaid upon
polylysine-coated coverslips for 60 min at 37°C. The coverslips were fixed, permeabilized, and
immunostained with an antibody specific for HIV capsid protein p24 (red). Overlapping green and red
signals appears as yellow in a merged image. Scale bars represent 2 µm. (B) AP-tag bearing, VSVG-
pseudotyped lentiviruses encoding a GFP reporter gene were produced and biotinylated. Half of the
biotinylated viruses were further labeled with streptavidin-QD525 (FUGW/VSVG+AP+QD525), and
the other half were not QD-labeled as the control (FUGW/VSVG+AP). 293T cells (2 × 10
5
) were spin-
infected with QD-labeled or unlabeled viruses. The resulting GFP expression was analyzed by flow
cytometry. (C) 293T cells were spin-infected with QD-labeled gamma-retrovirus
(MIG/VSVG+AP+QD) or unlabeled gamma-retovirus (MIG/VSVG+AP). The resulting GFP
expression was analyzed by flow cytometry.
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4.2.3 Photostability of QD-labeled viral particles
We examined the photostability of streptavidin-conjugated QDs bound to
viruses. The AP-tag-bearing lentiviruses were individually labeled with streptavidin
conjugated with either QD (QD525) or fluorescent dye (fluorescein isothiocyanate,
FITC). The labeled viruses were illuminated continuously with the Argon laser (8
mW) to excite QD or FITC. In order to clearly see viral particles, we needed to use a
63× oil-immersion objective. Because of high light intensity generated at the focal
plane, this type of objective usually requires good photostability of labeling reagents.
After continuous exposure by the laser light, the dye-labeled viruses showed marked
photobleaching, much more significant than that with QD-labeled particles (Figure
4.4A and 4.4B). Thus, photostable QDs therefore have a great advantage to be used as
fluorescent probes for virus imaging, especially for trafficking study of live viruses in
cells, in which long-term illumination under confocal light is likely required.

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Figure 4.4: Photostability comparison between QD-labeled or FITC-labeled viral particles. (A)
Biotinylated VSVG-pseudotyped lentiviruses were labeled with streptavidin-QD525 or streptavidin-
FITC and overlaid upon poly-lysine coated coverslips. The specimens were continuously illuminated
by argon laser at 488 nm over 3 min. Images were captured at ~10 s intervals. Scale bars represent 2
µm. (B) Kinetics of the fluorescence intensity of QD-labeled or FITC-labeled viral particles. The
fluorescent intensity of viral particles was measured using the software package for the Zeiss LSM
510.

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4.2.4 Trafficking of QD-labeled lentiviruses
Having established the QD-labeling of lentivirus with retained infectivity
(Figure 4.3B), we tested whether such labeling could facilitate single viral particle
tracking within target cells, which would be of importance for understanding the
mechanism of viral entry and transport. As a demonstration, we focused on
monitoring the movement of lentivirus to the endosomal compartment, where the
membrane of the VSVG-pseudotyped lentivirus and the endosomal membrane are
believed to fuse together, an important step of successful infection (Brandenburg and
Zhuang, 2007; Marsh and Helenius, 2006). To mark the endosomes in living cells, we
transfected the target cells with a construct to express a small GTPase (Rab5) fused
with a fluorescent protein (DsRed); Rab5 is well-known to be associated with early
endosomes (Zerial and McBride, 2001). It has been shown that the expression of this
kind of chimeric protein does not affect intracellular trafficking of virus and viral
infectivity (Vonderheit and Helenius, 2005). Lentiviral particles (QD-labeled, VSVG-
pseudotyped) were initially added to 293T cells in the cold to synchronize the
binding. Cells were then shifted to 37oC for different time periods (10, 30, and 60
min), fixed, and analyzed by confocal microscopy. At 10 min, no virus (green) was
colocalized with the early endosome marker Rab5 (red). After 30 min, a few viruses
were seen to be colocalized with Rab5, as evidenced by the appearance of the yellow
color after overlay of both QD-virus and Rab5 images. After 60 min, most of the viral
particles were observed to be located in Rab5+ endosomes. The quantification of
colocalization suggested that at 10 min, < 5% of viruses were located in Rab5+
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organelles (n=50), at 30 min, 31% in Rab5+ organelles (n=65), and at 60 min, 68% of
viruses were observed in Rab5+ organelles (n=58). At To monitor the transport of
QD-labeled virus to endosomes in real-time, we incubated viruses with 293T cells for
10 min at 37°C to allow the initial internalization, and then began live cell imaging
using time-lapse confocal fluorescence microscopy. Selected images obtained from a
time series were shown in Figure 4.5B. The green virus (indicated by arrow) was
initially separated from red endosomes (0-458 sec), and then a fluorescent DsRed
spot emerged that was centered on the virus at 472 sec; the colocalization of the virus
with an endosome was maintained for an extended period of time (up to 708 sec).
Thus, we demonstrated the use of the photophysical properties of QDs to monitor the
intracellular movement of lentiviruses in live cells.

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Figure 4.5: The trafficking of QD-labeled viral particles through endosomes. (A) 293T cells transiently
transfected with DsRed-Rab5 (red) were seeded on glass bottom dishes at 48 h posttransfection. QD-
labeled VSVG-pseudotyped lentiviruses (green) were then incubated with the cells for 30 min at 4°C
to synchronize infection. The cells were shifted to 37°C for various time periods (10, 30, 60 min) and
then fixed. (B) Real-time monitoring of QD-labeled virus transport to endosomes. Rab5 (red)
expressing 293T cells were incubated with QD-labeled VSVG-pseudotyped lentiviruses (green) for 30
min at 4°C and shifted to 37°C for 10 min to initiate virus internalization. Confocal time-lapse images
were then recorded. The arrows indicate the internalized viral particle. Scale bars represent 5 µm.

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4.2.5 Clathrin/Caveolin-dependent entry of VSVG-pseudotyped retroviruses
Many viruses such as vesicular stomatitis virus (VSV) enter cells through
endocytosis (Sun et al., 2005). It is generally believed that retroviruses, including
lentiviruses and gamma-retroviruses, once enveloped with VSVG, are also
internalized to low pH endosomes to infect target cells. However, many of the
molecular details of entry mechanisms for these pseudotyped viruses remain poorly
understood and their entry has not been directly visualized. We decided to test if QD-
labeling could be a useful tool to study some endocytic pathways exploited by these
viruses to enter cells. We focused on clathrin- and caveolar-mediated pathways as
these two are well-characterized pathways of endocytosis (Marsh and Helenius,
2006). To track the endocytic structures in cells, we made constructs capable of
expressing the fluorescent protein-tagged clathrin (DsRed-clathrin) and caveolin
(DsRed-caveolin) based on a previous report (Tagawa et al., 2005). We incubated
293T cells transfected to express either DsRed-clathrin or DsRed-caveolin with QD-
labeled lentiviruses (FUW/VSVG+AP+QD) and imaged the individual viral particles
and endocytic structures after different incubation time periods (10 min and 30 min).
We found that many viruses had been internalized into cells after 10 min of
incubation (Figure 4.6). However, no significant colocalization of lentivirus and
clathrin (Figure 4.6A) or caveolin (Figure 4.6B) was detected during these time
periods (up to 30 min), suggesting that clathrin and caveolin were not involved in the
entry of VSVG-pseudotyped lentiviruses.
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Although this observation is somewhat unexpected, considering that native VSV
enters cells by a clathrin-dependent route (Sun et al., 2005), it is consistent with a
previous biochemical study, in which the entry efficiency of lentiviruses enveloped
with VSVG was not altered by the expression of a dominant-negative dynamin in
host cells (Daecke et al., 2005); dynamin is the cellular GTPase that is essential for
clathrin- and caveolin-associated endocytosis (Damke et al., 1994). This result
highlights a fact that there are many different endocytic routes, some of which have
yet to be defined, that cells can use to uptake particles and ligands (Marsh and
Helenius, 2006; Mayor and Pagano, 2007; Medina-Kauwe et al., 2005; Muro et al.,
2003; Romer et al., 2007). Similarly, we investigated the viral entry of VSVG-
enveloped gamma-retroviruses (MIG/VSVG+AP+QD). QD-labeling had little effect
on the infectivity of gamma-retrovirus (Figure 4.3C). Interestingly, significant
colocalization (67%, n=60) of discrete clathrin structures and QD-labeled gamma-
viruses was seen at 10 min of incubation (Figure 4.6C, left). After incubation for 30
min, a lesser degree of colocalization (21%, n=57) was detected (Figure 4.6C, right),
suggesting that several viruses had already dissociated from uncoated clathrin
structures and were likely transported to early endosomes. Thus, we obtained clear
evidence that clathrin-mediated endocytosis was involved in the entry of VSVG-
pseudotyped gamma-retroviruses, which is consistent with the results from many
biochemical assays (Blanchard et al., 2006; Lee et al., 1999; Meertens et al., 2006).
To further validate the result from the confocal imaging of QD-labeled
viruses, we performed assays to examine the inhibitory effect of viral entry by drug
treatment.
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Chlorpromazine is a drug known to prevent clathrin polymerization and obstruct the
internalization mediated by clathrin-coated vesicles (CCV) (Wang et al., 1993). We
found that chlorpromazine at concentrations of 10 and 25 µg/ml, could markedly
inhibit VSVG-pseudotyped gamma-retroviruses to infect 293T cells, whereas no
inhibitory effect was observed for VSVG-pseudotyped lentiviruses (Figure 4.6D).
The treatment with filipin, a drug capable of depleting cholesterol to inhibit caveolin-
dependent internalization (Orlandi and Fishman, 1998), did not affect the entry of
both viruses (Figure 4.6E), indicating that their entry is independent of caveolin.
Thus, QD-labeling of viruses allows us to visualize the clathrin- and caveolin-
independent entry of VSVG-pseudotyped lentiviruses and the involvement of clathrin
for the entry of VSVG-pseudotyped gamma-retroviruses, which are consistent with
the results from infection assays using corresponding inhibitors. Our study also
suggests that envelope glycoprotein is not the sole determinant of the viral entry
pathway. The exact means of cell entry for VSVG-pseudotyped lentiviruses remains
to be established.

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Figure 4.6: Clathrin/caveolin-dependent entry of VSVG-pseudotyped retroviruses. 293T cells
transiently transfected with DsRed-clathrin (red) or DsRed-caveolin (red) were seeded on glass bottom
dishes at 48 h posttransfection. QD-labeled VSVG-pseudotyped lentiviruses (green) were incubated
with cells that express clathrin (A) or caveolin (B) for 30 min at 4°C to synchronize infection. The
cells were shifted to 37°C for various time periods (10 and 30 min) and then fixed. The confocal
images were acquired and colocalization was analyzed. Similarly, DsRed-clathrin-expressing 293T
cells were incubated with QD-labeled, VSVG-pseudotyped gamma-retroviruses
(MIG/VSVG+AP+QD) for 30 min at 4°C, shifted to 37°C, and fixed at different time points (10 and
30 min) (C). Arrows indicate the viral particles colocalized with clathrin. (D & E) Inhibition of
clathrin-dependent internalization by chlorpromazine (D) or caveolin-dependent internalization by
filipin (E). 293T cells were preincubated with chlorpromazine or filipin for 30 min at 37°C. The cells
(2 × 10
5
) were then spin-infected with supernatants of VSVG-pseudotyped lentivirus (FUGW/VSVG,
black bar) or gamma-retrovirus (MIG/VSVG, gray bar). The percentage of GFP
+
cells was measured
by flow cytometry. Both drug concentrations were maintained during the spin-infection. The drugs
were then removed and replaced with fresh media. Scale bars represent 5 µm.
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4.2.6 Intracellular trafficking of QD-labeled HIV
We further explored the potential utility of QD-labeling for the study of HIV.
To test whether our QD-labeling method could be used to tag HIV particles, virus-
producing cells were transiently transfected with the plasmids FUW and AP-TM, a
plasmid encoding a codon-optimized and CCR5-tropic HIV-1 envelope protein (Haas
et al., 1996), and other necessary packaging plasmids (gag, pol, rev). Viral
supernatants were harvested and concentrated to obtain HIV particles, which were
further subjected to biotinylation and labeling with streptavidin-conjugated QDs. The
resulting particles were incubated with HeLa cells expressing viral receptor CD4 and
coreceptor CCR5 (HeLa/CD4/CCR5) and stained with an anti-CD4 antibody. Green
fluorescence signals could be clearly detected on the surface of HeLa/CD4/CCR5
(Figure 7A, upper), indicating the successful labeling of HIV particles. When the
same viral particles were applied to HeLa cells lacking the expression of viral
receptors, no green signal was observed (Figure 4.7A, lower), confirming that the
specific interaction between envelope glycoprotein and its cognate receptor accounted
for the observed binding of HIV to HeLa/CD4/CCR5.
In addition to the well known interaction of HIV envelope glycoprotein with
CD4 and appropriate chemokine receptors (CCR5 or CXCR4), HIV also binds to DC-
SIGN, a C-type lectin predominately expressed on immature dendritic cells (DCs)
(Wu and KewalRamani, 2006). Instead of mediating infection, this HIV-DC binding
induces the internalization of intact HIV into a nonlysosomal compartment, where its
competence of infection could be retained for an extended period of time before
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transfer to target cells, resulting in the trans-enhancement of HIV infection to T cells
(Geijtenbeek et al., 2000; Kwon et al., 2002; Wu and KewalRamani, 2006). Although
the study of this internalization using QD-labeled envelope glycoprotein gp120 is
insightful (Cambi et al., 2007; Turville et al., 2004), there has been no report on direct
visualization of the binding, entry, and trafficking of HIV particle in DC-SIGN-
expressing cells. We tested the feasibility of using the QD-labeled HIV (QD-HIV) to
track this internalization. Upon incubation for 30 min at 4°C, QD signals could be
readily detected on the surface of 293T cells expressing DC-SIGN (293T/DC-SIGN)
(Figure 4.7B). Significant internalization of QD-HIV was seen after prolonged
incubation with 293T/DC-SIGN at 37°C. To examine whether the DC-SIGN-
mediated uptake of HIV was clathrin-dependent, we transfected 293T/DC-SIGN cells
with DsRed-clathrin. The resulting cells and QD-HIV were incubated for 10 min at
37°C, followed by fixation and confocal imaging. Approximately 65% of the QDs
(n=51) were colocalized with CCV, indicating that the entry is mediated by a clathrin-
dependent pathway. We further studied the colocalization of QDs with the Rab5
protein. About 62% of internalized HIV particles labeled with QDs (n=47) were
colocalized with DsRed-tagged Rab5, confirming that DC-SIGN-mediated
endocytosis of HIV particles relies on a clathrin-dependent pathway to enter the early
endosomes.

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Figure 4.7: Binding and intracellular trafficking of QD-labeled HIV. (A) Visualization of HIV viral
particles on the surface of HeLa cells expressing viral receptor CD4 and coreceptor CCR5. HeLa cells
or HeLa/CD4/CCR5 cells were seeded on glass bottom dishes overnight. The cells were incubated
with QD-labeled HIV viral particles (green) for 30 min at 4°C, fixed, and then immunostained with
anti-human CD4 antibodies (red) and counterstained with DAPI (blue). Arrows indicate QD-labeled
HIV viral particles bound to the cell surface. (B) 293T/DC-SIGN cells were seeded and incubated with
QD-labeled HIV viral particles (green) for 30 min at 4°C. The cells were fixed and immunostained
with anti-human DC-SIGN antibodies (red) and counterstained with DAPI (blue). Arrows indicate
HIV viral particles bound to DC-SIGN. (C) Involvement of clathrin-dependent internalization of HIV
in DC-SIGN-expressing cells. 293T/DC-SIGN cells were transiently transfected with DsRed-clathrin
and seeded on glass bottom dishes at 48 h posttransfection. The cells were incubated with QD-labeled
HIV viral particles for 30 min at 4°C to synchronize infection, shifted to 37°C for 10 min, and then
fixed. Arrows indicate HIV viral particles colocalized with clathrin. (D) HIV viral transport to
endosomes. 293T/DC-SIGN cells transiently transfected with DsRed-Rab5 were seeded on glass
bottom dishes. The cells were incubated with QD-labeled HIV viral particles for 30 min at 4°C, shifted
to 37°C for 30 min, and then fixed. Arrows indicate HIV viral particles colocalized with early
endosomes. Scale bars represent 5 µm.
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4.2.7 Covalent attachment of quantum dots on adeno-associated viruses
Since non-enveloped viruses that lack a lipid bilayer membrane are generally
believed to be more robust than enveloped viruses, the capsid proteins of purified
non-enveloped viruses are usually targeted for virus labeling via a direct chemical
labeling with dyes. One method of labeling adeno-associated virus with quantum dots
is through a coupling reaction illustrated in Fig. 4.8A. Quantum dot-AAV2 networks
were produced by covalent amide bonds formed by carbodiimide chemistry between
carboxylic moieties on the quantum dots and the primary amines from the lysine
residues on the virus capsid protein. The carboxyl group on the quantum dots were
incubated with 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC)
and N-hydroxysuccinimide (NHS) to modify the carboxyl group to an amine-reactive
NHS ester. Then, excess EDC and NHS were removed by a gel filtration column and
the primary amine source, AAV2, was added for a coupling reaction.

132

Figure 4.8: Covalent attachment of quantum dots on adeno-associated virus serotype 2 (AAV2). (A)
The schematic representation of the viral labeling with quantum dots. (B) Colocalization of QD705-
labeled AAV2 with anti-AAV2 antibody. QD705-labeled AAV2 (red) were overlaid upon poly-lysine
coated coverslips for 60 min at 37°C. The coverslips were fixed and immunostained with an antibody
specific to adeno-associated virus capsid protein (green). Overlapping green and red signals appears as
yellow in a merged image. Scale bars represent 2 µm. (C) Hela cells were (2 x 10
4
) were spin-infected
with QD-labeled or unlabeled viruses. The resulting GFP expression was analyzed by flow cytometer.
133
We confirmed that the quantum dots were indeed coupled to AAV2 by overlaying the
particle solutions onto coverslips and immunostaining the adhered viral particles with
an antibody specific for intact AAV2. The fluorescence signal of the quantum dots on
the viral surface was readily detected and almost all of the quantum dot signals were
colocalized with the signal generated by anti-AAV2 antibody staining (Fig. 4.8B). It
was also found that a similar transduction efficiency was obtained for the labeled and
unlabeled viruses, suggesting that this labeling strategy with quantum dots could
allow us to retain viral infectivity (Fig. 4.8C).

134
4.2.8 Photostability and detection sensitivity of QD-labeled AAV2
To compare the photostability of QD-labeled viruses to fluorescent dye-
labeled viruses, QD705- or FITC-labeled AAV2 were co-incubated with hela cells for
30min at 37°C. The cells were then fixed and were illuminated continuously with the
laser at 491nm (50 mW) to excite QD and FITC. As shown in Fig. 4.9A and 4.9B, the
dye labeled AAV2 showed remarked photobleaching, whereas QD-labeled AAV2
retained the fluorescent signal during live cell imaging.
Live cell imaging technology with fluorescent microscopy (e.g. spinning disk
confocal microscopy) has allowed tracking of the dynamic interactions between
viruses and cellular structures. In multicolor real-time imaging studies, it is always
desirable to acquire an image with exposure times as short as possible to monitor the
dynamics of virus trafficking in living cells in greater detail. However, the limited
number of fluorescent dyes that can be attached on a single virus without affecting
viral infectivity sets an upper limit to the exposure time needed to detect the
fluorescent signal emitted from a tiny virus. The use of quantum dots can potentially
offer advantages for fast, multicolor time-lapse imaging due to their remarkable
brightness compared with conventional fluorophores. The sensitivity of the quantum
dot-AAV2 particles was compared with fluorophore-labeled AAV2 and it was
determined that the quantum dot signal was much brighter and could be detected at a
shorter exposure time (Fig. 4.9C).

135

Figure 4.9: The photostability and detection sensitiviety of QD-labeled AAV2. (A) QD705- or FITC-
labeled AAV2 were co-incubated with hela cells for 30min at 37°C. The cells were then fixed and
were illuminated continuously with the laser at 491nm over 10 min. (B) Kinetics of the fluorescent
intensity of QD-labeled or FITC-labeled viral particles. (C) Detection sensitivity comparison between
QD705-labeled or FITC-labeled viral particles at different exposure times. AAV2 were labeled with
either QD705 or FITC and incubated with HeLa cells for 30 min at 37°C. The cells were then fixed
and illuminated by an argon laser at 491 nm at different exposure times. Scale bars represent 5 µm.

136
4.2.9 Clathrin/Caveolin-dependent entry of AAV2
It has been generally believed that AAV2 enter the cells through clathrin-
coated pits in dynamin-dependent manner (Duan et al., 1999). To confirm the role of
clathrin- or caveolin-mediated endocytosis in the entry of AAV2, we visualized the
individual viral particles and endocytic structures (clathrin or caveolin) in target cells
after 5 min incubation at 37°C. As shown in Fig. 4.10A, significant colocalization of
AAV2 particles with the discrete clathrin structures was detected. Although some
particles were overlaid with caveolin structures, remarked colocalization was not
observed. These imaging results were further confirmed by drug-inhibition assays
(Fig. 4.10B). It was found that chlorpromazine at concentration of 10 µg/ml could
significantly inhibit AAV2 to infect 293T cells, whereas no inhibitory effect of filipin
(5 µg/ml) was observed, indicating that clathrin-mediated endocytosis was involved
in AAV2 entry. It also showed that AAV2 infection is low pH dependent (BAF
treatment), but macropinocytosis is not involved for AAV2 entry (Amiloide
treatment). It also showed that cholesterol on the target cell membrane is not
associated with the AAV2 infection pathway (MβCD treatment).
To monitor the interactions between AAV2 and clathrin structures in real-
time, QD-labeled viruses were incubated with hela cells expressing DsRed-clathrin,
and then live-cell imaging began suing time-lapse spinning confocal microscopy.
Selected images obtained from a time series were shown in Fig. 4.10C. The viral
particle (green) that was initially separated from clathrin signal (red) began to
colocalize with clathrin at 72 s. This colocalization was maintained for ~ 150 s, and
137
then the viral particle was separated from the clathrin signal, indicating that the virus
was dissociated from uncoated clathrin vesicles (Fig. 4.10C and 10D).

Figure 4.10: Clathrin/caveolin-dependent endocytosis of AAV2. (A) Hela cells were incubated with
QD-AAV2 (red) at 4ºC for 30 min and were then warmed to 37ºC for 5 min, fixed, permeabilized, and
immunostained with anti-clathrin (green) or anti-caveolin-1 (green) antibodies. The boxed regions are
magnified and shown in individual panels. Scale bar represents 5 µm. (B) Drug inhibition of AAV2
transduction by chlorpromazine (CPZ), filipin, bafilomycin A1 (BAF), amiloride, or methyl-β
cyclodextrin (MβCD). 293T cells were preincubated with the drug for 30 min at 37ºC. The cells (2 ×
10
5
) were then spin-infected with AAV2. Drug concentrations were maintained, except MβCD, during
the spin-infection as well as for the additional 3 h incubation, after which the drug was removed and
replaced with fresh media. The percentage of GFP+ cells was analyzed by FACS. (C) Real-time
imaging of interaction between QD-labeled virus and clathrin structures in hela cells expressing
DsRed-clathrin. (D) The fluorescent intensity of the clathrin signal associated with the viral particles.

138
4.2.10 Trafficking of AAV2 through endosomes
Next, we demonstrated that AAV2 particles traffic through various endosomal
compartments by the colocalization experiment with early endosome antigen 1
(EEA1), cation-independent mannose 6-phosphate receptor (CI-MPR), and Rab11 as
the early endosome, late endosome, and recycling endosome markersm respectively.
As shown in Fig. 4.11A and 11C, many viral particles were observed in endosomes
positive for EEA1 After 15 min of incubation at 37ºC. At 30 min, viral particles are
detected both in late (CI-MPR) and recycling (Rab11) endosomes, suggesting that
AAV2 travels at least to late and recycling endosomes confirmed by quantification of
viral particles colocalized with EEA1, CI-MPR, and Rab11 (Fig. 4.11E).
To further investigate whether the viral trafficking into the early, late, and
recycling endosomal compartments are functionally required for AAV2 transduction,
the dominant-negative mutants of Rab proteins were used to disable either the early,
Rab5, the late, Rab7, or the recycling, Rab11, endosome function. 293T cells
transfected with either the wild-type or dominant-negative form of Rab5, Rab7, or
Rab11 were incubated with AAV2. As shown in 4.11F, expression of the Rab5
dominant-negative mutant reduced the transduction rate by ~ 70% as compared to the
transduction of wild-type Rab5-expressing cells, suggesting that AAV2 must be
trafficked through early endosomes for a successful transduction. The reduced
transduction rate in 293T cells expressing Rab7 or Rab 11 dominant-negative mutant
also indicated that the productive transduction pathways are associated with the
functional trafficking to the late and recycling endosomes.
139

Figure 4.11: Trafficking of AAV2 through endosomes. (A to D) Hela cells were incubated with QD-
AAV2 (green) at 4ºC for 30 min and were then warmed to 37ºC for 15 or 30 min, fixed, permeabilized,
and immunostained with anti-EEA1 (red) and anti-CI-MPR (blue) or anti-Rab11 (blue) antibodies. The
boxed regions are magnified and shown in individual panels. Scale bar represents 5 µm. (E)
Quantification of QD-labeled AAV2 colocalized with various endosomal markers. (F) Inhibition of
virus entry by Rab5, Rab7, and Rab11 dominant-negative mutants. 293T cells transiently transfected
with wild-type or dominant-negative mutant Rab5, Rab7, or Rab 11 were spin-infected with AAV2.
The percentage of GFP+ cells was analyzed by FACS.

140
4.2.11 Cytoskeleton-mediated AAV2 transport
It is also proposed that the transport of virus to the nucleus is involved in
microtubule and/or actin-filaments with drug inhibition studies (Sanlioglu et al.,
2000). Since the drug disruption of microtubule or actin-filaments may also inhibit
trafficking of other vesicles in the cell, it has been hampered to understand the
functional involvement of the cytoskeletons in intracellular transport of AAV2. The
direct visualization of QD-labeled AAV2 viral particles in living cells suggested that
the transport of the virus involves both microtubules and actin filaments (Fig. 4.12A).
Real-time monitoring of QD-labeled AAV2 particles in hela cells expression of GFP-
tubulin suggested that mirotubule-mediated movement of AAV2 was rapid and
directional toward the nucleus (up to 2.3 µm/s) although it was only small fractions of
viral particles (~ 10%) (Fig. 4.12B and 12C). Microtubule-independent movement of
viral particles was also observed, which was likely to be diffusion-associated
transport (Fig. 4.12D and 12E).

141

Figure 4.12: Cytoskeleton-mediated viral transport. (A) QD-labeled AAV2 (blue) were incubated with
hela cells for 30 min, and the cells were then immunostained against microtubules (green) and actin-
filaments (red). The boxed regions are magnified and shown in individual panels. Scale bar represents
5 µm. (C and D) The trajectory of QD-AAV2 in hela cells expressing GFP-tubulin. (E and F) Time
trajectories of the velocity of the virus indicated in C or D, respectively.
142
4.3 Discussion
To summarize, our goal in this study was to design and evaluate a general
method to label enveloped and non-enveloped viruses with QDs. Membrane-
enveloped retroviruses were demonstrated to efficiently incorporate a biotinylation
tag (AP tag), which could be biotinylated and labeled with streptavidin-conjugated
QDs. We demonstrated this method for tagging recombinant lentivirus and gamma-
retrovirus and for tagging HIV. The viruses labeled with QDs exhibited much better
photostability than that of organic dyes. Coupled with the nature of the extremely
stable binding between biotin and streptavidin, this QD-labeling could provide a
potentially practical means to track single virus particles for prolonged periods of
time. The small size of the AP tag and site-specific attachment of QDs make this
method less likely to affect the property of viral envelope glycoproteins. Compared to
the multiple layers of antibody-based QD-labeling (Agrawal et al., 2005; Bentzen et
al., 2005), our method of direct attachment of QD-conjugates to the small biotin
molecule could introduce less perturbation to the virus. We showed that this AP tag-
based QD labeling had little effect on the viral infectivity. The kinetics of the
internalization of the recombinant lentivirus enveloped with VSVG into the early
endosomes could be studied using QD labeling and live cell imaging could be used to
monitor the trafficking of QD-tagged virus to the Rab5+ endosomal compartments.
To further demonstrate that this labeling method could be a good tool to study the
molecular mechanisms of viral entry, we did a comparative study of the clathrin- and
caveolin-dependent pathways for the internalization of two different types of
143
retroviruses and demonstrated for the first time by direct visualization that VSVG-
pseudotyped lentivirus enters cells independent of clathrin- and caveolin pathways,
while the entry of VSVG-pseudotyped retrovirus occurs via the clathrin pathway.
Importantly, the results from this imaging study of QD-labeled virus are consistent
with the drug inhibition study by us and others (Blanchard et al., 2006; Daecke et al.,
2005; Lee et al., 1999; Meertens et al., 2006), suggesting that this labeling scheme
can be reliably used for single virus tracking. Our initial studies monitoring HIV
particles using QD-labeling showed that we could detect single virions on the surface
of target cells expressing either CD4/CCR5 or DC-SIGN. Further internalization
studies of QD-HIV evidently showed that the clathrin pathway is the major route for
DC-SIGN to uptake viruses, consistent to the previous study using HIV envelope
protein (Cambi et al., 2007; Turville et al., 2004). We also demonstrated that adeno-
associated virus that is non-enveloped and one of the smallest viruses can be
efficiently labeled with quantum dots. This quantum dot labeling approach can
potentially mitigate many concerns regarding small protein labeling; a limited number
of dye molecules than can be attached to a single virus or protein without causing
self-quenching or affecting their functions. Taken together, we reported and
demonstrated a general and efficient means based on QD-labeling for detecting and
tracking live viruses during infection. We believe that this labeling can take
advantage of the excellent fluorescence property of QDs and may represent an
attractive tool for elucidating the molecular details of entry and intracellular transport
of many kinds of enveloped and non-enveloped viruses.
144
Currently, quantum dots remain more bulky than conventional organic
fluorophores, which poses difficulties for their use in labeling internal viral
components without disrupting viral functionality. Studies are underway to reduce the
size of quantum dots to improve their applicability for biological detection studies.
There has also been difficulty in purifying conjugated quantum dots, such as
functionalized or antibody-bound, from non-conjugated quantum dots through
conventional purification methods. Molecular weight cut-off (MWCO) is the method
typically used to purify molecules by filtering them to sort by size. However, since
quantum dots are large and similar in size to proteins, they cannot be distinguished
and sorted out in a mixed population with such a setup. So far, size-exclusion
chromatography has been the most promising and widely used method to purify
conjugated quantum dots. Further studies are required to determine a quicker and
more convenient method to use.

145
4.4 Materials and Methods
4.4.1 Cell lines, antibodies, reagents
The 293T/DC-SIGN cell line was generated previously in our
laboratory.(Yang et al., 2008) 293T, 293T/DC-SIGN, and HeLa cells were
maintained in a 5% CO2 environment in Dulbecco’s modified Eagle medium
(Mediatech, Inc.) with 10% FBS (Sigma), and 2 mM L-glutamine (Hyclone).
HeLa/CD4/CCR5 cell line (TZM-bl) was obtained from the AIDS Research and
Reference Reagent Program (Division of AIDS, NIAID, NIH). The cells were
maintained in D10 media with gentamycin (50 µg/ml). QD525-streptavidin conjugate
and TexasRed-labeled goat anti-mouse IgG antibody were obtained from Molecular
Probes. PE- and FITC-conjugated streptavidin were purchased from BD Bioscience.
Alexa647-conjugated anti-human CD4 antibody was obtained from Biolegend.
Mouse monoclonal antibody to human DC-SIGN was obtained from Abcam.
Monoclonal antibody against HIV-1 p24 (AG3.0) was obtained from the NIH AIDS
Research and Reference Reagent Program (Division of AIDS, NIAID, NIH).
Chlorpromazine and filipin were purchased from Sigma.

4.4.2 Cloning and expression of biotin ligase (BirA)
We followed the previously described protocol to clone and produce the biotin
ligase (BirA) from E. coli.(Chen et al., 2005) Briefly, the gene encoding BirA was
PCR-amplified from E. coli genomic DNA and cloned into pET28 expression
146
plasmid to yield pET-BirA; the BirA gene contained a C-terminal His Tag and was
under the control of the T7 promoter. The plasmid (pET-BirA) was transformed into
E. coli expression strain BL21 by a heat-shock method. The single colony was picked
and cultured in 10 ml of LB media overnight. The resulting culture was expanded into
a 1 L culture. When OD reached ~0.6, BirA expression was induced by the addition
of isopropyl-β-D-thiogalactopyranoside (IPTG) to the final concentration of 0.42
mM. After shaking at 30°C for 3 hours (h), cells were pelleted by centrifugation. The
cell pellet was resuspended in lysis buffer (50 mM NaH2PO4, 300 mM NaCl, 10 mM
imidazole, 5 mM phenylmethylsulfonyl fluoride (PMSF), pH=8.0). BirA enzyme was
then purified using Ni-NTA agarose according to the manufacture’s protocol on
native protein purification (Qiagen). Eluted fractions were subjected to SDS-PAGE
analysis and those containing BirA were pooled and further purified using an ion-
exchange PD-10 column (Amersham Biosciences) to remove the imidazole.

4.4.3 Plasmids
For the construction of plasmid AP-TM, assembly PCR was employed to fuse
the DNA sequence of AP tag (synthesized as oligonucleotide; amino acid sequences:
GLNDIFEAQKIEWHE) to C-terminus of CD5 signal peptide using a synthesized
oligonucleoitde (5’-
GGTCTGAACGATATCTTCGAAGCTCAGAAAATCGAATGGCACGAAAGATC
TGCGGATCCACCA-3’) and was PCR-amplified using the forward primer 5’-
GAATTCTGCAGATGCCCATGGGGTCTCTGCAACCG-3’ and the backward
147
primer 5’- TGGTGGATCCGCAGATCTTTCGTGC-3’. The PCR product was then
cloned into the modified pcDNA3 (Invitrogen), downstream of the CMV promoter
but upstream of the transmembrane domain of CD7 via Pst1 and BamH1. The
cDNAs for human clathrin light chain (Gene accession #: M20472) were PCR-
amplified using the forward primer 5’-
TCGAGCTCAAGCTTATGCTGAGCTGGATCCGTTCGGCG-3’ and the backward
primer 5’- GGCCCGCGGTACCTCAGTGCACCAGCGGGGCCTG-3’. Rab5
(Rab5a, Gene accession #: AF498936) were PCR-amplified using the forward primer
5’- TCGAGCTCAAGCTTATGCTAGTCGAGGCGCAACAAGACCCAAC-3’
together with the backward primer 5’-
GGGCCCGCGGTACCTTAGTTACTACAACACTGATTCCTGGTTGGTTGTGTG
G-3’. The PCR product was then cloned into the pDsRed-monomer-C1 (Clontech) via
Hind3 and Kpn1to form DsRed-clathrin and DsRed-Rab5, respectively. For the
plasmid encoding DsRed-caveolin, the cDNA for caveolin-1 (Gene accession #:
NM_001753) was PCR-amplified using the forward primer 5’-
GAGCTCAAGCTTATGTCTGGGGGCAAATACGTAGACTCGGAG-3’ and the
backward primer 5’-
ACCGGTGGATCCATTTCTTTCTGCAAGTTGATGCGGACATTGC-3’ and then
inserted into the plasmid pDsRed-monomer-N1 (Clontech) via Hind3 and BamH1.
Production of AP-tagged virus. AP-tagged and pseudotyped lentiviruses were
produced by transient transfection of 293T cells using a standard calcium phosphate
precipitation method. 293T cells at 80% confluence in 6-cm culture dishes were
148
transfected with 5 µg of the lentiviral plasmid FUW, together with 2.5 µg each of AP-
TM, the envelope plasmid (VSVG or HIV gp160) and the packaging plasmids
(pMDLg/pRRE and pRSV-Rev). For production of AP-tagged retroviral viruses,
293T cells were transfected with 5 µg of the retroviral plasmid MIG, (Yang and
Baltimore, 2005) along with 2.5 µg each of AP-TM, the envelope plasmid (VSVG)
and the packaging plasmid (gag-pol). The viral supernatant was collected after 48-h
posttransfection, filtered through a 0.45-µm pore size filter, and then concentrated by
ultracentrifugation (Optima L-90 K ultracentrifuge, Beckman Coulter) either for 90
min at 82,700 × g for VSVG-pseudotyped viruses or for 60 min at 50,000 × g for HIV
virus. The pellets were then resuspended in an appropriate volume of cold PBS
containing 5 mM MgCl2.

4.4.4 Biotinylation and QD-labeling of virus
The concentrated viruses in PBS-MgCl2 were incubated with 2.5 µM BirA,
10 µM of biotin, and 1 mM ATP for 60 min at 4°C, followed by incubation with 30
nM of QD525-streptavidin for 60 min at room temperature. Viral aggregates were
removed with 0.45 µm pore size filters before imaging.

149
4.4.5 Viral transduction
293T cells (0.2 × 106 per well) were plated in a 24-well culture dish and spin-
infected with viral supernatants (1 ml per well) at 2,500 rpm and 30°C for 90 min by
using a Sorval Legend centrifuge. Then, the medium was removed and replaced with
fresh medium and cultured for 3 days before FACS analysis of GFP+ cells. For viral
transduction with drug-treated cells, 293T cells were pre-incubated with drugs
(chlorpromazine: 10 and 25 µg/ml; filipin: 1 and 5 µg/ml) for 30 min at 37°C and
then the cells (0.2 × 106 per well) were spin-infected with 1 ml of viral supernatants
in a 24-well culture dish. The drug concentration was maintained during the spin-
infection. The cells were further incubated for 60 min at 37°C. The drugs were then
removed and replaced with fresh D10 media.
For the production of adeno-associated virus 2, HEK 293 cells were triple
transfected with ΔF6 (adenoviral helper plasmid), an AAV cis plasmid, and an AAV
trans plasmid. At 60 h posttransfection, cells were harvested, and then viruses were
purified by cesium chloride gradient density centrifuging for 20 h at 25,000 rpm with
Beckman SW-28 rotor. The viral fraction was desalted with Amicon Ultra-15
centrifugal concentrator (Millipore, MWCO: 100kd)

4.4.6 Quantum dot-AAV2 network
Quantum dot-AAV2 networks were produced by covalent amide bonds
formed by carbodiimide chemistry between carboxylic moieties on the quantum dots
150
and the primary amines from the lysine residues on the virus capsid protein. The
carboxyl group on the quantum dots were incubated with 1-ethyl-3-[3-
dimethylaminopropyl]carbodiimide hydrochloride (EDC) and N-hydroxysuccinimide
(NHS) to modify the carboxyl group to an amine-reactive NHS ester. Then, excess
EDC and NHS were removed by a gel filtration column and the primary amine
source, AAV2, was added for a coupling reaction for 1 h at room temperature. The
reaction was quenched by adding 10M Tris buffer pH 7.4. Uncoupled quantum dots
were removed by using HiTrap Heparin column (GE Health).

4.4.7 Confocal imaging
Fluorescent images were acquired on a Zeiss LSM 510 META laser scanning
confocal microscope equipped with Argon, red HeNe and green HeNe lasers as well
as attached to a Coherent Chameleon Ti-Sapphire laser for multiphoton imaging.
Images were acquired using a Plan-apochromat 63×/1.4 oil immersion objective.
Some images were collected by a Yokogawa spinning-disk confocal scanner system
(Solamere Technology Group, Salt Lake City, UT) using a Nikon eclipse Ti-E
microscope equipped with a 60×/1.49 Apo TIRF oil objective and a Cascade II: 512
EMCCD camera (Photometrics, Tucson, AZ, USA). For the detection of individual
viral particles, QD-labeled viruses were overlaid upon polylysine-coated coverslips
for 60 min at 37°C. The coverslips were then rinsed, fixed with 4% formaldehyde,
permeabilized, and immunostained with monoclonal antibody specific to p24 capsid
protein. The coverslips were mounted in Vectashield (Vector Laboratories), which is
151
an antifade mounting medium. Images were analyzed with the use of the Zeiss LSM
510 software version 3.2 SP2.
For photostability comparisons of fluorescent dye and QD-labeled viral
particles, viral particles were labeled with FITC-streptavidin (20 µg/ml) or QD525-
streptavidin (30 nM). The labeled viruses were overlaid upon polylysine-coated
coverslips for 60 min at 37°C. The viruses were then continuously exposed to the
Argon laser over 3 min. Images were captured at ~10 seconds (s) intervals.
Fluorescence intensity versus time within the regions of interest was measured by
using the Zeiss LSM 510 software package.
For the viral trafficking studies using clathrin, caveolin-1, and Rab constructs,
293T, 293T/DC-SIGN, or hela cells were transfected with individual plasmids
encoding either DsRed-clathrin, DsRed-caveolin1, or DsRed-Rab5. At 24 h
posttransfection, cells were seeded onto glass-bottom culture dishes and grown at
37°C overnight. QD-labeled viruses were incubated with cells for 30 min at 4°C to
synchronize infection. The cells were shifted to 37°C for the different time periods,
and then fixed with 4% formaldehyde.
To visualize the interaction between HIV virus and target cells,
Hela/CD4/CCR5 or 293T/DC-SIGN cells were incubated with QD-labeled HIV virus
for 30 min at 4°C, fixed, and then immunostained with anti-human CD4 antibodies or
anti-human DC-SIGN and counterstained with DAPI.

152
For the real-time observation of colocalization of the labeled virus with early
endosomes or clathrin, QD-labeled viruses (FUW/VSVG+AP+QD or QD-AAV2)
were incubated with cells for 30 min at 4°C to allow virus binding. The cells were
then warmed to 37°C for various time points to induce viral internalization, and
confocal time-lapse images were then recorded.

153

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Abstract (if available)

Abstract A strategy to target lentiviral vectors to specific cell types holds great promise for future clinical applications of gene therapy. We have previously developed an efficient method to target lentivirus-mediated gene transduction by introducing a targeting antibody and pH-dependent fusogenic protein as two distinct molecules on the lentiviral surface. However, the molecular mechanism that controls the targeted infection needs to be defined. To elucidate the endocytic pathway of the engineered lentivirus, we have monitored intracellular trafficking of the individual lentiviruses in the targeted cells by various direct visualization approaches. This study proposed that the fusogen-mediated membrane fusion could be a rate-limiting step of targeting lentivirus transduction. However, the specific features of the fusogen-associated membrane fusion that control the targeted infection still remain largely unknown. Therefore, we further demonstrated the intracellular behaviors of two engineered lentiviruses displaying a class I fusogen derived from Sindbis virus glycoprotein or a class II fusogen derived from influenza virus hemagglutinin by tracking the individual viral particles in target cells. Our results suggest that both engineered lentiviruses enter target cells through clathrin-dependent endocytosis. However, the different kinetics of virus-endosome fusion as well as the distinct requirement of endosomal traffic for viral fusion of two engineered lentiviruses was suggested by imaging multiple sequential steps of fusion event in target cells. These imaging studies shed some light on the infection mechanism of the engineered lentivirus and is beneficial to the design of more efficient gene delivery vectors.

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Study of dendritic cell-specific lentivector vaccine system

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Study of dendritic cell targeting by engineered lentivectors

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Entry and intracellular trafficking of measles virus glycoprotein pseudotyping lentivector for transduction in target cells

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Engineering nanoparticles for gene therapy and cancer therapy

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Engineering viral vectors for T-cell immunotherapy and HIV-1 vaccine

Asset Metadata

Creator Joo, Kye Il (author)

Core Title Dissecting the entry mechanism of targeting lentiviral vectors in living cells and developing quantum dot labeling of viruses for single virus tracking

School Andrew and Erna Viterbi School of Engineering

Degree Doctor of Philosophy

Degree Program Chemical Engineering

Publication Date 10/05/2009

Defense Date 09/10/2009

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

Tag gene delivery,intracellular trafficking,lentiviral vectors,OAI-PMH Harvest,quantum dots,single virus tracking

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Wang, Pin (committee chair), Hamm-Alvarez, Sarah F. (committee member), Shing, Katherine S. (committee member)

Creator Email kijoo75@gmail.com,kjoo@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m2643

Unique identifier UC1492679

Identifier etd-Joo-3292 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-258648 (legacy record id),usctheses-m2643 (legacy record id)

Legacy Identifier etd-Joo-3292.pdf

Dmrecord 258648

Document Type Dissertation

Rights Joo, Kye Il

Type texts

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

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu