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Engineering nanoparticles for gene therapy and cancer therapy
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Engineering nanoparticles for gene therapy and cancer therapy
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Content
ENGINEERING NANOPARTICLES FOR GENE THERAPY AND
CANCER THERAPY
by
Yarong Liu
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMICAL ENGINEERING)
May 2014
Copyright 2014 Yarong Liu
ii
Dedication
This thesis is dedicated to my family.
iii
Acknowledgements
First, I wish to thank my advisor Dr. Pin Wang, who has supported and guided me
throughout the years. His understanding, patience, and more importantly, his
friendship during my graduate studies help me to reach my full potential. He
encouraged me to do experiments independently and fully supported me when I was
pregnant. For everything you’ve done for me, Dr. Wang, I thank you. I would also like
to thank my Dissertation committee Dr. Katherine Shing and Dr. Don Arnold for their
suggestion and support.
I am very thankful for all the help and fun brought by the group members in RTH-515
at USC, Yuning Lei, April Tai, Chi-lin Lee, Bingbing Dai, Biliang Hu, Steve Froelich,
Paul Bryson, Chupei Zhang, Man Ji, Jinxu Fang, Natnaree Siriwon, Yu-Jeong Kim and
Jennifer Rohrs. I would like to show my special thanks to Kye-il Joo for tutoring me
when I joined the lab. It was also my pleasure to work with all the other lab members
for their support of my work.
I would also like to show my thanks to the professors and staffs in Chemical
Engineering department. Thanks to Dr. Steven Nutt for lots of valuable information he
provided and full support when I applied for scholarship.
Last but not the least, this dissertation is dedicated to my family for their unconditional
love and support. Without my husband Chuanxi, I could not reach this far in my Ph.D.
study. Thanks for taking care of Andrew and me. You and Andrew make my life filled
with joy and love.
iv
Table of Contents
Dedication………………………………………………………………………………...ii
Acknowledgements……………………………………………………………………....iii
List of Figures……………………………………………………………………...........viii
List of Tables……………………………………………………………………................x
Abstract……………………………………………………………………........................xi
Chapter 1 Introduction 1
1.1. Gene Therapy………………………………………………………….........................2
1.1.1 Viral vector-mediated gene delivery...…………………………..........................2
1.1.2 Adeno-associated viruses (AAV) vector in gene delivery……………………..3
1.1.3 Increasing transduction efficiency and specificity AAV2 targeting vectors….5
1.1.3.1 Cellular barriers limiting viral transduction……………………………..5
1.1.3.2 Site-specific modification…………………………………………………8
1.1.3.3 Cell-permeable peptides…………………………………………………10
1.2. Cancer therapy 11
1.2.1 Nanomedicine in cancer therapy……………………………………………...11
1.2.2 Main issues of liposomes for drug delivery…………………………………..13
1.2.3 Co-delivery of chemotherapeutics to induce synergistic effect……………...15
1.2.4 Co-delivery of chemotherapeutics to overcome multi-drug resistance……..17
v
Chapter 2 Site-specific Modification of Adeno-Associated Viruses via a Genetically
Engineered Aldehyde Tag 20
2.1. Abstract…………………………………………………………………....................21
2.2. Introduction………………………………………………………………................22
2.3. Results and Discussion……………………………………………………...............24
2.3.1 Confirming and visualizing aldehyde tag on modified AAV nanoparticles...24
2.3.2 Evaluate the effect of modification on AAV function………...........................30
2.3.3 Enhanced transduction of AAV2
Ald13
by conjugating with targeting ligands..37
2.4. Materials and Methods……………………………………………………...............40
Chapter 3 Enhancing Gene Delivery of Adeno-Associated Viruses by Cell-Permeable
Peptides 49
3.1. Abstract…………………………………………………….........................................50
3.2. Introduction……………………………………………..............................................51
3.3. Results……………………………………………........................................................53
3.3.1 Enhanced AAV2 transduction mediated by CPPs..............................................53
3.3.2 Enhanced viral uptake with faster kinetics by CPPs...........................................56
3.3.3 Entry mechanism of AAV2-CPP complexes.......................................................58
3.3.4 Involvement of endosomes in viral transduction mediated by AAV2-CPP
complexes………………………………................................................................60
3.3.5 CPPs enhance viral transduction of AAV2 in primary cells and tissues…….65
3.3.6 CPPs enhance AAV2-mediated gene delivery in vivo………………………..67
vi
3.3.7 CPPs increase viral transduction of AAV8 both in vitro and in vivo…………72
3.4. Discussion……………………………….......................................................................73
3.5. Materials and Methods………………...........................................................................77
Chapter 4 Combinatorial Drug Delivery of Doxorubicin and Paclitaxel via cMLVs
Enables Synergistic Antitumor Activity 87
4.1. Abstract…………………………………………………….........................................88
4.2. Introduction……………………………………………….........................................89
4.3. Materials and Methods…………………………………….........................................92
4.4. Results and Discussions……………………………………........................................99
4.4.1 Characteristics of combinatorial drug delivery via cMLVs................................99
4.4.2 In vitro analysis of doxorubicin: paclitaxel for drug ratio-dependent
synergy…………………………………………………………………………102
4.4.3 Drug ratio-dependent efficacy of cMLV(Dox+PTX) in tumor treatment…106
4.4.4 Drug ratio-dependent efficacy of co-encapsulated Dox:PTX on tumor
apoptosis……………………………………………………………………….107
4.4.5 In vivo cardiac toxicity evaluation of drug combinations in cMLV
formulations…………………………………………………………………...108
4.4.6 In vivo maintenance of drug ratios in cMLV formulations………………...110
4.5. Conclusion…………………………………………………………………............113
vii
Chapter 5 Co-delivery of Doxorubicin and Paclitaxel via Crosslinked Liposomal
Formulations To Overcome Multidrug Resistance in Tumor………….....................114
5.1. Abstract…………………………………………………….......................................115
5.2. Introduction………………………………………………........................................116
5.3. Materials and Methods…………………………………...........................................118
5.4. Results……………………………………………………..........................................125
5.4.1 In vitro efficacy study by XTT assay………………..........................................125
5.4.2 Cellular uptake study of doxorubicin and paclitaxel........................................127
5.4.3 Effect of codelivery nanoparticles on P-gp expression.....................................129
5.4.4 Efficacy of dual drug-loaded cMLVs against a breast cancer model………..131
5.4.5 Histology study……………………………………….......................................133
5.5. Discussion………………………………………........................................................137
5.6. Conclusion……………………………………...........................................................139
References……………………………………....................................................................141
viii
List of Figures
Figure 1. Map of the wild-type AAV2 genome.
Figure 2. Wild type AAV life cycle.
Figure 3. Stages of rAAV transduction.
Figure 4. Schematic representation of EPR effect by which nanoparticles can deliver
drugs to tumors.
Figure 5. AAV2 nanoparticles can be site-specifically modified by genetically encoded
aldehyde tags.
Figure 6. Covalent attachment of amine-functionalized gold (Au) particles onto
AAV2Ald13 nanoparticles and characterization of Au-labeled AAV2
Ald13
nanoconjugates.
Figure 7. The hydrodynamic size of unconjugated AAV2Ald13 and Au-AAV2
Ald13
measured by dynamic light scattering.
Figure 8. Quantitative analysis of the number of attached dyes per AAV2
Ald13
particle.
Figure 9. The expression and chemical modification of aldehyde tag on AAV2 capsid
proteins.
Figure 10. LC/MS/MS spectrum of the peptide fragment QTGLFGlyTPSR from
AAV2
Ald13
.
Figure 11. Comparison of nanoparticle titers and functional titers of AAV2
WT
and
AAV2
Ald13
.
Figure 12. Intracellular trafficking of AAV2
Ald13
in HeLa cells.
ix
Figure 13. Targeting of anti-HLA-conjugated AAV2
Ald13
nanoparticles to receptor-
bearing cells.
Figure 14. Targeted gene transduction mediated by antibody-conjugated AAV2
Ald13
vectors.
Figure 15. Targeting of peptide-conjugated AAV2Ald13 nanoparticles to tumor cells.
Figure 16. Antp, TAT-HA2 and LAH4 improve AAV2 transduction in permissive and
non-permissive cells.
Figure 17. Interaction of CPPs with AAV2 facilitates enhanced viral transduction.
Figure 18. CPPs enhance AAV2 uptake by cells.
Figure 19. Entry mechanisms of AAV2-CPP complexes.
Figure 20. The effect of CPPs on the endosomal escape of AAV2 particles.
Figure 21. CPPs enhance viral transduction in primary cells and tissues.
Figure 22. CPPs facilitate gene delivery of AAV2 in mouse muscles.
Figure 23. No cytotoxicity of CPPs was detected in vivo.
Figure 24. Antp, TAT-HA2 and LAH4 improve AAV8 transduction in target cells.
Figure 25. CPPs facilitate AAV8-mediated gene delivery in mouse muscles.
Figure 26. Characteristics of cMLV (Dox+PTX).
Figure 27. Determination of the ratio of drug combinations to induce synergy.
Figure 28. IC50 values of Dox and PTX in cMLV formulation or free drug solution in
B16 melanoma or 4T1 breast tumor cells.
Figure 29. Drug ratio-dependent efficacy of cMLV(Dox+PTX) in tumor treatment.
x
Figure 30. Drug ratio-dependent efficacy of co-encapsulated Dox:PTX on tumor cell
apoptosis.
Figure 31. In vivo toxicity.
Figure 32. In vivo maintenance of Dox:PTX ratios in cMLV formulations.
Figure 33. Overcoming drug resistance by codelivery of Dox and Taxol via MLVs.
Figure 34. Celluar uptake of doxorubicin and paclitaxel.
Figure 35. Effect of codelivery nanoparticles on P-gp expression.
Figure 36. In vivo efficacy of drug combinations via cMLVs against 4T1 tumor model.
Figure 37. Effect of co-delivery cMLVs on tumor apoptosis.
Figure 38. Effect of co-delivery cMLVs on P-gp expression in tumros.
List of Tables
Table 1. Representative examples of nanocarrier-based drugs on the market.
Table 2. Amino acid sequences of cell-permeable peptides used in this study
xi
Abstract
Virus-based nanoparticles have shown promise for mediating gene delivery due to
their well-defined nanostructure and intrinsic bioactive functionality. Adeno-
associated virus (AAV) has been considered as a promising vehicle for human gene
therapy based on its ability to infect both dividing and nondividing cells, as well as
establish long-term gene expression in vivo without known pathological consequence
of infection. However, because of their native tropisms, the applicability of AAV
nanoparticles is often limited to the restricted ranges of cells or tissues. Studies
proposed that low expression of receptors/coreceptors on cell surface and an impaired
intracellular trafficking pathway of vectors could be the rate-limiting steps of AAV-
mediated transduction. In this study, we have developed two strategies for enhancing
AAV-mediated gene delivery by overcoming these two biological barriers. The first
strategy we used is generating a targeted AAV2 vector by genetically encoding an
aldehyde tag on viral capsids. Such a tag can be exploited for site-specifi c attachment
of targeting molecules and allows for further introduction of targeting antibodies or
ligands. The results showed that the site-specific conjugation of targeting antibodies
could significantly enhance viral transduction to those target cells that have otherwise
exhibited very low permissiveness to AAV2 infection. This method also allows the
functional incorporation of RGD peptides onto AAV2 for enhanced delivery with
implications for cancer gene therapy. Another strategy we used to enhance AAV
transduction, both in vitro and in vivo, is incubating AAV vectors with cell-permeable
xii
peptides (CPPs). We show that CPPs increase internalization of viral particles into
cells by facilitating both energy-independent and energy-dependent endocytosis.
Moreover, CPPs can significantly enhance the endosomal escape process of viral
particles, thus enhancing viral transduction to those cells that have exhibited very low
permissiveness to AAV2 infection as a result of impaired intracellular viral processing.
We also demonstrated that this approach could be applicable to other AAV serotypes.
Non-viral based nanoparticles offers new hope for cancer detection, prevention and
treatment due to their potentials to deliver drugs to tumors. Liposomes constitute one
of the most popular nanocarriers for the delivery of cancer therapeutics. However,
since their potency is limited by incomplete drug release and inherent instability in the
presence of serum components, their poor delivery occurs in certain circumstances. In
this study, we address these shortcomings and demonstrate an alternative liposomal
formulation, termed crosslinked multilamellar liposomal vesicles (cMLVs). With its
properties of improved sustainable drug release kinetics and enhanced vesicle stability,
cMLVs can achieve controlled delivery of cancer therapeutics. We further showed that
the cMLVs are potential in combination therapy by loading drugs with different
hydrophilicities into the same cMLV in a precisely controllable manner over drug
ratios. The stability of cMLVs allows an improved loading efficiency and sustained
release of doxorubicin (Dox) and paclitaxel (PTX), maximizing the combination
therapeutic effect and minimizing the systemic toxicity. Furthermore, in vivo
experiments showed that the robust cMLV formulation maintains drug ratios for
xiii
prolonged times, enabling the ratio-dependent combination synergy translating from
in vitro to in vivo antitumor activity. We also demonstrated that this combinatorial
delivery system (cMLV(Dox+PTX)) achieves enhanced drug accumulation and
retention, resulting in improved cytotoxicity in tumor cells, including drug resistant
cells. Moreover, cMLV(Dox+PTX) significantly overcomes MDR by reducing the
expression of P-glycoprotein (P-gp) in cancer cells, thus improving antitumor activity
in vivo. With a combined therapeutic ability that enhances drug delivery efficacy to
tumors and lowers the apoptotic threshold of individual drugs, cMLV(Dox+PTX)
represents a potential multimodal therapeutic strategy to overcome MDR in cancer
therapy.
1
Chapter 1
INTRODUCTION
2
1.1 Gene Therapy
1.1.1 Viral vector-mediated gene delivery
Gene therapy is broadly defined as the introduction of nucleic acid, either RNA or
DNA, into a target cell or tissue to prevent a disease. And it has been increasingly
considered as the most promising method to treat or eliminate the cause of disease by
insertion of functional genes into a target cell [1,2,3]. Although it has been reported
uptake of “naked” DNA occurred in a few tissues such as muscle, the limited efficiency
of delivery and expression raised problems. To improve the efficiency and stability of
gene delivery, many efforts are made on developing nanoparticles including viral and
nonviral vectors for gene transfer. Nonviral vectors such as liposomes have been
developed as a carrier to deliver DNA into cells due to its advantages of no insert-size
limitation and nonpathogenic. However, they have the notable disadvantage of low
transfection efficiency [1,4].
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 (Rv), adenovirus (Ad)
and adeno-associated virus (AAV). Many retroviral vectors have been used in clinical
use due to their capability of stably integrating into the host DNA in the infected cell
and expressing a therapeutic gene for the life of that cell. However, current retroviral
vectors are better suited for ex vivo than in vivo gene therapy because they require cell
division for integration [4]. Another disadvantage of retroviral vectors is the random
3
insertion into the host genome, possibly leading to oncogene activation or tumor-
suppressor gene inactivation. Adenoviruses attracted attention as gene delivery vectors
because they can efficiently infect and express their genes in both dividing and non-
dividing cells. And their genome is 36kb in length and is easy to manipulate using
classical recombinant DNA techniques [5]. However, it has notable disadvantages of
limited duration of transgene expression and immunogenicity in vivo. Adeno-
associated viruses (AAV) have attracted considerable interests in gene therapy due to
their capability of infecting both dividing and nondividing cells to establish substantial
transfection and long-term gene expression without any known pathological
consequence of infection [6,7,8].
1.1.2 Adeno-associated viruses (AAV) vector in gene delivery
Adeno-associated virus harbors a single-stranded genome approximately 4.7 kb long,
which contains two open reading frames (rep and cap) flanked by inverted terminal
repeat elements (ITRs)[9], shown in Figure 1A. The cap gene encodes for three
structural capsid proteins (VP1, VP2, and VP3) that share the same C-terminal
domain, while VP1 and VP2 contain additional N-terminal sequences. Sixty total
copies of these three capsid proteins VP1, VP2 and VP3 at a molar ratio of
approximately 1:1:10 self-assemble to form the viral capsid. Thus cap plays a great role
in the viral gene transduction properties.
4
Figure 1. Map of the wild-type AAV2 genome. (A) Rep and Cap genes flanked by ITRs.
The wild type AAV genome is capable of integrating into a specific locus on human
Chromosome 19. However, the replication of wild type AAVs requires sequences
provided in trans by a helper virus, such as adenovirus or HSV (Fig 2A, AAV life
cycle). Although the molecular mechanisms of the integration remain unclear, the
sustained gene expression of AAV vectors is a major advantage in treating chronic
diseases. The long-lasting transgene expression has been reported in lung, muscle, liver
and the central nervous system in mice. This is at least partly contributed by an
undetectable cytotoxic-T-lymphocyte (CTL) response to AAV-transfected cells.
Although AAV show a great promise in gene therapy, only a few clinical protocols
involving AAV have been initiated, mainly because of manufacturing difficulties.
Recombinant AAV (rAAV) vectors are constructed by co-transfection of two
plasmids, but a helper virus is still required for rAAV infection. Alternatively, a
strategy was proposed by using a plasmid to provide helper virus-like functions in
5
trans [10] . The improved manufacturing processes allowing high-titer production
without a helper virus increase more AAV-mediated gene therapies in clinical trials.
Different serotypes of AAVs have been shown to share a common genome structure,
but have unique capsid proteins that can be recognized by different cell surface
receptors [11]. To date, more than 100 distinct primate AAV capsid sequences have
been isolated [12], and 12 primate serotypes of AAVs (AAV1 to AAV12) have been
evaluated as recombinant gene therapy vectors [13]. Although AAV2, a relatively well-
characterized AAV serotype, has been extensively investigated in clinical trials [6,14],
many alternative serotypes are currently in preclinical and clinical development due to
their different tissue tropisms and enhanced gene transfer efficiency [8,15,16].
1.1.3 Increasing transduction efficiency and specificity of AAV2 vectors
1.1.3.1 Cellular barriers limiting viral transduction
The AAV2 vector was the first primate AAV to be cloned, and promising results have
been obtained with this vector in clinical gene transfer, including retinal degenerative
disorders [14,17,18] and haemophilia B [19,20]. However, the transduction of AAV2 is
inefficient in a number of nonpermissive cell types [21,22,23,24], limiting its
application in many areas of gene therapy. As shown in Figure 3, several biologic
barriers that limit the effectiveness of AAV2 for gene therapy have been proposed
based on the AAV2 infectious process [25].
6
Figure 2. (A) Wild type AAV life cycle. (B) Schematic representation of rAAV production.
First, AAV2 must bind to the surface of target cells through receptor and coreceptor.
Thus, the most significant factor affecting this stage includes the abundance of AAV2
receptors and /or coreceptors [25]. Heparan sulfate proteoglycan (HSPG) is known to
be one of the primary attachment receptor for binding of AAV2 to the surface of many
cell types [26,27]. In addition to attachment receptors, AAV2 also requires several
coreceptors for efficient internalization to cells, including αVβ5 integrin and fibroblast
growth receptor 1 [28,29]. It has been reported that the limited expression of receptor
7
and/or coreceptors on the cell surface of nonpermissive cells, significantly prevents
binding of the viral vector to the cell surface [30,31,32].
Figure 3. Stages of rAAV transduction.
After AAV internalize into cells through endocytosis, the multi-step intracellular
events (stage 3-5) is considered as barriers to AAV transduction [25]. For example, it
has been reported that AAV2 vectors bind efficiently to murine fibroblast cells and are
internalized successfully, but the trafficking to nucleus is significantly impaired, thus
limiting transgene expression in the cells [24]. Furthermore, a comparative study
showed that a remarkably low efficiency of AAV2 transduction was obtained in
endothelial cells compared to that of highly permissive cell types, even though the
receptors for AAV2 are expressed in both situations [33]. This study revealed that
8
AAV2 virions tended to accumulate in the Golgi area of permissive cell lines, but not
endothelial cells, indicating that the intracellular trafficking of the virus can impact its
transduction efficiency. It has become increasingly recognized that successfully
processing of AAV virions through the endosomal compartment is essential to
determine the efficacy of AAV transduction. For AAV2, it has been proposed that
processing through endosomal compartments, including early and late endosomes
[30,34,35], is required to induce a conformational rearrangement of the viral capsid for
nuclear transport and uncoating [36]. Moreover, it has been reported that a limited
AAV2-mediated transduction was attributed to the impaired endosomal escape
process in NIH3T3 cells [24], a non-permissive cell line for AAV2 vectors. Therefore,
overcoming these cellular barriers with a novel method of AAV2 infection would
greatly increase the applicability of AAV2 vectors in gene therapy.
1.1.3.2 Site-specific modification of capsid proteins to enhance viral transduction
Retargeting the viral tropism of AAV2 to cells that are normally resistant to AAV2
infection could increase transduction efficiency, thus expanding its utility for gene
delivery. However, while targeting AAV2 vectors to specific tissues of interest is highly
desirable, it is also a challenging task. Nonetheless, many attempts have been made to
generate targeted AAV2 vectors by modifying virus-cell interaction, which is generally
hypothesized as an essential step in virus infection. Significant effort has also been
devoted to the alteration of capsid proteins, which are responsible for virus binding to
cell surface receptors and mediating cellular entry [37,38,39].
9
Generally, engineering viruses to enhance the targeted specificity is focus on genetic
modification and chemical modification. Currently, the majority of targeting efforts
rely upon genetic modification of AAV capsid proteins by insertion of targeting
peptide motif to encode ligands or antibodies that can direct vectors to specific cell
types. For example, genetically displaying retargeting peptides on virus surface has
been successfully employed to retarget AAV to arterial endothelium [40], striated
muscles [41], and brain basculature [42]. However, the major technical challenges in
this manipulation include the low production yield, dramatic reduction of vector titer,
or significant drop of DNA packaging efficiency [40,43]. In order to sidestep the issues
of virion assembly and size limit for insertion of targeting peptides, chemical
modification methods have been explored. Chemical manipulation utilizes the
reactivity of functional groups, such as the amino group of lysine and the thiol group
of cysteine, which have the potential to introduce targeting ligands able to attach virus
to specific cells. However, random modification of lysine residues can lead to
inactivation of the protein [44]; therefore, the lysine modification of the viral capsid
could potentially alter virus-host cell interaction and/or cause a loss of viral infectivity.
Another drawback to this approach is the requirement of extensive prior purification,
which is time-consuming and occasionally ineffective. Additionally, chemical
modifications are dependent on naturally occurring residues, leading to a potential loss
of precision. Therefore, site-specific modification is highly desirable for both labeling
and targeting of virus vector.
10
1.1.3.3 Cell-permeable peptides
An alternative way to increase cell-surface interactions of viral particles may be
through cell-permeable peptides (CPPs). CPPs, small poly-basic peptides, show a
superior ability to penetrate cell membranes and have been used to deliver biologically
active substrates such as proteins [45], plasmid DNAs [46], liposomes [47] and viruses
[48] into various cell types and tissues. The main feature of CPPs, with a net positive
charge at physiological pH [49], enables them to penetrate the cell membrane at low
micromolar concentrations both in vitro and in vivo without using any chiral receptors
and without causing significant membrane damage [50]. For example, the third α-
helix of Antp homeodomain or an 11-amino acid motif from HIV Tat protein, have
been shown to be able to cross the cell membrane through a receptor-independent
mechanism [51,52]. Thus, these cell-permeable peptides show potential to increase the
efficiency of the initial binding step of viral vectors in cells, allowing substantially
enhancing viral-mediated gene transfer.
Moreover, CPPs show promise as an alternative strategy to chemical reagents that
promote endosomal membrane disruption [53,54] because CPPs can promote the
destabilization of the endosomal membrane upon acidification of the endosomal
compartment [55] without significant toxicity. For instance, arginine-rich and
histidine-rich peptides have shown the ability to disrupt the endosomal membrane via
a buffering effect within the acidic endosomal environment [56,57,58]. Therefore,
11
CPPs might enhance AAV2 mediated transduction in various cell types by overcoming
these two major obstacles.
1.2. Cancer Therapy
1.2.1 Nanomedicine in cancer therapy
Although cancer remains one of the most devastating diseases in the world, recent
advances in nanotechnology have offered new hope for cancer detection, prevention
and treatment. Nanomedicine for cancer therapy is advantageous over conventional
medicine because it has the potential to enable the preferential delivery of drugs to
tumors owing to the enhanced permeability and retention (EPR) effect [59,60,61,62].
As shown in Figure 4, nanoparticles can escape into the tumor tissues via the leaky
blood vessels and accumulate in poor lymphatic drainage in tumor regions via EPR
effect, allowing the release of chemotherapeutics into tumor cells.
Other advantages of nanocarriers in drug delivery include protecting the drugs from
premature degradation, controlling the pharmacokinetics and drug tissue distribution
profile, and improving intracellular penetration by conjugating with targeting ligands
or peptides. Moreover, it has been increasingly attractive to develop nanocarriers with
an extended circulating half-life in serum and more controlled release profile. Over 20
nanoparticles for delivering therapeutics have been approved by the FDA for clinical
use, including polymer-drug conjugates, lipid-based carriers and so on
[59,60,63](Table 1).
12
Active cellular
targeting
Ligand
Drug
Receptor
Nuc leus
iii
iii
Blood
vessel
Endothelial cell
Passive tissue targeting Nanopartic le
EPR e!ect
Angiogenic
vessels
Ine!ective
lymphatic drainag e
Tu mour
Lymphatic vessel
Enl argement of tu m our cell
x
Figure. 4. Schematic representation of EPR effect by which nanoparticles can deliver drugs to
tumors.
Table 1. Representative examples of nanocarrier-based drugs on the market.
13
Among the nanocarriers, polymers were one of the earliest reported materials for
cancer therapy[59]. Recently, a nanoparticle formulation of paclitaxel, in which
albumin is included as a carrier and has been applied in the clinic for the treatment of
metastatic breast cancer as shown in Table 1. However, their inherent structural
heterogeneity has been concerned[59]. It has been more and more attractive to develop
lipid-based carrier because of their composition, which makes them biocompatible and
biodegradable [59]. Moreover, lipid-based carriers have the capability of delivering
drugs with different lipophilicities by encapsulating strongly lipophilic drugs in lipid
bilayer and strongly hydrophilic drugs in the aqueous compartment. Another
attraction lies in the easy alternation of their properties, such as size, surface chemistry
and so on. Currently, several kinds of cancer drugs have been approved to use in
cancer treatment by using lipid-based system. As shown in Table 1, liposomal
formulations of doxorubicin (Doxil, Myocet) and daunorubicin (DaunoXome) have
been approved for the treatment of metastatic breast cancer, ovarian cancer and
Kaposi’s sarcoma [59].
1.2.2 Main issues of liposomes in drug delivery
Phospholipid vesicles (liposomes) constitute a class of nanoparticulate drug carriers
generally characterized by composition of one or more phospholipid bilayer
membranes, and capability of delivering aqueous or lipid drugs [64]. Based on their
ability to encapsulate both hydrophilic and hydrophobic drugs, liposomal
formulations of anticancer drugs have been extensively evaluated for treating cancers
14
[60]. Among the many benefits of liposomal delivery of anthracyclin, e.g., doxorubicin
or daunorubicin, compared with the administration of the fee drug, is reduced cardiac
toxicity with remaining therapeutic efficacy to tumors [65,66].
Although it has been reported that liposomes are promising drug carriers for cancer
therapeutics, the inherent instability of conventional unilamellar liposomes in the
presence of serum components, which is apparently related to their drawback of a
rapid drug release profile, have limited their utility for delivering anticancer agents
[64]. It has been shown that the release rate of liposomal Dox is intrinsically linked to
the toxicity level and therapeutic activity. Briefly, slower release rates usually result in
lower toxicity and higher therapeutic efficiency. To overcome these challenges, a
PEGylated liposomal drug formulation was developed by using an ammonium sulfate
gradient loading procedure, enabling stable drug entrapment into liposomes and their
extended blood-circulation time. For example, pegylated liposomal doxorubicin has
been approved for the treatment of Kaposi’s sarcoma, ovarian cancer and metastatic
breast cancer [59]. Indeed, such liposomal drug formulations do appear to improve
accumulation of liposomes at the tumor site. However, slow and incomplete drug
release could still lead to low drug bioavailability within tumor tissue, limiting, in turn,
therapeutic activity [67,68]. Furthermore, a lack of controlled-release properties of
encapsulated drug may lead to toxic side effects, such as palmar-plantar
erythrodysesthesia that is thought to result from unwanted drug distribution to skin
during prolonged circulation of liposomal Dox [69], thus requiring further
15
development or improvement in liposomal drug formulation. Attempts were made to
improve the drug release rates of Dox by altering liposomal lipid compositions, but
this method led to uncontrolled and rapid drug release kinetics, which also lowered
therapeutic efficacy [70]. Therefore, a strategy to improve liposome-based anticancer
drugs should involve the development of a stable liposomal formulation with
improved drug release from the carrier in a controlled and sustained manner, thereby
enhancing bioavailability.
1.2.3 Co-delivery of chemotherapeutics to induce synergistic effect
Currently target-based drug design has successfully been used to develop numerous
drugs acting on novel molecular targets; however, they have shown poor efficacy in
clinical trials. This can be attributed to the compensatory mechanism, or drug-
mitigating response, used by the complex diseases such as cancer [71,72]. Overcoming
this compensation often requires high drug doses that can induce drug resistance at
the disease target site or side effects in other tissues [73], thus limiting the efficacy of
many potential drugs in cancer therapy. These limitations of mono-therapy can be
overcome by synergistic combination of two or more agents, which allows for reduced
dosage of each compound and can attack the disease system through multiple targets
[74,75]. However, current combination methods, through the cocktail administration,
have shown limited improvements in clinical studies due to the distinctive
pharmacokinetics of individual drugs which leads to non-uniform distribution after
systemic administration [76,77]. Moreover, an unexpected clinical outcome of
16
increasing toxicity was reported with these cocktail combinations, raising the concerns
that synergistic combination therapies would induce synergistic toxicity [78]. For
instance, although a doxorubicin (Dox) and paclitaxel (PTX) combinations have been
widely used in the treatment of tumors, particularly in metastatic breast cancer, the
clinical result was not as effective as expected and an increased cardiotoxicity was
observed [79,80,81,82]. Clinical pharmacokinetic studies also revealed an altered
plasma disposition of Dox and PTX when given in combination [83,84], rendering in
vitro data ineffective in predicting therapeutic efficacy of combination therapy in vivo.
Thus, a more effective combination strategy with the ability to unify the
pharmacokinetics and biodistribution of various drug molecules is highly desirable to
maximize the combinatorial effects without synergistic toxicity, yet challenging to
implement.
Development of nanotechnology has provided a novel combination strategy by
delivering multiple types of drugs simultaneously to the site of interest via a single
vehicle[77]. Nanoparticles are considered promising drug delivery vehicles for cancer
therapy based on their ability to prolong drug circulation time, reduce systemic
toxicity, and increase drug accumulation at tumor sites through the enhanced
permeation and retention (EPR) effect [85,86,87,88]. The pharmacokinetic behavior of
the co-formulated drugs can be determined by the pharmacokinetic behavior of the
drug carriers. Thus, nanoparticles offer the potential to coordinate the plasma
elimination and biodistribution of multiple drugs, enabling dosage optimization to
17
maximize cytotoxicity while minimizing the chances of cell resistance to any one drug.
Compared to other nanoparticles, liposomes have shown superior ability to co-deliver
multiple drugs that have vast differences in their hydrophobicity, to the sites of action
[89,90]. However, the poor stability and limited loading efficiency of hydrophobic
drugs remain significant concerns for most liposomal formulations, limiting their
utility in cancer therapy [91,92]. For example, a number of studies reported that
optimal drug-to-lipid molar ratio of paclitaxel-encapsulated conventional liposome
formulation was below 4% [93,94,95,96], thwarting the practical applications of
liposomes as drug carriers. Moreover, fine-controlling of the comparative loading yield
and release kinetics of multiple drugs via conventional liposomes remains an unmet
need. Thus, a stable liposomal formulation enabling improved drug loading and drug
release from the carrier in a controlled and sustained manner is necessary for
combinatorial drug delivery.
1.2.4 Co-delivery of chemotherapeutics to overcome multidrug resistance
The development of multidrug resistance (MDR) against a variety of conventional and
novel chemotherapeutic agents has been a major impediment to the success of cancer
therapy [97,98]. As one of the most important mechanisms involved in MDR, P-
glycoprotein (P-gp), an active drug efflux transporter, is overexpressed in the plasma
membrane of various cancer cells and capable of effluxing a broad range of anticancer
agents such as taxanes and anthracyclines [99,100]. For example, the efficacy of
doxorubicin (Dox) and paclitaxel (PTX), two agents widely used for the treatment of
18
various cancers, is often compromised by P-gp mediated MDR [101,102]. Therefore,
strategy of inhibition P-gp expression or bypass of P-gp-mediated drug efflux has been
developed to overcome MDR. For instance, a large number of P-gp inhibitors and
siRNAs targeting the gene encoding P-gp have been combinatorial delivered with
anticancer agents into cancer cells to down-regulate P-gp expression, enabling the
sufficient drug concentration of therapeutics for induction of cytotoxicity
[103,104,105]. However, clinical trials have been disappointing due to the high
toxicities of P-gp inhibitors (functional inhibitors or siRNAs) and the enhanced side
effects of chemotherapy in normal cells [106,107].
Combined therapy with multiple therapeutics provides a promising strategy to
suppress MDR, as different drugs may attack cancer cells at varying stages of their
growth cycles, thus decreasing the concentration threshold for individual drugs that is
required for cytotoxicity [108]. It has been reported that various drug combinations
successfully induce synergistic antitumor activity and prevent disease recurrence
[109,110]. For example, co-administration of Dox and PTX in cocktails, has been
considered as one of standard combined anthracycline-taxane regimens for various
tumors, as they are able to overcome drug resistance [111,112,113]. However, one
major challenge of combinatorial therapy is to unify the pharmacokinetics and cellular
uptake of combined therapeutics, which limited the clinical success of combination
therapy [83,114].
19
To overcome this challenge, strategy that allows loading multiple types of therapeutics
into a single drug-delivery vehicle and then concurrently delivering them the site of
action has been extensively explored [115,116]. Several drug-delivery systems have
shown the ability to co-deliver multiple drugs to tumors and improved antitumor
activity, potentially overcoming drug resistance by reducing the dosage of individual
drugs [117,118]. Indeed, nanoparticles are known to efficiently deliver therapeutics to
the tumor site through the enhanced permeability and retention (EPR) effect, thus
enhancing the concentration of therapeutics in tumors that is required for cytotoxicity
[85,86,88]. Moreover, nanoparticles can enter cancer cells by an endocytosis pathway,
which was independent from the P-gp pathway, thereby increasing the cell entry of
therapeutics to induce cytotoxicity [119,120,121]. Thus, a nanoparticular delivery
system that allows high efficiency of cellular entry of nanoparticles and subsequent
release of multiple anticancer drugs intracellularly to overcome MDR is highly
desirable.
20
Chapter 2
Site-specific Modification of Adeno-Associated Viruses via a Genetically
Engineered Aldehyde Tag
Yarong Liu, Yun Fang, Yu Zhou, Ebrahim Zandi, Chi-Lin Lee, Kye-Il Joo,
and Pin Wang
(Small, 2013, 9: 421-429)
21
2.1 Abstract
As a consequence of their well-defined nanostructure and intrinsic bioactive
functionality, virus-based nanoparticles have shown promise for mediating gene
delivery. Adeno-associated virus (AAV) nanoparticles, which possess an excellent
safety profile and therapeutic potential, hold potential for use in human gene therapy.
However, because of their native tropisms, the applicability of AAV nanoparticles is
often limited to the restricted ranges of cells or tissues. Thus, retargeting AAV particles
to the desired cell populations has continued to be a major research focus in many
gene therapy applications. In this study, we report a general strategy for nanoparticle
targeting. This involves the site-specific modification of AAV type 2 (AAV2) by
genetically incorporating a short peptide, in this case, an aldehyde tag, in the viral
capsid. Such a tag can be exploited for site-specific attachment of targeting molecules
and allows for further introduction of targeting antibodies or ligands. It was shown
that this aldehyde tag-based modification neither affected the level of infectious viral
titer nor intracellular trafficking properties. Furthermore, the site-specific conjugation
of targeting antibodies could significantly enhance viral transduction to those target
cells that have otherwise exhibited very low permissiveness to AAV2 infection. This
method also allowed for the functional incorporation of RGD peptides onto AAV2 for
enhanced delivery with implications for cancer gene therapy. Hence, by such
retargeting of AAV nanoparticles, our data reveal their expanded potential to
transduce various cell types and achieve targeted gene delivery.
22
2.2 Introduction
The development of nanoscale delivery vectors that allow for delivering genes to target
cells with prolonged circulation time has progressed substantially in recent years.
Although nonviral nanoparticle vectors such as polymers, liposomes, or dendrimers,
are promising in terms of safety concern in gene therapy, their relatively low gene-
transfer efficiency raises greater challenges [122,123]. Virus-based nanovectors,
considered as genetically encoded nanoscale structures, have been broadly used in gene
therapy clinical trials based on their unique properties and functionalities [124]. Many
efforts have been made to engineer viral nanoparticles including cowpea mosaic virus
[125], hepatitis B virus [126], and bacterial virus [127] for targeted gene delivery by
utilizing their unique sets of characteristics such as structures and functions. Among
various viral nanoparticles, adeno-associated virus (AAV) has been considered as a
promising vehicle for human gene therapy based on its ability to infect both dividing
and nondividing cells, as well as establish long-term gene expression in vivo without
known pathological consequence of infection [6,7,8,128,129,130]. The AAV type 2
(AAV2) nanoparticles constitutes the first primate AAV to be cloned, and promising
results have been obtained with this nanovector in clinical gene transfer, including
cystic fibrosis [131], retinal degenerative disorders [14,132,133] and haemophilia B
[19,134].
Although AAV2 nanovectors possess a high safety profile and remarkable potential in
several disease models, their utilization in many areas of gene therapy is limited by the
restricted tissue tropism of AAV2. Furthermore, the use of native viral tropism can
23
also cause non-specific infection of many cells, which raises concerns over feasibility
and safety for systemic administration of the nanovector. Therefore, retargeting the
viral particle tropism of AAV2 to desired target cells could not only expand its utility
for gene delivery applications, but also mitigate safety issues for use in human gene
therapy. Significant effort has been devoted to either an directed evolution approach
that can create novel AAV variants with desired gene delivery properties [135,136,137]
or the alteration of capsid proteins, which are responsible for virus binding to cell
surface receptors and mediating cellular entry [37,38,138].
Currently, most targeting efforts rely upon genetic modification of AAV capsid
proteins by insertion of targeting peptide motifs that can direct nanovectors to specific
cell types. This method has been successfully employed to retarget AAV to arterial
endothelium,[40] striated muscles [41], and brain vasculature [139]. However, the
major technical challenges in this manipulation include the low production yield,
dramatic reduction of vector titer, or significant drop of DNA packaging efficiency
[40,43]. In order to sidestep the issues of viral particle assembly and size limit for
insertion of targeting peptides, chemical modification methods have been explored.
Chemical manipulation utilizes the reactivity of functional groups, such as lysine and
cysteine residues, to introduce targeting ligands on viral particles. However, random
modification of lysine residues can lead to inactivation of the protein [44]; therefore,
the lysine modification of viral capsids may potentially alter virus-host cell interaction
and/or cause a loss of viral infectivity. Therefore, site-specific modification is highly
desirable for both labelling and targeting of viral nanoparticles.
24
In this study, we report a general method for site-specific modification of AAV2 by
introducing a small aldehyde tag onto the virus capsid, which can be exploited for the
selective modification on the surface of viral nanoparticles. It was found that this
aldehyde tag-based nanoparticle modification had little effect on the virus production
and infectivity. In addition, it was demonstrated that this aldehyde tag enabled
covalent attachment of hydrazide-functionalized molecules including antibodies and
peptides to AAV2 nanoparticles in a site-specific manner. It was shown that antibody
conjugation to AAV2 could significantly enhance viral transduction in both permissive
and nonpermissive cell lines. Furthermore, RGD peptide-conjugated AAV2
nanoparticles exhibited tumor targeting in vitro. These results demonstrated a general
and efficient means for retargeting AAV nanovectors and could also expand the utility
of AAV nanovectors for human gene delivery.
2.3 Results and Discussion
2.3.1 Confirming and Visualizing Aldehyde Tag on Modified AAV Nanoparticles
As shown in Figure 5A, our strategy involves genetic insertion of a 13-amino acid
consensus sequence (LCTPSRAALLTGR) into the cap gene at the amino acid position
587 located within the VP1/VP2/VP3 regions, which have been reported to be capable
of tolerating insertion of functional amino acid sequences and generating modified
AAV nanoparticles with titers comparable to unmodified particles [140,141]. The
inserted sequence could be recognized by cellular formylglycine generating enzyme
(FGE), which converts cysteine (Cys) to aldehyde-bearing formylglycine (FGly)
25
residue [142,143,144,145]. Consequently, the AAV2 capsid proteins were outfitted
with an aldehyde group that allows for site-specific and chemoselective modification of
AAV2 nanoparticles with hydrazide- or aminooxy-functionalized targeting/labeling
elements, such as peptides, antibodies, and fluorophores, without significant loss of
viral function and infectivity.
Figure 5. AAV2 nanoparticles can be site-specifically modified by genetically encoded
aldehyde tags. (A) General strategy for the site-specific modification of AAV2 nanoparticles by
genetically encoded aldehyde tag. A 13-amino-acid sequence can be inserted to AAV2 capsids.
Upon expression, the encoded cysteine is modified to an aldehyde tag, which can be covalently
conjugated with hydrazide- or hydroxylamine-functionalized molecules. (B) Fluorophore
labeling of aldehyde tag. Wild-type (AAV2
WT
) and aldehyde tag-bearing (AAV2
Ald13
)
nanoparticles were incubated with the Alexa488-hydrazide (green) and subsequently probed
with a mouse monoclonal antibody (A20) specific for intact AAV2 (red). The labeled aldehyde
tags (green) were co-localized with AAV2
Ald13
nanoparticles (red). Overlapping green and red
signals appear as yellow in a merged image. (C) The percentages of colocalization between
AAV2
Ald13
(anti-AAV2) and aldehyde tags (Alexa488-hydrazaide) measured by viewing more
than 80 randomly chosen field of views. The percentages of colocalization were calculated
using the Mander’s overlap coefficient.
26
To determine whether the aldehyde tag could be incorporated onto viral particles, the
wild-type AAV2 (AAV2
WT
) and aldehyde-tagged AAV2 (AAV2
Ald13
) nanoparticles
were produced and reacted with Alexa488-hydrazide at pH 6.5 for 2 hr. Subsequently,
the particle-containing solutions were overlaid onto coverslips and immunostained
with an antibody specific for intact AAV2 nanoparticles. Most of the hydrazide signals
(> 60%) were colocalized with the AAV2 signals (Figure 4B, bottom; Figure 4C),
whereas no significant hydrazide signals were observed for viral particles lacking the
aldehyde tag (Figure 5B, upper), indicating that aldehyde groups are efficiently
expressed on the surface of AAV2
Ald13
.
We further confirmed the incorporation of aldehyde tags on viral particles by a
transmission electron microscope (TEM) assay. The gold (Au) nanoparticles with
amine functional groups (NH 2-Au) were first reacted with a crosslinker, SANH
(succinimidyl 4-hydrazinonicotinate acetone hydrazone), to create a free hydrazide,
allowing for further conjugation to AAV2
Ald13
via the aldehyde tag (Figure 6A). The Au
nanoparticles on viral surfaces were readily detected, and the TEM images showed that
the size of individual Au-conjugated AAV2
Ald13
particles was approximately 30 nm in
diameter (Figure 6B, bottom), which is slightly bigger than the unconjugated
AAV2
Ald13
(Figure 5B, upper). Furthermore, the hydrodynamic radii of AAV2
Ald13
and
Au-AAV2
Ald13
nanoparticles in aqueous solutions were measured by dynamic light
scattering (DLS), and the results indicated an increase in the mean radius of Au-
AAV2
Ald13
(71.1nm) as compared to that of AAV2
Ald13
(35.9 nm), suggesting that Au
particles were indeed conjugated to AAV2
Ald13
nanoparticles (Figure 7). The sizes of
27
particles detected by DLS are generally larger than that measured by TEM, presumably
by the influence of the counterion cloud on particle mobility and the intrinsic
aggregation tendency of AAV particles in aqueous solutions [146,147,148].
Figure 6. Covalent attachment of amine-functionalized gold (Au) particles onto AAV2
Ald13
nanoparticles and characterization of Au-labeled AAV2
Ald13
nanoconjugates. (A) Schematic
representation of the method used to conjugate AAV2
Ald13
with Au particles via a crosslinker,
SANH. (B) TEM images of unconjugated AAV2
Ald13
nanoparticles (upper) and Au-labeled
AAV2
Ald13
nanoparticles (bottom). Scale bars represent 20 nm. (C) Histogram of number of Au
particles per AAV2
Ald13
nanoparticle obtained from TEM images.
28
0
2
4
6
8
10
12
14
16
18
0.01 0.10 1.00 10.00 100.00 1.0E+3 1.0E+4 1.0E+5
0.01 0.10 1.00 10.00 100.00 1.0E+3 1.0E+4 1.0E+5
0
2
4
6
8
10
12
14
16
18
20
AAV2
Ald13
Au-AAV2
Ald13
R (nm)
R (nm)
% Intensity % Intensity
Figure 7. The hydrodynamic size of unconjugated AAV2
Ald13
and Au-AAV2
Ald13
measured by
dynamic light scattering.
Additionally, analysis of the TEM images suggested that approximately 6~9 Au
nanoparticles were detected in the majority of individual AAV2
Ald13
nanoparticles
(Figure 6C), implying that approximately 6~9 aldehyde tags per AAV2
Ald13
nanoparticle are available for conjugation with hydrazide/hydroxylamine. The number
of aldehyde tags functionally displayed on individual AAV2
Ald13
nanoparticles was
further determined by measuring the spectroscopic property of the purified AAV2
Ald13
conjugated with Alexa488-hydrazide dye at different initial ratios as described in
29
Materials and Methods. The spectroscopic measurement indicated an average dye-to-
nanoparticle ratio (dye/AAV2
Ald13
molar ratio) of approximately 7 (Figure 8). Based on
the fact that 1) a 13-amino acid consensus sequence was inserted into the cap gene
within VP1/VP2/VP3 regions and 2) the AAV2 capsid is composed of 60 capsid
protein subunits, one can calculate that a single AAV2 nanoparticle should contain 60
Cys-bearing 13-mer peptides that are prone to conversion by the cellular FGE enzyme
to FGly-bearing peptides. Hence, the ratio of 7 aldehyde tags to 60 Cys-containing 13-
mer peptides reflects an approximate 12% conversion of Cys to FGly, although this
estimation is based on the assumption that the conjugation is quantitative and that all
displayed FGly side-chains are accessible for the conjugation chemistry.
Figure 8. Quantitative analysis of the number of attached dyes per AAV2
Ald13
particle. The
numbers were determined by absorption measurement on solutions of purified Alexa488-
hydrazide-conjugated AAV2
Ald13
by comparison of the intensities of dyes to that of viral
particles.
The expression of an aldehyde tag on AAV2 capsid proteins VP1/VP2/VP3 was
further confirmed by Western blot against hydrazide-functionalized biotin, whereas no
30
signal was detected in AAV2
WT
(Figure 9B). As a control, the VP1, VP2, and VP3
proteins, which were expressed in the appropriate stoichiometric ratio of 1:1:10, for
both unmodified (AAV2
WT
) and modified (AAV2
Ald13
) particles were readily detected
by an anti-AAV B1 antibody (Figure 9A). Furthermore, tryptic digestion of the
aldehyde-tagged capsid proteins allowed for direct identification of FGly and Cys by
mass spectrometry (Figure 9C). The presence of Cys was further confirmed by
identification of carbamidomethyl Cys after treatment with 2-iodoacetamide. These
peptide identities were also corroborated by the LC/MS/MS analysis (Figure 10),
further validating the incorporation of the aldehyde tag.
2.3.2 Evaluate the Effect of Modification on AAV Function
It was next determined how modified AAV2 nanoparticles might affect their biological
function. To examine whether the modified AAV2 remained infectious, either
unmodified or modified AAV2 carrying a GFP reporter gene was used to infect 293T
cells. A similar level of production (genome copies/ml) and functional titer
(transducing units/ml) was found in AAV2
Ald13
and AAV2
WT
(Figure 11A and 11B),
indicating that the introduction of the aldehyde tag has minimal perturbation on
production and transduction capability of AAV2 nanoparticles.
It has been reported that the intracellular transport properties of viral nanoparticles
can be altered by the modification of AAV capsid [149,150]. Thus, it is important to
determine whether the incorporation of aldehyde tag and its site-specific modification
of AAV2 capsid protein alter the intracellular trafficking of AAV2 in cells.
31
Figure 9. The expression and chemical modification of aldehyde tag on AAV2 capsid proteins.
(A) Detection of AAV capsid proteins (VP1, VP2, VP3) by Western blot analysis using a B1
antibody. (B) Site-specific labeling and detection of aldehyde tag with the biotin probe by
Western blot analysis. AAV2
WT
and AAV2
Ald13
were incubated with biotin hydrazide and
probed with streptavidin-conjugated antibody coupled to HRP. (C) Mass spectra confirming
the presence of formylglycine (FGly) and Cys in a tryptic peptide from AAV2
Ald13
nanoparticles by LC-MS [(M+2)/2]. (Left) Mass spectrum of the tryptic fragment
incorporating FGly (theoretical: 473.2362 m/z; observed: 473.24 m/z). (Middle) Mass spectrum
of the tryptic fragment incorporating unmodified Cys (theoretical: 482.2362 m/z; observed:
482.24 m/z). (Right) Mass spectrum of tryptic fragment incorporating unmodified Cys after
treatment with 2-iodoacetamide (theoretical: 510.7470 m/z, observed: 510.75 m/z).
32
Figure 10. (A) LC/MS/MS spectrum of the peptide fragment QTGLFGlyTPSR from AAV2
Ald13
.
(B) LC/MS/MS spectrum of the peptide fragment QTGLCTPSR from AAV2
Ald13
. (C)
LC/MS/MS spectrum of the peptide fragment QTGLCTPSR from AAV2
Ald13
after reacting with
2-iodoacetamide.
VPSIM_ITIT #3987 RT: 50.77 AV: 1 NL: 8.19E1
T: ITMS + p NSI Full ms2 473.24@cid35.00 [130.00-2000.00]
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
m/z
0
5
10
15
20
25
30
35
40
45
50
55
60
65
70
75
80
85
90
95
100
Relative Abundance
464.33
455.25
437.33
468.08
427.17
261.42 327.25
648.42
747.42
537.83 366.08 550.92
800.83 393.08 527.08 718.67 760.75
301.08 212.92 875.50 660.75 154.92
579.42 270.42 602.25 668.67 799.33
199.17
163.58 813.33 471.08 831.50
522.25
y9
2+
-NH
3
(464.33)
b9
2+
-NH
3
(455.25)
y5
+
-H
2
O (527.08)
y2
+
(261.42)
b2
+
-H
2
O
(212.92)
y8
+
-NH
3
(799.33)
b5
+
-NH
3
(468.08)
Q T GL FGly TP SR
y9 y5
b2 b9
y2
b5
y8
A
m / z
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
30
10
0
20
100
80
90
70
50
60
40
Relative Abundance
VPSIM_ITIT #3982 RT: 50.70 AV: 1 NL: 8.18E1
T: ITMS + p NSI Full ms2 482.24@cid35.00 [130.00-2000.00]
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
m/z
0
5
10
15
20
25
30
35
40
45
50
55
60
65
70
75
80
85
90
95
100
Relative Abundance
464.33
472.83
459.92
388.92
446.33
393.08
475.58 243.42
634.92 411.00
382.25
620.58 558.75
337.42 234.33
726.50
302.08 656.75 707.67
806.58
276.00 601.83 217.25
745.92 488.50
211.08 814.67
801.67 499.58
549.08
162.75 834.92
797.83
526.33
b9
2+
-NH
3
(464.33)
y9
2+
-NH
3
(472.83)
y2
+
-H
2
O
(243.42)
b4
+
-H
2
O
(382.25)
b2
+
-H
2
O
(211.08)
y4
+
(459.92)
Q T GL C TP S R
y9
y4
b2 b9
b4
y2
B
30
10
0
20
100
80
90
70
50
60
40
Relative Abundance
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
m / z
VPSIM_ITIT #3940 RT: 50.17 AV: 1 NL: 1.02E2
T: ITMS + p NSI Full ms2 510.75@cid35.00 [140.00-2000.00]
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
m/z
0
5
10
15
20
25
30
35
40
45
50
55
60
65
70
75
80
85
90
95
100
Relative Abundance
492.25
501.75
410.17 480.00
819.42
478.67
820.75
418.50 312.08
474.33
402.33
457.50 266.75 746.25 659.67
602.42 401.50 327.25
805.58 544.92 201.00
258.67 678.42 709.25 392.25 874.00 631.42 586.00 516.58
791.42 906.17
561.17 292.25
173.08 930.83 227.42
872.83 998.50 963.75
849.42 153.42
b9
2+
-H
2
O (492.25)
y9
2+
-NH
3
(501.75)
y5
+
-H
2
O (602.42)
b6
+
(659.67)
y8
+
-H
2
O (874.00)
b4
+
(401.50)
Q TGL CT PSR
y8 y5
b4 b6
y9
b9
C
150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000
m / z
30
10
0
20
100
80
90
70
50
60
40
Relative Abundance
33
Figure 11. Comparision of nanoparticle titers and functional titers of AAV2
WT
and AAV2
Ald13
.
(A) Nanoparticle titers (genome copies) were determined by quantitative PCR assay. (B)
Functional titers were measured by gene transduction assay in 293T cells. 293T cells (0.1×10
6
)
were spin-infected with unmodified AAV2 (AAV2
WT
) or modified AAV2 (AAV2
Ald13
)
nanoparticles containing a GFP transgene at the same multiplicity of infection (MOI) (genome
copies/cell). The cells were then analyzed by flow cytometry for GFP expression at 72 hr post-
infection. All data are shown as the means of triplicate experiments from three different
nanoparticle preparations.
34
Merged EEA1 AAV2
Ald13
Merged CI-MPR AAV2
Ald13
Merged Rab11 AAV2
Ald13
30 min 45 min 45 min
15 min 15 min
AAV2
Ald13
+ Clathrin AAV2
Ald13
+ Caveolin-1 A B
C D E
Figure 12. Intracellular trafficking of AAV2
Ald13
in HeLa cells. (A-E) Alexa488-labeled
AAV2
Ald13
vectors (green) were added to HeLa cells for 30 min at 4 °C to synchronize infection
and were then shifted to 37 °C for different periods (indicated in A-E). The cells were fixed,
permeabilized, and immunostained with antibodies against clathrin (A), caveolin-1 (B), EEA1
(C), CI-MPR (D) or Rab11 (E) and counterstained with TO-PRO-3 (blue). The boxed regions
are enlarged in the right panels (A and B), or the bottom panels (C to E). The arrows indicate
the viral particles colocalized with endocytic markers. Scale bar represents 10 µm.
35
Figure 13. Targeting of anti-HLA-conjugated AAV2
Ald13
nanoparticles to receptor-bearing cells.
(A) Schematic representation of the method used to conjugate AAV2
Ald13
nanoparticles with an
anti-HLA antibody. (B) Transduction of 293T and HepG2 cells by AAV2
WT
, AAV2
Ald13
, and
AAV2
Ald13
conjugated with an isotype control antibody (AAV2
Ald13
-ctrlAb), or AAV2
Ald13
conjugated with an anti-HLA antibody (AAV2
Ald13
-αHLA) to express GFP. The percentage of
GFP
+
cells was analyzed by flow cytometry. Error bars represent the standard deviation of the
mean from experiments conducted in triplicate (*p < 0.05). (C) GFP fluorescence microscopy
images of 293T cells infected by indicated particles. Upper: Bright field image. Lower: GFP
Fluorescence. Scale bar represents 50 µm.
36
Figure 14. Targeted gene transduction mediated by antibody-conjugated AAV2
Ald13
vectors.
(A) Schematic of anti-CD20 antibodies conjugated to the AAV2
Ald13
vector. Upper: Addition of
a hydrazide group to antibodies using SANH (succinimidyl 4-hydrazinonicotinate acetone
hydrazone). Lower: Covalent conjugation of hydrazide-functionalized antibodies to AAV2
Ald13
vector. (B) Transduction of 293T.CD20 cells (~1000 viral genome copies per cell) mediated by
AAV2
WT
, AAV2
Ald13
, isotype control antibody-conjugated AAV2
Ald13
or anti-CD20-conjugated
AAV2
Ald13
at the same MOI (genome copies/cell). The percentage of GFP-positive cells was
analyzed by FACS at 72 h post-infection. Error bars represent the standard deviation of the
mean from triplicate experiments.
To this end, we utilized Alexa488-hydrazide to label AAV2
Ald13
and monitored its
various entry routes. It has been generally hypothesized that AAV2 internalizes to the
cells through clathrin-mediated endocytosis [148,151,152,153,154]. To confirm the
role of clathrin- or caveolin-mediated endocytosis in the entry of modified AAV2, we
visualized the dye-labeled viral nanoparticles and endocytic structures (clathrin or
caveolin) in HeLa cells after 15 min incubation at 37 °C. As shown in Figure 12A and
37
12B, a significant colocalization of modified AAV2 nanoparticles with clathrin
structures was detected, while no significant colocalization of particles with caveolin
was observed. By the colocalization experiment, we also demonstrated that the
modified AAV2 nanoparticles migrate from the early endosomes to both the late and
recycling endosomes. As shown in Figure 12C to 12E, after 30 min incubation, most
nanoparticles were colocalized with EEA1 [155,156,157,158], an early endosome
marker, while after 45 min, most particles were observed in the late endosome (CI-
MPR) [155,158,159] and recycling endosome (Rab11) [160,161]. These trafficking
properties are in good agreement with the wild-type AAV2 [34,162]. Taken together,
the results suggested that this method of modification neither affected virus titer nor
intracellular trafficking routes of the viral nanoparticles.
2.3.3 Enhanced Transduction of AAV2
Ald13
by Conjugating with Targeting Ligands
To evaluate the targeting ability of AAV2Ald13 upon site-specific introduction of
targeting motifs, a mouse monoclonal antibody against human leukocyte antigen
(HLA) was conjugated with viral nanoparticles via the crosslinker SANH (Figure 13A).
Because significant proportions of human cell types express HLA, including HepG2
cells that have exhibited very limited permissiveness to AAV2 infection,
[22]
such
modification might have a tendency to expand the natural tropism of AAV2
nanoparticles. Indeed, flow cytometric analysis showed that transduction in 293T cells
mediated by anti-HLA-conjugated AAV2
Ald13
was significantly increased from 15.7% to
36.7% (p < 0.05) as compared to that mediated by unmodified AAV2 (Figure 13B, left).
Moreover, the enhanced transduction was specific for HLA because AAV2
Ald13
38
conjugated with an isotype control antibody showed no marked effect on transduction
efficacy. The increased transduction of AAV2
Ald13
conjugated with anti-HLA was also
observed in HepG2 cells, as shown in Figure 13B (right, p < 0.01). The enhanced
transduction of anti-HLA-conjugated-AAV2
Ald13
in 293T cells was further confirmed
by the GFP expression detected by fluorescence microscopy (Figure 13C). Finally,
transduction of AAV2
Ald13
was also remarkably enhanced in CD20-expressing 293T
(293T.CD20) cells by site-specific conjugation of anti-CD20 through the crosslinker
SANH (Figure 14). In addition, the number of antibody molecules conjugated to each
AAV2
Ald13
nanoparticle was determined by measuring the spectroscopic property of
the purified antibody-conjugated AAV2
Ald13
. The measurement indicated that the
average number of antibodies conjugated to each viral nanoparticle is approximately
5.4. Thus, this modification method can be employed to specifically target certain
tissues or cell types that are otherwise poorly transduced, thereby enhancing the
applicability of AAV2 nanoparticles for gene delivery.
Gene delivery to tumors by active targeting methods has been considered a potential
strategy for cancer gene therapy, although nanoscale delivery vectors already have
intrinsic capability for passive accumulation at the tumor site by the enhanced
permeability and retention (EPR) effect. Genetic incorporation of tumor-specific
peptides or functional motifs to viral nanovectors has been explored for tumor
targeting. For example, Arginine-Glycine-Aspartate (RGD) motif has been reported to
mediate enhanced gene transfer of modified AAV2 to tumor cells by using biotin
ligase.
[163]
In addition, genetically modified RGD-AAV2 has been demonstrated to
39
efficiently infect HSPG receptor-negative cell lines that lack the expression of heparin
sulfate proteoglycan (HSPG), via RGD-integrin interaction [164]. Here, we chemically
modified the cyclic RGD peptide (c(RGDfC)) with EMCH (3,3’-N-[e-
maleimidocaproic acid] hydrazide) to introduce a free hydrazide group, which was
further exploited for covalent linkage to AAV2
Ald13
nanoparticles via the aldehyde tag
(Figure 15A). The average number of cyclic RGD peptides conjugated to each
nanoparticle was approximately 7.4. As shown in Figure 15B, the c(RGDfC)-
conjugated AAV2
Ald13
displayed markedly enhanced transduction (11%) in HeLa cells
as compared to that of unconjugated AAV2
Ald13
(3.7%, p< 0.05).
Figure 15. Targeting of peptide-conjugated AAV2Ald13 nanoparticles to tumor cells. (A)
Schematic representation of the method used to conjugate AAV2Ald13 with a cyclic RGD
peptide. Upper: addition of a hydrazide group to the cyclic RGD peptide (c(RGDfC)) using
EMCH (3,3ʹ -N-[e-Maleimidocaproic acid] hydrazide, trifluoroacetic acid salt). Lower:
covalent conjugation of hydrazide-RGD peptides to AAV2Ald13 nanoparticles. (B)
Transduction of HeLa cells by AAV2Ald13 or c(RGDfC)-conjugated AAV2Ald13
(AAV2Ald13-(CfDGR)c) nanoparticles. The data are presented as the mean values ± SD (n
>3).
40
4. Experimental Section
Cell lines, antibodies, and reagents: 293, 293T, HepG2, 293T.CD20, and HeLa cells
were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Gibco) supplemented
with 10% FBS (Sigma-Aldrich, St. Louis, MO) and 2 mM L-glutamine (Hyclone
Laboratories, Inc., Omaha, NE). Mouse monoclonal antibodies against the intact
AAV2 (A20) and capsid proteins (B1) were purchased from American Research
Products, Inc. (Belmont, MA). Mouse monoclonal antibodies against clathrin and
caveolin-1 and the rabbit polyclonal antibody specific to CI-MPR were obtained from
Abcam (Cambridge, MA). The mouse monoclonal anti-EEA1 antibody, anti-CD20
(2H7), isotype control antibody and the rabbit polyclonal anti-Rab11 antibody were
purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). The mouse
monoclonal antibody to HLA class I antigen was obtained from Sigma-Aldrich. Texas
Red- or Alexa488-conjugated goat anti-mouse IgG antibody, Alexa647-conjugated
goat anti-rabbit IgG antibody and Alexa488-hydrazide were obtained from Invitrogen
(Carlsbad, CA). Cyclic-RGD peptide (c(RGDfC)) was obtained from Peptides
International (Louisville, KY). EMCH (3,3ʹ-N-[e-Maleimidocaproic acid] hydrazide
trifluoroacetic acid salt) and SANH (succinimidyl 4-hydrazinonicotinate acetone
hydrazone) were purchased from Pierce Biotechnology (Rockford, IL).
41
Plasmids: For construction of plasmid pAAV2-Ald13, assembly PCR was employed to
insert the DNA sequence (TGLCTPSRAALLTGRGLS; aldehyde tag region is
underlined, and linkers (TG and GLS) are included to promote efficient display and
flexibility of the motif) into the cap gene at amino acid position 587. Briefly, DNA
primers were designed to encode the sequence of aldehyde tag, and PCR amplifications
were performed using the forward primer (5’-TAT-CGT-ACG-TCC-TCG-GCT-
CGG-CGC-ATC-3’) and the reverse primer (5’-GCA-GGG-CGG-CCC-GGC-TGG-
GGG-TGC-ACA-GAC-CAG-TTT-GTC-TGT-TGC-CTC-TCT-GGA-GGT-3’) or the
forward primer (5’-CAG-CCG-GGC-CGC-CCT-GCT-GAC-CGG-CCG-GCT-GAG-
CGC-AGC-TAC-CGC-AGA-TG-3’) and the reverse primer (5’-TTA-TCT-AGA-
GCA-TGG-CTA-CGT-AGA-TAA-GTA-GCA-TGG-C-3’). The two fragments were
fused using assembly PCR, and the PCR product was cloned into the plasmid pAAV2-
wt via restriction sites BsiW1 and Xba1.
Recombinant AAV2 production: Recombinant AAV2 nanovectors (AAV2
WT
and
AAV2
Ald13
) were produced in HEK-293 cells as previously described. Forty 150-cm
dishes of subconfluent 293 cells were triple-transfected with 650 µg each of AAV2 cis-
plasmid carrying a GFP reporter gene under the control of the CMV promoter (CB7)
and AAV2 trans-plasmid containing the AAV2 rep and cap genes (pAAV2-wt) or
encoding the aldehyde tag sequence (pAAV2-Ald13) and 1300 µg of the adenovirus
helper plasmid pΔF6, using the calcium phosphate precipitation method. After
overnight incubation, the medium was replaced with fresh D10 media, and the cells
42
were harvested at 72 h post-transfection, followed by three cycles of freeze and thaw.
The cell lysates were then mixed with a stock solution of 40% PEG 8000 (final
concentration: 8%, Sigma, St Louis, MO, USA) and 2.5 M NaCl, followed by
incubation on ice for 2h.
[165]
The mixture was then centrifuged at 2500 g for 30 min,
and the supernatant was discarded. The pellet containing viruses was then resuspended
in buffer and purified by cesium chloride gradient density centrifugation at 25,000 rpm
and 15°C for 20 h (Optima L-90 K Ultracentrifuge, SW-28 rotor, Beckman Coulter,
Brea, CA). Viral nanoparticles recovered from the first round ultracentrifugation were
pooled and subjected to isopycnic separation by a second CsCl centrifugation at 13,000
rpm and 15°C for 20 h in a SW-32 Ti Roter. Fractions containing AAV2 determined
by refractive index were further desalted in PBS using an Amicon Ultra 100 kDa
MWCO centrifugal filter device (Millipore, Billerica, MA).
Probe ligation and antibody conjugation: For fluorescent labeling of AAV2
Ald13
, the
purified viral nanoparticles were incubated with Alexa488-hydrazide (500 µM) at pH
6.0-7.0 at RT for 2 hr. Excess unreacted dyes were removed by a gel filtration column.
For the conjugation of RGD peptide onto AAV2 nanoparticles, cyclic-RGD peptide
(c(RGDfC)) was reacted with EMCH at RT for 2 hr to introduce a free hydrazide
group, followed by a gel filtration column to remove unbound EMCH. Subsequently,
the hydrazide-functionalized cyclic RGD peptide was incubated with AAV2
Ald13
at pH
7.0 for 8 hr at RT. For antibody conjugation to viral nanoparticles, anti-CD20, anti-
HLA, and control antibody were modified with SANH to create a free hydrazide to
43
antibodies. After removing excess SANH by the gel filtration column, the modified
antibodies were conjugated to AAV2
Ald13
for 8 hr at RT.
For the spectroscopic measurement to determine the number of dyes bound to
AAV2
Ald13
particle, AAV2
Ald13
nanoparticles were labeled with various concentrations
of Alexa488-hydrazide dye in order to saturate any remaining aldehyde tags on the
viral particle. After purification by a gel filtration column, the number of attached dyes
per AAV2
Ald13
nanoparticle was then calculated by measuring the absorbance of
purified Alexa488-hydrazide-labeled AAV2
Ald13
particles at 494 nm for Alexa488-
hydrazide (ε=71,000 M
-1
cm
-1
) and at 280 nm for AAV particles (ε=6.61 x 10
6
M
-1
cm
-
1
).
[166]
The quantification of the number of peptides and antibodies bound per AAV2
Ald13
nanoparticle was also determined by using the absorption measurements. However, to
avoid misleading absorbance measurement of AAV particles at 280 nm after
peptides/antibodies attached on viral particles, fluorescent dye-labeled AAV2
Ald13
,
antibodies, and peptides were used as a tracer in the coupling reaction. First, AAV2
Ald13
nanoparticles were labeled with Cy5-NHS ester (GE Healthcare, Piscataway, New
Jersey) at 2.5 molar ratio of Cy5 to AAV2
Ald13
. Similarly, FITC-labeled antibodies (1.2
molar ratio of FITC to IgG) and FITC-labeled peptides (1 molar ratio of FITC to
peptide) were prepared for coupling reaction. For conjugation reaction, Cy5-labeled
AAV2
Ald13
nanoparticles were incubated with a saturating concentration of FICT-
labeled antibodies or peptides. The extent of conjugation of antibodies or peptides on
AAV2
Ald13
nanoparticles was then calculated by measuring the absorbance of the
44
purified coupling products at 650 nm for Cy5 (ε =250,000 M
-1
cm
-1
) and at 494 nm for
FITC (ε =68,000 M
-1
cm
-1
).
Viral nanoparticle transduction: Target cells (0.2×10
6
) were seeded onto a 24-well
culture dish and spin-infected with nanoparticle preparations as described above
(~1000 viral genome copies per cell) at 2,500 rpm for 90 min at 25 °C, using a Sorval
Legend centrifuge. After an additional 3 hr of incubation, the medium was removed
and replaced with fresh culture medium. Gene transduction was determined after 3
days of infection by flow cytometry analysis of GFP
+
cells and inverted fluorescent
microscope. All transduction assays were performed in triplicate, and the results are
presented as mean values ± SD. Analysis was performed using Student’s t-test for
comparison of two groups. Results were considered significant when P <0.05.
Quantification of genome copies by qPCR: Viral DNA was extracted via the QIAamp
MinElute virus spin kit (Qiagen, Valencia, CA) according to the manufacturer’s
protocol. The genome copies of vectors were quantified by the Bio-Rad MyiQ RT-PCR
(Bio-Rad, Hercules, CA) system by using a pair of primers specific for the GFP
transgene: 5’-GACATCAT-GAAGCCCCTTGAG-3’ (forward) and 5’-
GGTGGTCGAAATTCA-GATCAAC-3’ (reverse). AAV2 cis-plasmid (CB7) was
performed as a reference.
45
Confocal imaging: Fluorescence images were acquired 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 Plan-apochromat 60×/1.49 oil
immersion objective and a Cascade II: 512 EMCCD camera (Photometrics, Tucson,
AZ). Illumination powers at 491, 561, and 640 nm solid-state laser lines were provided
by an AOTF (acousto-optical tunable filter)-controlled laser-merge system with 50mW
for each laser.
To confirm the expression of aldehyde tag on the viral particles, Alexa488-hydrazide-
conjugated nanoparticles (AAV2
WT
or AAV2
Ald13
) were overlaid on polylysine-coated
glass-bottom dishes (MatTek Corporation, Ashland, MA) for 1 hr at 37 °C. The dish
was then washed three times with PBS and then immunostained with an antibody
(A20) against intact AAV2 nanoparticles. To quantify the extent of colocalization, the
percentages of colocalization were calculated using Mander's overlap coefficient by
viewing more than 80 randomly chosen field of views. The Mander's overlap
coefficient was generated by using the Nikon NIS- Elements software.
For the colocalization study with endocytic markers, HeLa cells were seeded on glass-
bottom dishes overnight. Alexa488-hydrazide-conjugated AAV2
Ald13
nanoparticles
were added to the cells (~1000 viral genome copies per cell) and incubated for 30 min
at 4 °C for binding. After washing with PBS, the cells were incubated at 37 °C to
initiate nanoparticle internalization for the indicated time periods. The cells were then
fixed, permeabilized by 0.1% Triton X-100, immunostained with corresponding
antibodies specific to clathrin, caveolin, EEA1, CI-MPR, or Rab11, and counterstained
46
with TO-PRO-3 (Invitrogen). All images were analyzed using Nikon NIS-Elements
software.
Western blot: Western blot was performed via a standard protocol. Briefly, both
AAV2
WT
and AAV2
Ald13
nanoparticles were reacted with biotin-hydrazide at a final
concentration of 500 µM for 2 hr at pH 6.0-7.0. The biotinylated viral particles were
denatured by boiling and separated in reducing 12% polyacrylamide gel. The separated
proteins were then transferred to polyvinylidene difluoride membranes. For detection
of biotinylated capsid proteins, the blot was probed with a streptavidin-conjugated
HRP. For immunological detection of capsid proteins, AAV2
WT
and AAV2
Ald13
nanoparticles were denatured and separated using the same protocol described above.
Immunodetection of capsid proteins (VP1, VP2 and VP3) was carried out with
antibodies specific to AAV2 capsid proteins (B1) and anti-mouse IgG conjugated to
HRP. Detection of the bands was done using ECL chemiluminescent substrate.
Tryptic digestion of AAV2
Ald13
nanoparticles: Capsid proteins of the modified
nanovector were reduced by adding dithiothreitol to a final concentration of 10 mM at
95°C for 10 min. The proteins were digested with trypsin (Promega, Madison, WI) at
the ratio of 1:20 (mass ratio, enzyme/substrate) at 37°C overnight. The digests were
then diluted and injected into LC-ESI-MS (Waters Micromass Q-TQF). For
LC/MS/MS analysis, samples were analyzed using an Eksigent NanoLC Ultra 2D
(Dublin, CA) and Thermo Fisher Scientific LTQ Orbitrap XL (San Jose, CA). Briefly,
47
peptides were separated in a 10 cm column (75 µm inner diameter) packed in-house
with 5 µm C18 beads on an Eksigent NanoLC Ultra 2D system. Target peptide
sequence analysis was performed using SIM scan mode in ion trap, followed by IT-
MS/MS fragmentation. For carbamidomethylation of unconverted cysteine, capsid
proteins of the modified nanovector was first treated with Tris(2-carboxyethyl)
phosphine hydrochloride (TCEP, 5 mM) for 30 min at 37°C, followed by reaction with
2-iodoacetamide (20 mM) at room temperature for 30 min in the dark. And then the
proteins were reduced and digested with the same procedure described above.
Dynamic light scattering (DLS) analysis and transmission electron microscope (TEM)
imaging: For gold particle conjugation to AAV2 nanoparticles, amine-functional Au
nanoparticles (Nanocs Inc, New York, NY) were modified with SANH to create free
hydrazides to the Au particles. After removing excess SANH by gel filtration columns,
the hydrazide-functionalized Au particles were then conjugated to AAV2
Ald13
for 2 hr
at RT. The hydrodynamic sizes of AAV2
Ald13
and Au-labeled AAV2
Ald13
were measured
by dynamic light scattering (Wyatt Technology, Santa Barbara, CA).
For transmission electron microscope imaging, unlabelled AAV2
Ald13
nanoparticles
were negatively stained for 10 min with 2% uranyl acetate in alcoholic solution (50%
ethanol) and were then deposited on the formvar/carbon-coated electron microscope
grids (Ted Pella, Inc., Redding, CA). The grid deposited with AAV2
Ald13
was placed on
the suspension drop containing hydrazide-funciotionalized Au particles for 30 min.
48
After washing, images of the grid were obtained using the JEOL JEM-2100 LaB6
operated at 100kV.
49
Chapter 3
Enhancing Gene Delivery of Adeno-Associated Viruses
by Cell-Permeable Peptides
Yarong Liu, Young Joo Kim, Man Ji, Jinxu Fang, Natnaree Siriwon,
Li I. Zhang, and Pin Wang
(Molecular Therapy- Methods & Clinical Development, 2014)
50
3.1 Abstract
Adeno-associated virus type 2 (AAV2) is considered a promising gene delivery vector
and has been extensively applied in several disease models; however, inefficient
transduction in various cells and tissues has limited its widespread application in many
areas of gene therapy. In this study, we have developed a general, but efficient, strategy
to enhance viral transduction, both in vitro and in vivo, by incubating viral particles
with cell-permeable peptides (CPPs). We show that CPPs increase internalization of
viral particles into cells by facilitating both energy-independent and energy-dependent
endocytosis. Moreover, CPPs can significantly enhance the endosomal escape process
of viral particles, thus enhancing viral transduction to those cells that have exhibited
very low permissiveness to AAV2 infection as a result of impaired intracellular viral
processing. We also demonstrated that this approach could be applicable to other AAV
serotypes. Thus, the membrane-penetrating ability of CPPs enables us to generate an
efficient method for enhanced gene delivery of AAV vectors, potentially facilitating its
applicability to human gene therapy.
51
3.2 Introduction
Adeno-associated virus type 2 (AAV2), a relatively well-characterized AAV serotype,
has been evaluated in gene therapy clinical trials owing to its ability to establish long-
term gene expression in both dividing and nondividing cells without known
pathological consequences of infection [6,7,8,129]. However, the transduction of
AAV2 is inefficient in a number of nonpermissive cell types [21,22,23,24], limiting its
application in many areas of gene therapy. Two obstacles have been proposed to
contribute to the inefficient infection of AAV2 in cells. First, low expression of
receptors and/or coreceptors on the cell surface of nonpermissive cells prevents
binding of the viral vector to the cell surface [30,31,32]. Second, after binding of viral
vectors to the cell surfaces and successful internalization, an impairment occurs in the
multistep intracellular trafficking process of AAV2 [24]. A novel method of enhancing
AAV2 infection could overcome these obstacles, thereby expanding the utility of
AAV2 vectors in gene therapy.
Several approaches have been explored to improve the transduction efficiency of
AAV2 in nonpermissive cell types. For example, insertion of targeting peptides into
capsid proteins of vectors enables AAV2 to bind and internalize into nonpermissive
cells via alternative receptors/coreceptors [40,41,167]. However, this type of capsid
modification could result in low production yield, reduction of vector titer, or
inefficient DNA packaging [40,43]. An alternative way to increase cell-surface
interactions of viral particles may be through cell-permeable peptides (CPPs). CPPs,
small poly-basic peptides, show a superior ability to penetrate cell membranes and
52
have been used to deliver biologically active materials, such as proteins [45], plasmid
DNAs [46], liposomes [47] and viruses [48], into various cell types and tissues. CPPs
show promise as an alternative strategy to chemical reagents that promote endosomal
membrane disruption [53,54] because CPPs can promote the destabilization of the
endosomal membrane upon acidification of the endosomal compartment without
significant toxicity [55]. For instance, arginine-rich and histidine-rich peptides have
displayed the ability to disrupt the endosomal membrane via a buffering effect within
the acidic endosomal environment [56,57,58]. Therefore, CPPs could enhance AAV2-
mediated transduction in various cell types for gene delivery to target cells and thus
overcome the obstacles noted above.
In this study, we have investigated the potential of three cell-permeable peptides (Antp,
TAT-HA2 and LAH4) to enhance AAV2-mediated gene delivery. We found that these
CPPs significantly increase AAV2-mediated transduction into tissues and cells,
including both permissive and non-permissive cells. We also demonstrated that CPPs
facilitate internalization of AAV2 via various endocytosis pathways, including the
clathrin-dependent route and energy-independent pathway. Furthermore, CPPs
showed a prominent ability to promote AAV2 particle escape from endosomal
membranes. We found that LAH4 exhibits a superior ability to increase viral
transduction via faster internalization kinetics and higher ability to penetrate
endosomal membrane, as compared to TAT-HA2 and Antp. Moreover, CPPs
complexed with AAV8 were shown to significantly improve AAV8-mediated gene
delivery to cells and tissues. These results demonstrate a general and efficient means
53
for enhancing viral vector-mediated gene delivery, both in vitro and in vivo, thus
expanding the utility of AAV for clinical applications.
3.3. Results
3.3.1 Enhanced AAV2 transduction mediated by CPPs
Our main hypothesis was that preincubation of AAV2 with cell-permeable peptides
(Antp, TAT-HA2 and LAH4; amino acid sequences shown in Table 2) could generally
enhance transduction. To test this hypothesis, each of these CPPs was incubated with
an AAV2 vector encoding GFP, and GFP expression was assayed after infection of
HEK293T cells. Increasing concentrations of CPPs (0.1-200 µM) were used against
fixed amounts of AAV2 (multiplicity of infection (MOI) of 400). As shown in Figure
16a-16c, Antp, TAT-HA2 and LAH4 induced a dose-dependent enhancement of viral
transduction in HEK293T cells. The peptide-mediated enhancement of viral
transduction was further investigated in HepG2 cells, which are known to have very
limited permissiveness to AAV2 infection [22]. Indeed, flow cytometric analysis
showed that transduction of AAV2 in HepG2 cells mediated by AAV2-CPPs was
significantly increased from 5% up to 45% (p < 0.05) as compared to that mediated by
AAV2 alone (Figure 16d-16f). LAH4 was the most effective peptide for enhancing viral
transduction in both HEK293T and HepG2 cells, as compared to that of Antp and
TAT-HA2.
54
CPP Peptide sequence
Antp
TAT-HA2
LAH4
RQIKIWFQNRRMKWKKC
CRRRQRRKKRGGDIMGEWGNEIFGAIAGFLG
KKALLALALHHLAHLALHLALALKKAC
!
Table 2. Amino acid sequences of cell-permeable peptides used in this study
a
0
10
20
30
40
50
GFP
+
cells (relative %) GFP
+
cells (relative %)
Antp concentration (µM)
Antp concentration (µM)
d
0
10
20
30
40
50
60
b
TAT-HA2 concentration (µM)
0 0.1 1 10 200 100
0
10
20
30
40
50
0 0.1 1 10 200 100
TAT-HA2 concentration (µM)
e
c
0
10
20
30
40
50
60
LAH4 concentration (µM)
0 0.1 1 10 200 100
0
10
20
30
40
50
0 0.1 1 10 200 100
LAH4 concentration (µM)
f
0 0.1 1 10 200 100
0 0.1 1 10 200 100
0
10
20
30
40
50
60
Figure 16. Antp, TAT-HA2 and LAH4 improve AAV2 transduction in permissive and non-
permissive cells. (a-c) GFP expression levels of HEK293T cells infected with AAV2
(MOI=400) alone or precomplexed with Antp (a), TAT-HA2 (b), or LAH4 (c). (d-f) GFP
expression levels of HepG2 cells infected with AAV2 (MOI=400) alone or precomplexed with
Antp (d), TAT-HA2 (e), or LAH4 (f). Error bars represent the standard deviation of the mean
from triplicate experiments.
55
To characterize the effect of these peptides on the target cells, the viability of cells was
determined by an XTT assay after incubation with either AAV2-CPP complexes or
AAV2 alone. As shown in Figure 17a, no cytotoxicity was observed when AAV2 was
preincubated with CPPs at concentrations up to 200 µM. We then investigated the
complex formation between CCPs and AAV2 by confocal imaging. The purified
AAV2-CPP complexes were overlaid to coverslips and immunostained with an
antibody specific for intact AAV2 particles. Most of the FITC-labeled CPPs were
colocalized with AAV2 signals (Figure 17b), confirming the formation of AAV2-CPP
complexes. To further characterize the AAV2-CPP complexes, the number of CPPs
functionally bound to individual AAV2 particles was determined by measuring the
spectroscopic property of the purified AAV2-CPP complexes at different initial ratios
as described in Materials and Methods. We found that the average maximal numbers
of Antp, TAT-HA2 and LAH4 bound to each AAV2 particle are approximately 2540,
2478, and 2706, respectively (Figure 17c).
Because we found that addition of CPPs could enhance transduction of multiple cell
types, we next hypothesized that combining these CPPs with our viral vectors could
reduce the amount of vectors necessary to achieve comparable levels of transduction.
To test this hypothesis, we incubated constant amounts of Antp, TAT-HA2 or LAH4
(200 µM) with increasing amounts of AAV2 particles and infected 293T cells at a MOI
of 10−2500. Based on the results shown in Figure 17d, the concentration of CPPs (200
µM) was saturating, even for the highest amount of AAV2, with MOI of 2500. The
56
data in Figure 17d show that a ~3-, 10- or 20-fold lower titer of AAV2 is sufficient for
similar transduction level when AAV2 is preincubated with Antp, TAT-HA2 or LAH4,
respectively. The superior potential of LAH4 on enhancing viral transduction was
further confirmed by its 10-fold and 15-fold enhancement on viral transduction titers
in HEK293T and HepG2 cells, respectively (Figure 17e).
To examine whether the positively charged nature of CPPs is critical for enhancing
viral transduction via complex formation with viruses, Antp, TAT-HA2 and LAH4
(200 µM) were incubated with AAV2 particles in increasing concentrations of
phosphate buffer (0.1-0.5M). As shown in Figure 17f, increasing concentrations of
phosphate did not affect AAV2 transduction, but did significantly reduce the ability of
Antp, TAT-HA2, and LAH4 to enhance viral transduction, indicating electrostatic
forces in the formation of complexes between AAV2 and peptides are critical for
enhancing viral transduction.
3.3.2 Enhanced viral uptake with faster kinetics by CPPs
We next sought to determine whether the enhanced viral transduction mediated by
CPPs resulted from an increased cellular uptake of viral particles in the presence of
CPPs. To measure the cellular uptake of AAV2, Alexa488-labeled AAV2 particles were
preincubated with CPPs (200 µM), and the uptake of viral particles was determined by
flow cytometry 30 min after incubation with cells. As shown in Figure 18a and 18b, a
significant increase in integrated mean fluorescence intensity (iMFI) of viruses was
observed when cells were incubated with AAV2-CPP complexes, as compared to
57
AAV2 alone, indicating that CPPs can facilitate the uptake of viral particles (p < 0.01).
LAH4 showed the most prominent enhancement of AAV2 uptake as compared to
TAT-HA2 and LAH4 (p < 0.01). Indeed, LAH4 increased cellular uptake of AAV2 at a
much faster rate compared to TAT-HA2 and Antp, as shown in Figure 18c.
Figure 17. Interaction of CPPs with AAV2 facilitates enhanced viral transduction. (a) Cellular
cytotoxicity of AAV2-CPP complexes. HEK293T cells were incubated with the AAV2-CPP
complexes with increasing concentrations of peptides for 48 h. Subsequently cell viability was
determined using the XTT assay. (b) Visualization of the AAV2-CPP complexes. AAV2 were
incubated with FITC-CPP (green) and subsequently probed with a mouse monoclonal
antibody (A20) specific for intact AAV2 (red). Overlaping green and red signals appear as
yellow in the merged image. Scale bar represents 5 µm. (c) Quantitative analysis of the number
of remaining CPPs per AAV2 particle. The numbers were determined by measuring the
spectroscopic property of the purified AAV2-CPP complexes 30 min after incubation. (d) GFP
expression levels of HEK293T cells infected with increasing MOI of AAV2 in the presence of a
fixed concentration of CPPs (200 µM). (e) Transduction titers of AAV2 in the presence or in
the absence of LAH4 in HEK293T cells and HepG2 cells. (f) GFP expression levels of
HEK293T cells infected with AAV2 alone or the AAV2-CPP complexes formed in the
presence of increasing concentrations of buffered phosphate (0.1-0.5 M, pH 7.4). All error bars
represent the standard deviation of the mean from triplicate experiments.
58
Figure 18. CPPs enhance AAV2 uptake by cells. (a, b) Internalization of AAV2 particles in the
presence of CPPs to HEK293T cells (a) or HepG2 cells (b). AAV2 particles were labeled with
Alexa488 dye and incubated with CPPs for 30 min at 37°C. Cells were incubated with AAV2-
CPP complexes for 30 min at 37°C. The cellular uptake of dye-labeled AAV2 was determined
by measuring dye fluorescence using flow cytometry. (c) Internalization kinetics of AAV2
particles in the presence of CPPs. AAV2 particles were labeled with Alexa488 dye and
incubated with CPPs for 30 min at 37°C. HEK293T cells were incubated with AAV2-CPP
complexes at 37°C for different indicated time points. After incubation, the cells were washed
to remove the unbound complexes, and cellular uptake of dye-labeled AAV2 was determined
by measuring dye fluorescence using flow cytometry.
3.3.3 Entry mechanism of AAV2-CPP complexes
Having shown that CPPs enhanced uptake of viral particles into the cells, we next
sought to examine the mechanisms by which CPPs facilitate this viral uptake. We first
investigated whether CPPs could enhance viral uptake in cells at 4°C because AAV2
internalizes into cells primarily through clathrin-coated pits in an energy-dependent
manner [168,169,170]. As shown in Figure 19a, when cells were incubated with AAV2-
59
CPP complexes at 4°C for 30 min, Antp and TAT-HA2 modestly increased viral
uptake compared to AAV2 alone, while LAH4 significantly enhanced the
internalization of AAV2, indicating that LAH4 is the most efficient in improving
energy-independent uptake of AAV2. However, the enhanced effect of CPPs on viral
uptake was more apparent when AAV2-CPP complexes were incubated with cells at 37
°C, indicating that energy-dependent entry routes are also involved in the
internalization of complexes.
It has been proposed that clathrin-, caveolin- and macropinocytosis-mediated
pathways are the main routes of endocytosis for viruses, proteins, and nanoparticles.
To assess the role of these three endocytic pathways in the entry of AAV2-CPP
complexes, drug-inhibition assays were performed. Chlorpromazine (CPZ) is known
to block clathrin-mediated internalization [171], while filipin is a cholesterol-binding
reagent that can inhibit the caveolin-dependent entry pathway [172], and amiloride is
used to inhibit the macropinocytosis [173] by preventing induction of membrane
ruffling. The addition of CPZ significantly decreased the transduction of AAV2-Antp,
AAV2-TAT-HA2, and AAV2-LAH4 by 81.4%, 69.5% and 18%, respectively (Figure
19b), indicating that the clathrin-mediated pathway is involved in endocytosis of all
three AAV2-CPP complexes and that AAV2-LAH4 is the least dependent on clathrin-
mediated internalization. No significant inhibitory effect of filipin was observed,
suggesting that caveolin may not be involved in the entry of AAV2-CPP complexes.
Moreover, only amiloride significantly decreased the transduction mediated by AAV2-
60
LAH4 complexes, suggesting that the macropinocytosis-mediated pathway is also
involved in the internalization of AAV2-LAH4 complexes, but not AAV2-Antp or
AAV2-TAT-HA2 complexes.
It is generally believed that heparin sulfate proteoglycan (HSPG) is the primary
attachment receptor that mediates AAV2 binding to the surface of many cell types
[174]. To assay whether HSPG is involved in the mechanism of CPP transduction
enhancement, heparin and HSPG antibody were used to competitively bind to HSPG
receptors [26,174]. As shown in Figure 19c and 19d, both heparin and HSPG antibody
treatments significantly reduced AAV2 transduction, while no inhibitory effects on
viral transduction mediated by AAV2-CPP complexes were observed, indicating that
CPPs facilitate viral uptake through receptors other than HSPG.
3.3.4 Involvement of endosomes in viral transduction mediated by AAV2-CPP
complexes
Endosomal transport of viral particles is thought to be a critical step for viruses to
achieve successful infection. For AAV2, it has been proposed that processing through
endosomal compartments, including early and late endosomes [30,34,35], is required
to induce a conformational rearrangement of the viral capsid for nuclear transport and
uncoating [36]. To investigate whether the endosomal environments are required for
productive infection of AAV2-CPP complexes, bafilomycin A1, which specifically
inhibits vacuolar proton ATPases [175], was used to block low pH-associated
endosomal processes in cells. As shown in Figure 20a, the flow cytometric analysis of
61
GFP
+
cells showed that transduction mediated by AAV2-CPP complexes was
significantly inhibited by bafilomycin A1, indicating that low-pH endosomal processes
are essential for productive infection of AAV2-CPP complexes.
Figure 19. Entry mechanisms of AAV2-CPP complexes. (a) The internalization of AAV2 or
AAV2-CPP complexes in cells after 30 min incubation at 37°C or 4°C. (b) The effect of
inhibitory drugs on viral transduction of cells infected by AAV2 or AAV2-CPP complexes.
HEK293T cells were preincubated with chlorpromazine (CPZ, 30 nM), filipin (15 nM), or
Amiloride (1 mM) for 30 min at 37 °C. Treated and untreated cells were spin-infected with the
AAV2 alone or complexes of AAV2 with peptides (final concentration: 200 µM) for 90 min in
the presence of drugs. GFP expression was analyzed 2 days after infection. All error bars
represent the standard deviation of the mean from triplicate experiments. Asterisks indicate
statistical significance compared to the no drug treatment group (* P< 0.05, ** P<0.01). (c, d)
The effect of HSPG receptor blocking by heparin (c) or HSPG antibody (d) on viral
transduction mediated by AAV2 or AAV2-CPP complexes. The cells were incubated with
heparin (200 µg/ml) or HSPG Ab (1:100) for 30 min at 37°C. The treated and untreated cells
were spin-infected with the AAV2 alone or complexes of AAV2 with peptides (final
concentration: 200 µM) for 90 min. GFP expression was analyzed 2 days after infection. All
data are shown as the means of triplicate experiments. Asterisks indicate statistical significance
compared to the no drug treatment group (** P<0.01).
62
To further investigate the functional involvement of different endosomes, including
early and late endosomes, in AAV2-CPP-mediated transduction, the dominant-
negative mutants of Rab proteins were employed to perturb either early (Rab5) [176]
or late (Rab7) [177] endosome function. HEK293T cells transfected with wild type or
the dominant-negative form of Rab5 or Rab7 were transduced with AAV2 alone or
AAV-CPP complexes. As shown in Figure 20b, expression of dominant-negative Rab5
significantly reduced the transduction rate of AAV2 and AAV2-CPPs, as compared
with the transduction of wild-type Rab5-expressing cells, indicating that early
endosomes are required for the AAV2-CPP infection pathway. AAV2-CPP-mediated
transduction in cells expressing dominant-negative Rab7 was also remarkably
decreased, compared to that of wild type-expressing cells (Figure 20c), suggesting that
AAV2-CPP complexes traffic through both early and late endosomes for successful
transduction.
It has been proposed that the increasingly acidic environment found in both early
endosome (~pH 6.0) and late endosome (~pH 5.0) favors membrane penetration of
AAV2 particles. Moreover, it has been reported that the acidic pH of endosomes could
affect some peptide structures, such as histidine-rich molecules and poly(amido amine)
polymers [56,57,58], resulting in disruption of the endosomal membrane. To
investigate whether the CPPs could assist endosomal escape of AAV2 particles for
enhanced transduction, we designed a membrane penetration assay using a transwell,
63
in which a lipid bilayer membrane was formed on the bottom of the upper
compartment to mimic endosomal membranes [178]. This assay enabled us to
quantify the number of viruses transiting through the membrane toward the lower
compartment when either AAV2 or AAV2-CPP complexes were exposed to PBS at
different pH values (7.4, 6.0 or 5.0) in the upper compartment. These pH values mimic
the cytosolic, early and late endosomal environments. As shown in Figure 20d, a
moderate enhancement of virus transport was observed in AAV2-Antp complexes
treated with PBS with different pH values, compared to that of AAV2 at the same pH
values, suggesting that Antp could facilitate viral penetration of multiple membranes,
including cell membranes and endosomal membranes. Moreover, a significant
enhancement of genome copy number was observed in AAV2-TAT-HA2 complexes
pretreated with buffer of pH 6.0 and 5.0, as compared to that of AAV2-Antp,
indicating that TAT-HA2 could be more beneficial for endosomal escape of AAV2 in
an endosomal environment. In addition, AAV2-LAH4 complexes treated with PBS
with pH 7.4, 6.0 and 5.0 showed a significant movement of particles across the
membrane, as compared to that of AAV2-TAT-HA2 complexes with the same
treatment, suggesting that LAH4 peptide is the most efficient one in facilitating AAV2
membrane penetration, including cell membranes and endosomal membranes.
64
Figure 20. The effect of CPPs on the endosomal escape of AAV2 particles. (a) The effect of
bafilomycin-A1 (BAF) on viral transduction mediated by AAV2-CPP complexes. HEK293T
cells were preincubated with BAF for 30min at 37 °C. The cells were then spin-infected with
AAV2 or AAV2-CPP complexes for 90 min in the presence of BAF. GFP expression was
analyzed 2 days after infection. All data are shown as the means of triplicate experiments.
Asterisks indicate comparisons to the no drug treatment group (* P<0.05). (b, c) Functional
involvement of early and late endosomes in the viral transduction mediated by AAV2-CPP
complexes. HEK293T cells were transiently transfected with a wild-type or dominant-negative
mutant form of Rab5 (b) or Rab7 (c). Twenty-four h after transfection, cells were spin-infected
with AAV2 or AAV2-CPP complexes for 90 min. GFP expression was analyzed 2 days after
infection. All data are shown as the means of triplicate experiments. Asterisks indicate
statistical significance compared to the wild-type form of Rab treatment groups (* P< 0.05). (d)
The effect of CPPs on the endosomal escape of AAV2 particles. AAV2 (107 genome copies per
well) or AAV2-CPP complexes (final concentration of CPP: 200 µM) in PBS with different pH
values (pH 7.4, 6.0 or 5.0) were placed in the upper compartment of a 24-well transwell plate.
After incubation at 37 °C for 12 h, the AAV2 particles transferred from upper compartment to
the lower compartment with pH 7.4 PBS were collected. The intracellular viral genome copies
were quantified by qPCR. (e) The GFP expression of NIH3T3 cells infected by AAV2 alone or
AAV2-CPP complexes (final concentration of CPP: 200 µM). All error bars represent the
standard deviation of the mean from triplicate experiments. Asterisks indicate statistical
significance compared to the AAV2 alone treatment group (* P< 0.05, ** P<0.01). (f)
Transduction titers of AAV2 in the presence or in the absence of LAH4 (final concentration:
200 µM) in NIH3T3 cells. All data are shown as the means of triplicate experiments.
65
To further confirm that CPPs could facilitate endosomal escape of AAV2 particles, we
examined the transduction of AAV2-CPP complexes in NIH3T3 cells, in which
limited AAV2-mediated transduction has been attributed to impaired intracellular
trafficking [24,179]. As shown in Figure 20e, Antp significantly enhanced AAV2-
mediated transduction in NIH3T3 cells, while TAT-HA2 induced a higher GFP
expression compared to Antp. Moreover, the AAV2-LAH4 complex demonstrated the
highest GFP expression in NIH3T3 cells, consistent with our previous experiments
showing the superior ability of LAH4 in facilitating the endosomal escape of AAV2
particles. Indeed, we found that inclusion of AAV2-LAH4 led to a 25-fold
enhancement of the transduction unit titer (∼25x106 TU/ml) in NIT3T3 cells, as
compared to that of AAV2 (∼1x106 TU/ml) (Figure 20f).
3.3.5 CPPs enhance viral transduction of AAV2 in primary cells and tissues
We have shown that cell-permeable peptides can enhance viral-mediated transduction
in both permissive (HEK293) and nonpermissive (HepG2, NIH3T3) cell lines. Next,
we determined whether this enhanced infection in the presence of CPPs could be
achieved in primary cells. To determine this, mouse bone marrow-derived cells
(BMDCs) were incubated with either AAV2 alone or AAV2-CPP complexes, and GFP
expression was detected by flow cytometry 2 days after infection. As shown in Figure
21a, CPPs indeed significantly enhanced viral transduction in BMDCs, compared to
AAV2 alone. As observed in the cell lines tested, LAH4 again showed a superior
efficiency in increasing viral transduction compared to TAT-HA2 and Antp. Next, we
66
tested the effect of CPPs on the viral transduction in murine primary mesenchymal
stem cells (MSCs), in which AAV2-mediated transduction is inefficient [24]. As shown
in Figure 21b, the transduction efficiency mediated by AAV2-CPPs was 3 to 4-fold
higher than that of the AAV2 alone in murine MSCs.
Figure 21. CPPs enhance viral transduction in primary cells and tissues. (a) The GFP
expression levels of BMDC cells infected by AAV2 alone or AAV2-CPP complexes. (b)
Enhancement of gene delivery of AAV2 in the presence of CPPs in murine MSCs. (c, d) CPPs
facilitate gene delivery of AAV2 in mouse cochlear neuron. Mouse cochlear neurons were
treated with AAV2-CPP complexes or AAV2 alone for 4 h, and the medium was replaced by
fresh medium. After 7 days, tissues were imaged by fluorescence microscopy (c). Upper: GFP
fluorescence. Lower: Myosin VI image. Scale bar represents 100 µm. (d) Quantification of GFP
expression levels in cochlear neuron. To quantify GFP-positive cells, 4 regions of interest
(ROI) were randomly chosen per image at x10 magnification. The data are expressed as % of
total area of GFP-positive in the region of AAV-CPP-treated tissues normalized by that of
tissues treated by AAV2 alone. All data are shown as the means of triplicate experiments.
Asterisks indicate statistical significance compared to the AAV2 alone treatment group (* P<
0.05, ** P<0.01).
67
Additionally, we assessed the effects of the CPPs in cochlear gene transfer, which has
shown promise as a potential strategy for the treatment of hearing loss [180]. Previous
studies suggested that AAV2 was unable to efficiently transduce the cells of the cochlea
in vitro [181]. Here, AAV2-CPP complexes displayed markedly enhanced GFP
expression in cochlear tissues when compared to that of AAV2 alone (Figure 21c), as
quantified in Figure 21d. It is also noteworthy that the cochlear neurons infected by
AAV2-CPP complexes showed strong GFP expression in spiral ganglion cells (SGCs),
while gene expression was not detected in SGCs of cochlear neurons transduced by
AAV2 alone (Figure 21c). Previous research showed that no spiral ganglion cells could
be transduced by a majority of serotypes of hybrid recombinant AAV vectors [182],
demonstrating that CPPs could enhance gene transfer in tissues that cannot be
achieved by alternative AAV serotypes. Taken together, this enhanced transduction
mediated by AAV2-CPP complexes occurs not only in cell lines, but also in primary
cells and tissues.
3.3.6 CPPs enhance AAV2-mediated gene delivery in vivo
Recombinant AAV2 vector-based muscle gene therapy has been widely explored for
inherited diseases, such as muscular dystrophies, by the ability of AAV2 vector to
establish persistent transgene expression in muscles. However, it has been reported
that low viral titers have limited the success of clinical trials [183]. Many different
approaches have been tested to increase the efficacy of AAV2-mediated gene delivery
to muscles in order to improve the therapeutic benefit in clinical trials. Here, we
68
investigated the potential effects of the cell-permeable peptides in an in vivo setting by
examining gene transfer in muscle. Complexes of AAV2 (10
9
particles) preincubated
with CPPs (final concentration: 10 µM) and AAV2 alone were injected
intramuscularly into mice, and the GFP expression in muscles was detected 14 days or
30 days post-administration. As shown in Figure 22a, intramuscular injection of AAV2
in the presence of CPPs significantly increased the GFP expression in muscles as
compared to that of AAV2 alone 14 days post-injection. This enhanced gene
expression via CPPs could be maintained for the duration of the 30-day study,
demonstrating that the enhancement is of the long-lived nature. The potential of cell-
permeable peptides on promoting in vivo gene delivery was further demonstrated by
the quantification data shown in Figure 22b. In addition, we investigated whether
CPPs could alter the biodistribution of viral vectors. As shown in Figure 22c, the
vector DNA was mainly present in muscles of both AAV2-injected and AAV2-LAH4-
injected mice; no significantly different distribution of viral vectors was observed
between these two groups.
Next, we determined whether CPPs could cause any cytotoxicity. As shown in Figure
23a, no significant histopathologic change was observed for the muscle tissues of the
mice treated with AAV2-CPPs or with AAV2 alone. Immunohistochemical analysis
revealed that AAV2-CPPs-injected muscles did not show higher rates of infiltration of
CD4
+
and CD8
+
T lymphocytes than that of the AAV2-injected muscles, indicating
that CPPs do not induce detectable cytotoxicity. To further assay for induction of
69
innate immune gene transcripts following the intramuscular injection of AAV2-CPPs
or AAV2, we performed quantitative RT-PCR 14 days post-injection to detect the
inflammatory cytokine transcript levels. Transcript levels of IL-1b, IL-6 and TNF-α
were quantified and normalized by those in AAV2-treated mice. As shown in Figure
23b, no significant difference of induced cytokine transcripts was observed in the
AAV2-CPP-injected muscles as compared to that in the AAV2-injected muscles.
Taken together, CPPs were shown to significantly enhance gene delivery in vivo
without detectable cytotoxicity.
70
Figure 22. CPPs facilitate gene delivery of AAV2 in mouse muscles. (a) Transgene expression
in muscle tissues from BALB/c mice injected with AAV2 or AAV2-CPP complexes. Muscle
cross sections were taken at day 14 or day 30 after intramuscular injection of AAV2 alone (109
particles/ mice) or AAV2-CPP complexes from injected legs. Muscle sections were stained
with GFP antibody (red), followed by nuclear staining (blue). Scale bar represents 100 µm. (b)
Quantification of GFP expression levels in muscles shown in (a). To quantify GFP-positive
cells, 4 regions of interest (ROI) were randomly chosen per image at x10 magnification. The
data are expressed as % of total nuclear area stained by GFP in the region. All error bars
represent the standard deviation of the mean from triplicate experiments. Asterisks indicate
statistical significance compared to the AAV2 alone treatment group (** P<0.01). (c)
Biodistribution of viral vectors in AAV2-injected and AAV2-LAH4-injected mice. Organs
were collected from AAV2-injected and AAV2-LAH4-injected mice 14 days post-
administration. AAV2 genome copies were assessed by qPCR assay. Levels of vectors were
standardized using primers against housekeeping gene Apob. All error bars represent the
standard deviation of the mean from triplicate experiments.
71
Figure 23. No cytotoxicity of CPPs was detected in vivo. (a) Analysis of T lymphocyte
infiltration in muscle tissues from BALB/c mice injected with AAV2 or AAV2-CPP complexes
14 days post-administration. Serial cross-sections were stained with hematoxylin and eosin
(H&E), and were immunohistochemically stained with antibodies against CD4 and CD8 (red)
followed by nuclear staining (blue). Scale bar represents 100 µm. (b) Analysis of inflammation
markers in muscles from mice injected with AAV2 or AAV2-CPP complexes 14 days post-
administration. Total RNA was isolated from muscles and mRNA levels of IL-1b, IL-6 and
TNF-α were assessed by qPCR. 2ΔCt (ΔCt=-Ctcytokine+CtGAPDH) method was used to
calculate the relative cytokine expression. All of the data were then normalized by the mean
percentage of cytokine expression in AAV2-injected mice. All error bars represent the
standard deviation of the mean from triplicate experiments.
72
3.3.7 CPPs increase viral transduction of AAV8 both in vitro and in vivo
We have showed that CPPs could significantly enhance AAV2-mediated gene delivery
in various cells and tissues both in vitro and in vivo. It would be beneficial if this
approach could be applicable to other serotypes, such as AAV8, which is more
frequently used in recent clinical trials due to its unique in vivo transduction profiles
[184,185]. We first investigated whether CPPs could increase AAV8-mediated gene
delivery in cell lines, where it shows low transduction efficiency. To test this, each of
CPPs was incubated with an AAV8 vector encoding GFP, and GFP expression was
assayed after transduction of HEK293T or Huh7 cells. Increasing concentrations of
CPPs (0.1-200 µM) were used with a fixed amount of AAV8 (MOI=1000). As shown
in Figure 24a-24c, Antp, TAT-HA2 and LAH4 induced a dose-dependent
enhancement of viral transduction in HEK293T cells. In addition, transduction of
AAV8 in Huh7 cells mediated by AAV8-CPPs was significantly increased as compared
to that mediated by AAV8 alone (Figure 24d-24f).
Moreover, we investigated the potential effects of the cell-permeable peptides on
AAV8-mediated gene transfer in muscles. Complexes of AAV8 (10
8
particles)
preincubated with or without CPPs (final concentration: 10 µM) were injected
intramuscularly into mice, and the GFP expression in muscles was analyzed 14 days
post-administration. As shown in Figure 25a, no significant histopathologic change
was observed for the muscle tissues from the mice treated with the AAV8-CPPs as
73
compared to that treated with AAV8 alone, confirming that CPPs did not induce
detectable cytotoxicity. More importantly, intramuscular injection of AAV8 in the
presence of CPPs significantly increased the GFP expression in muscles as compared
to that of AAV8 alone (Figure 25b). Thus, the fact that CPPs can enhance both AAV2
and AAV8 suggests that the approach to using CPP to improve gene delivery could be
potentially applicable to various other AAV serotypes.
3.4. Discussion
Many efforts have been made to improve the efficacy of AAV2-mediated gene transfer
in cells, especially in nonpermissive cells, by increasing the internalization and
endosomal escape process of viral particles. Cell-permeable peptides have been used as
a promising carrier to deliver bioactive hydrophilic molecules, such as DNA, proteins,
or liposomes into cells, based on their ability to penetrate cell membranes in a
receptor-independent manner and to facilitate endosomal escape of molecules by
disrupting endosomal membranes [45,46,47,48,53,54,55]. The present investigation
was designed to elucidate the role of three different CPPs (Antp, TAT-HA2 and LAH4)
in AAV2-mediated gene delivery. The data presented here show that a simple pre-
incubation of these CPP peptides with AAV2 particles markedly enhanced cellular
entry and endosomal escape of AAV2 particles, thus improving both in vitro gene
expression and in vivo gene delivery.
74
Figure 24. Antp, TAT-HA2 and LAH4 improve AAV8 transduction in target cells. (a-c) GFP
expression levels in HEK293T cells infected with AAV8 (MOI=1000) alone or precomplexed
with Antp (a), TAT-HA2 (b), or LAH4 (c). (d-f) GFP expression levels in Huh7 cells infected
with AAV8 (MOI=1000) alone or precomplexed with Antp (d), TAT-HA2 (e), or LAH4 (f).
Error bars represent the standard deviation of the mean from triplicate experiments.
Figure 25. CPPs facilitate AAV8-mediated gene delivery in mouse muscles. (a, b)
Histochemical and immunofluorescence analyses of muscle tissues from BALB/c mice injected
with AAV8 or AAV8-CPP complexes. Muscle cross sections were taken at day 14 after
intramuscular injection of AAV8 alone (108 particles/ mice) or AAV8-CPP complexes. Muscle
sections were stained with hematoxylin and eosin (a), or GFP antibody (red), followed by
nuclear staining (blue, b). Scale bar represents 100 µm.
75
The ability of CPPs to enhance viral uptake into cells was demonstrated as a key
mechanism promoting improved viral-mediated transduction of cells. Moreover, the
LAH4 peptide showed a superior ability to facilitate viral uptake with a faster rate
compared to Antp and TAT-HA2, even though the number of peptide molecules
bound to one particle was similar. CPPs facilitate the energy-independent step of
AAV2 uptake. At the same time, however, uptake requires energy-dependent
internalization to achieve optimal efficiency of transduction. Although it is generally
believed that AAV2 is internalized into cells via clathrin-coated pits, this study showed
that LAH4 is able to facilitate the internalization of AAV2 through both clathrin- and
macropinocytosis-mediated pathways. Interestingly, the competition assay suggested
that CPPs facilitate the uptake of AAV2 through receptors other than HSPG. Although
the detailed molecular mechanism underlying the different internalization processes of
AAV-CPP complexes remains unclear, our data suggest that the differential
involvement of endocytic pathways could contribute to the enhanced viral
transduction in nonpermissive cell lines with limited expression of HSPG receptors
and/or coreceptors.
It is generally believed that AAV2 vectors must transit through the early and late
endosomes for successful infection [22,34]. Our experiments with dominant-negative
mutants of Rab constructs suggested that both early and late endosomes are required
for efficient transduction of AAV2-CPPs. Moreover, viral escape from the endosomal
membrane to cytosol has been considered another key barrier to viral transduction in
76
target cells. Our previous data suggested that exposure to the acidic endosomal
environment could trigger AAV2 to penetrate endosomal membranes [178]. Here, by
using an in vitro transwell assay, we show that CPPs could further enhance viral
penetration from endosomal membranes when exposed to the acidic endosomal
environment (pH 6.0 and 5.0). The ability of CPPs to facilitate endosomal escape of
viral particles was further confirmed by their significant enhancement on viral
transduction in NIH3T3 cells, in which previous work showed that impaired
endosomal processing limited viral transduction [24].
It has been reported that AAV2-mediated transduction of primary cells, such as
BMDCs and MSCs, is inefficient. Tyrosine phosphorylation of AAV2 capsid proteins
has been suggested to promote ubiquitination and degradation of the AAV2 capsid,
leading to impairment of viral transport to the nucleus and thus poor transduction
[186,187]. This hypothesis was supported by the enhanced transduction of murine
MSCs mediated by AAV2 tyrosine-mutant vectors [188]. Here, we show that
preincubation of AAV2 with CPPs significantly enhances viral transduction in both
murine BMDCs and MSCs, offering a general and simple method that can bypass the
need for manipulating AAV2 vector. Our in vitro transwell assay demonstrates that the
addition of CPPs allows the virus to penetrate multiple types of membranes, including
proteasomal membranes, and may enable viral particles to escape from the
degradation process.
Alternative serotypes were usually proposed to overcome the limitations of AAV2 such
77
as poor infectivity in certain tissues. However, certain cells such as the spiral ganglion
cells in cochlear neurons are resistant to transduction by a majority of AAV serotypes
[182]. In this study, we showed that AAV2-CPP complexes could successfully
transduce the spiral ganglion cells, raising the possibility that CPPs complexed with
viral particles might be useful for cochlear gene therapy. We also showed that CPPs
significantly enhance AAV8-mediated gene delivery both in vitro and in vivo,
suggesting that this approach could be applicable to other AAV serotypes. In
conclusion, our studies provided a general method to markedly improve AAV-
mediated gene delivery to cells and tissues using cell-permeable peptides.
3.5. Materials and Methods
Cell lines and reagents. HEK293, HEK293T and HepG2 cells were cultured in
Dulbecco’s modified Eagle’s medium (DMEM; Gibco) supplemented with 10% FBS
(Sigma-Aldrich, St. Louis, MO) and 2 mM L-glutamine (Hyclone Laboratories, Inc.,
Omaha, NE). Total bone marrow cells were harvested from naïve C57BL/6 mice, and
BMDCs were obtained by culturing bone marrow cells in the presence of GM-CSF.
Murine bone marrow-derived MSCs were isolated from 6- to 8-week-old C57BL/6
mice. Briefly, bone marrow cells were flushed out with DMEM supplemented with 15%
FBS, centrifuged down at 1200 rpm and plated in a 125cm
2
flask. The medium was
changed after 24 h and then every 12 h until 72 h after collection to remove excess
CD45+ cells. MSCs were collected after filtering out CD45+ cells using CD45
microbeads (MACS Miltenyi Biotech) according to the manufacturer’s protocol.
78
Bafilomycin A1, chlorpromazine, filipin, heparin and amiloride were obtained from
Sigma-Aldrich and used at appropriate concentrations according to the manufacturer’s
protocols. Anti-heparin/heparin sulfate antibody (MAB2040) was purchased from
Millipore (Billerica, MA). Mouse monoclonal antibody against the intact AAV2 (A20)
was purchased from American Research Products, Inc. (Belmont, MA). Alexa647-
conjugated goat anti-mouse IgG antibody was obtained from Invitrogen (Carlsbad,
CA).
Peptides. The Antp (amino acid sequence: RQIKIWFQNRRMKWKKC), TAT-HA2
(amino acid sequence: CRRRQRRKKRGGDIMGEWGNEIFGAIAGFLG), LAH4
(amino acid sequence: KKALLALALHHLAHLALHLALALKKAC) and FITC-labeled
Antp, FITC-labeled TAT-HA2 and FITC-labeled LAH4 peptides were synthesized by
GenScript Inc. (Piscataway, NJ). Peptides were dissolved in deionized water (stock
solution, 10mM).
AAV production. Recombinant AAV2 and AAV8 vectors were produced in HEK293
cells as previously described [178]. Forty 15-cm dishes of subconfluent HEK293 cells
were triple-transfected with 650 µg each of AAV cis-plasmid and AAV2 or AAV8
trans-plasmid containing the rep and cap genes and 1300 µg of the adenovirus helper
plasmid pΔF6 using the calcium phosphate precipitation method. For AAV2
production, after an additional 16 h of incubation, the medium was replaced with fresh
medium. The cells were harvested at 3 days post-transfection, followed by three cycles
of freeze and thaw. For AAV8 vectors, the virus supernatants were harvested every 12
79
h and replaced with fresh medium for 7 days. AAV2 or AAV8 viruses were then
purified by cesium chloride gradient density centrifugation [189] at 25,000 rpm and
15°C for 20 h (Optima L-90 K Ultracentrifuge, SW-28 rotor, Beckman Coulter, Brea,
CA). Viral particles recovered from the first round of ultracentrifugation were pooled
and subjected to isopycnic separation by a second CsCl centrifugation at 13,000 rpm
and 15°C for 20 h in a SW-32 Ti rotor. Fractions containing AAV2 or AAV8
determined by refractive index were further desalted in PBS using an Amicon Ultra
100,000 MWCO centrifugal filter device (Millipore, Billerica, MA).
Plasmids. The plasmid encoding the dominant-negative mutant of DsRed-Rab7
(Rab7T22N) was generated by site-directed mutagenesis as described [190]. The
constructs for wild-type and dominant-negative forms of DsRed-Rab5 and DeRed-
Rab7 were obtained from Addgene (Cambridge, MA).
Cell infection. Purified AAV2 particles (MOI of 400) or AAV8 (MOI of 1000) were
preincubated with the peptides (final concentration: 0.1-200 µM) in PBS for 30 min at
37 °C. The AAV alone or AAV-peptide complexes were then added to the HEK293T
cells, HepG2 cells or Huh7 (2 × 10
4
cells per well) in a 96-well culture dish by spin
infection at 2,500 rpm for 90 min at 25°C using a Sorvall Legend centrifuge. The
medium was replaced with fresh medium after an additional 3 h of incubation. GFP
+
cells were detected by flow cytometry 2 days post-infection.
80
To assess the degree to which peptides influenced the titer necessary for expression,
peptides (fixed concentration: 200 µM) were preincubated with increasing MOI of
AAV2. HEK293T cells were then spin-infected with the complexes for 90 min. GFP
expression was analyzed 2 days after infection.
For viral transduction with drug-treated cells, HEK293T cells or HepG2 cells were pre-
incubated with Bafilomycin A1 (BAF, 50 nM), Chlorpromazine (CPZ, 25 µg/ml),
filipin (5 µg/ml), Amiloride (1 mM), heparin (200 µg/ml) or HSPG Ab (1:100) for 30
min at 37°C. The treated and untreated cells were spin-infected with the AAV2 alone
or complexes of AAV2 with peptides (final concentration: 200 µM) for 90 min. GFP
expression was analyzed 2 days after infection.
For viral transduction with PO 4
−
blocking, AAV2 alone or the complexes were
incubated with increasing concentrations of phosphate (0.1-0.5 M) in the
preincubation buffer, and the cells were then spin-infected at 2500 rpm for 90 min at
30 °C using a Sorval Legend centrifuge (Newport Pagnell, England) [178]. After an
additional 3h of incubation at 37°C, the medium was then removed and replaced with
fresh medium and cultured for 72h before flow cytometry analysis of GFP
+
cells. For
viral transduction with Rab protein-expressing cells, HEK293T cells were seeded in a
24-well dish overnight at 37°C. The cells were then transfected with DsRed-Rab5 or -
Rab7 (either wild-type or dominant-negative mutants) by the calcium phosphate
precipitation method. After 4 h of incubation at 37 °C, the media were then replaced
with fresh D10 media (DMEM containing 10% FBS). At 24 h post-transfection, the
81
complexes of AAV2 with peptides were spin-infected with the transfected cells for 90
min, and the percentage of GFP-positive cells was analyzed by flow cytometry at 48 h
post-infection.
In vitro cytotoxicity. HEK293T cells were plated at a density of 5 x 10
3
cells per well in
96-well plates and grown for 6 h. The cells were then exposed to complexes of AAV2
with different concentrations of peptides for 48 h, and cell viability was assessed using
the Cell Proliferation Kit II (XTT assay) from Roche Applied Science according to the
manufacturer's instructions. Cell viability percentage was determined by subtracting
absorbance values obtained from media-only wells from drug-treated wells and then
normalizing to the control cells without complexes.
In vitro internalization study. AAV2 (1 × 10
10
particles) was incubated with 50 nmol of
Alexa488-TFP ester (Invitrogen) for 2 h in 0.1 M sodium bicarbonate buffer (pH=9.3).
After 2 h incubation, the reaction was stopped, and unbound dye molecules were
removed via buffer exchange into PBS (pH=7.4) using a Zeba desalting spin column
(Fisher Scientific). The Alexa488-labeled AAV2 were then incubated with different
concentrations of peptides for 30min. HEK293T or HepG2 cells were then incubated
with the complexes (MOI=400) for 30 min at 37 °C. After incubation, the cells were
washed and treated with trypsin to eliminate unbound particles and those not uptaken.
Cellular uptake of AAV particles was determined by measuring Alexa488 fluorescence
using flow cytometry.
82
For internalization kinetics of complexes, Alexa488-labeled AAV2 was preincubated
with fixed concentration of peptides (200 µM) for 30 min. Then the complexes were
incubated with cells for different time points (5, 15, 30, 60 and 120 min). After
incubation, the cells were washed and treated with trypsin to eliminate unbound
particles and those not uptaken. Cellular uptake of AAV particles was determined by
measuring Alexa488 fluorescence using flow cytometry.
Quantification of number of peptides bound per AAV2 particle. The quantification of
the number of peptides bound per AAV2 particle was determined by using the
absorption measurements. First, AAV2 particles were labeled with Cy5-NHS ester (GE
Healthcare, Piscataway, New Jersey) for 2h. After purification by a gel filtration
column, the number of attached dyes per AAV2 particle was then calculated by
measuring the absorbance of purified Cy5-labeled AAV2 particles at 650 nm for Cy5-
NHS ester (ε=250,000 M
-1
cm
-1
) and at 280 nm for AAV2 particles (ε=6.61 x 10
6
M
-1
cm
-1
) [166]. A different initial ratio of AAV2 particles to FITC-labeled peptides was
incubated for 30 min at 37 °C. After purification by a gel filtration column, the extent
of peptides on AAV2 particles was then calculated by measuring the absorbance of the
purified coupling products at 650 nm for Cy5 (ε =250,000 M
-1
cm
-1
) and at 494 nm for
FITC (ε =68,000 M
-1
cm
-1
).
In vitro model of endosomal membrane. 24-well transwells with 0.4 µm pore filters on
the bottom of the upper compartment were used to study the endosomal escape of the
complexes of AAV2 with peptides. A planar lipid bilayer was formed by applying 5 µl
83
of 1% L-α-phosphatidycholine (Sigma) on the porous filters of the upper
compartment. The complexes of AAV2 (10
7
genome copies per well) with peptides
(final concentration: 200 µM) in PBS with indicated pH values were incubated in the
upper compartment of transwell for 12 h. The vectors transferred to the lower
compartment with pH 7.4 PBS were collected, and intracellular genomes were
extracted and quantified by qPCR assay.
Cochlear tissue culture and infection. C57BL/6J mice were sacrificed at P0-P2 by
decapitation using procedures approved by the USC Animal Committee. Individual
cochlears were dissected in cold Leibovitz's L-15 (L-15) medium and explanted onto a
glass-bottom dish previously coated with CellTek (BD Biosciences). Explants were
then incubated in DMEM/F-12 supplemented with 10% FBS, N2 supplement
(Invitrogen), and penicillin at 37°C. For viral transfection experiments, the complexes
of AAV2 (10
9
particles) with CPPs (Antp, TAT-HA2 and LAH4) and AAV2 vectors
alone were applied for 4 hours directly to culture media. Tissues were placed back in
culture for 7 days to allow for stable and robust expression of the transfected
constructs. The culture media were exchanged daily. At the end of each experiment,
cultures were fixed in 4% paraformaldehyde (PFA) for 2 hours and processed for
imaging. Specimens were examined using an Olympus FV1000 confocal microscope.
Images were obtained with identical settings with 10x objective lens. To quantify GFP-
positive cells, 4 regions of interest (ROI) were randomly chosen per image at x10
magnification. Within one region, area of GFP- positive cells and area of tissue in
84
bright field were counted by software. The data are expressed as % of total area of GFP-
positive in the region of AAV-CPP-treated tissues normalized by that of tissues treated
by AAV2 alone.
Animal experiments and immunohistochemistry. AAV2 (10
9
particles/mouse) or
AAV8 (10
8
particles/mouse) was pre-incubated with Antp, TAT-HA2 or LAH4 (final
concentration 10 µM) for 30 min at 37 °C. The AAV or AAV-CPP complexes were
intramuscularly injected into BALB/c mice. Fourteen days or thirty days after
injection, mice muscles were excised, fixed, frozen, cryosectioned, and then mounted
onto glass slides. Frozen sections were then fixed and rinsed with cold PBS. After
blocking and permeabilization, the slides were washed by PBS and then incubated with
mouse anti-GFP (Invitrogen, Carlsbad, CA), rat anti-mouse CD4 (Biolegend, San
Diego, CA), or biotin anti-mouse CD8 (Biolegend, San Diego, CA), followed by
secondary antibody, and counterstained with DAPI (Invitrogen, Carlsbad, CA).
Fluorescence images were acquired by a Yokogawa spinning-disk confocal scanner
system (Solamere Technology Group, Salt Lake City, UT) using a Nikon Eclipse Ti-E
microscope. Illumination powers at 405, 491, 561, and 640 nm solid-state laser lines
were provided by an AOTF (acousto-optical tunable filter)-controlled laser-merge
system with 50mW for each laser. All images were analyzed using Nikon NIS-Elements
software. To quantify GFP-positive cells, 4 regions of interest (ROI) were randomly
chosen per image at x10 magnification. The data were expressed as % of total nuclear
area stained by GFP in the region. For toxicity, the frozen muscle sections were stained
85
with hematoxylin and eosin. Histopathologic specimens were examined by light
microscopy at ×20 magnification.
Quantification of genome copies and cytokines. DNA of vectors was extracted using the
QIAamp MinElute Virus Spin Kit (Qiagen, Valencia, CA) according to the
manufacturer's protocol. Quantitative PCR was performed on the Bio-Rad MyiQ real-
time system using a pair of primers specific for the GFP transgene: 5'-
GACATCATGAAGCCCCTTGAG-3' (forward) and 5'-
GGTGGTCGAAATTCAGATCAAC-3' (reverse).
For in vivo biodistribution of AAV2, fourteen days post-injection, organs from AAV2-
injected and AAV2-CPP-injected mice were collected, homogenized and purified
using DNeasy Blood & Tissue kit (Qiagen). qPCR was then performed for
quantification of GFP and endogenous mouse apolipoprotein B (Apob) in various
organs. A primer set of Apob was used (forward primer 5'-
CGTGGGCTCCAGCATTCTA-3', reverse primer 5'-
TCACCAGTCATTTCTGCCTTTG-3'). Apob DNA template was purchased from
Bio-Rad (Hercules, CA).
For detection of cytokines in muscles, fourteen days post-injection, total RNA was
isolated from muscles using RNeasy Mini kit (Qiagen), and first-strand cDNA was
synthesized using a QuantiTect Reverse Transcription kit (Qiagen). mRNA was
detected using primer sets of IL-1b (forward primer 5'-
86
GCAACTGTTCCTGAACTCAAC-3', reverse primer5'-
ATCTTTTGGGGTCCGTCAACT-3'), IL-6 (forward primer 5'-
TAGTCCTTCCTACCCCAATTTCC-3', reverse primer 5'-
TTGGTCCTTAGCCACTCCTTC-3') and TNFα (forward primer 5'-
CTGAACTTCGGGGTGATCGG-3', reverse primer 5'-
GGCTTGTCACTCGAATTTTGA-3'). For an internal control, a primer set of
GAPDH was used (forward primer 5'-AGGTCGGTGTGAACGGATTTG-3', reverse
primer 5'-TGTAGACCATGTAGTTGAGGT-3').
Statistical analysis. The results were expressed as means ± standard deviation. The
significance of the difference in the means was determined by Student t test.
87
Chapter 4
Codelivery of Doxorubicin and Paclitaxel by Crosslinked Multilamellar
Liposome Enables Synergistic Antitumor Activity
Yarong Liu, Jinxu Fang, Yu-Jeong Kim, Michael K. Wong, and Pin Wang
88
4.1. ABSTRACT
Combination therapy holds a significant promise in tumor chemotherapy; however,
the clinical success of combination therapy is limited by distinctive pharmacokinetics
and non-unified biodistribution of combined drugs. In this study, we report a new
robust approach to load drugs with different hydrophilicities into a same crosslinked
multilamellar liposomal vesicle (cMLV) in a precisely controllable manner over drug
ratios. The stability of cMLVs allows an improved loading efficiency and sustained
release of doxorubicin (Dox) and paclitaxel (PTX), maximizing the combination
therapeutic effect and minimizing the systemic toxicity. Furthermore, in vivo
experiments showed that the robust cMLV formulation maintains drug ratios for
prolonged times, enabling the ratio-dependent combination synergy translating from
in vitro to in vivo antitumor activity. This could be beneficial for enhancing the efficacy
of combination therapy. Therefore, this combinatorial delivery system may provide a
new strategy for synergistic delivery of multiple chemotherapeutics with a ratiometric
control over encapsulated drugs to treat cancer and other diseases.
KEYWORDS: crosslinked multilamellar liposomal vesicle, combination therapy,
doxorubicin, paclitaxel, synergy, combination ratios, nanomedicine
89
4.2. INTRODUCTION
Currently target-based drug design has been successfully used to develop numerous
drugs acting on novel molecular targets; however, they have shown poor efficacy in
clinical trials. This can be attributed to the compensatory mechanism, or drug-
mitigating response, used by the complex diseases such as cancer [71,72]. Overcoming
this compensation often requires high drug doses that can induce drug resistance at
the disease target site or side effects in other tissues [73], thus limiting the efficacy of
many potential drugs in cancer therapy. These limitations of monotherapy can be
overcome by synergistic combination of two or more agents, which allows for
reduction of dose for each compound and provides a new strategy to hit multiple
targets for treatment of complex diseases [74,75]. However, current combination
methods, through the cocktail administration, have shown limited improvement in
clinical studies due to the distinctive pharmacokinetics of individual drugs, which leads
to non-coordinate distribution after systemic administration [76,77]. Moreover, an
unexpected clinical outcome of increasing toxicity was reported with these cocktail
combinations, raising concerns about the induction of synergistic cytotoxicity by
combination therapies [78]. For instance, although a combination of doxorubicin (Dox)
and paclitaxel (PTX) has been widely used in the treatment of tumors, particularly in
metastatic breast cancer, the clinical result was not as effective as expected and an
increased cardiotoxicity was observed [79,80,81,82]. Clinical pharmacokinetic studies
also revealed an non-coordinate plasma disposition of Dox and PTX when given in
combination [83,84], rendering in vitro data ineffective in predicting in vivo
90
therapeutic efficacy of combination therapy. Thus, a more effective combination
strategy with the ability to coordinate the pharmacokinetics and biodistribution of
various drug molecules is highly desirable to maximize the combinatorial effects
without significant toxicity.
Development of nanotechnology has provided a novel combination strategy by
delivering multiple types of drugs simultaneously to the site of interest via a single
vehicle [77]. Nanoparticles are considered promising drug delivery vehicles for cancer
therapy based on their ability to prolong drug circulation time, reduce systemic
toxicity, and increase drug accumulation at tumor sites through the enhanced
permeation and retention (EPR) effect [85,86,87,88]. The pharmacokinetic behavior of
the co-formulated drugs can be determined by the pharmacokinetic behavior of the
drug carriers. Thus, nanoparticle deliver system offers the potential to coordinate the
plasma elimination and biodistribution of multiple drugs, enabling dosage
optimization to maximize cytotoxicity while minimizing the chances to develop drug
resistance. Compared to other nanoparticle deliver systems, liposomes have shown
superior ability to co-deliver multiple drugs that have vast differences in their
hydrophobicity, to the sites of action [89,90]. However, the poor stability and limited
loading efficiency of hydrophobic drugs remain the most significant concerns for
conventional formulations of liposome, limiting their clinical utility in cancer therapy
[91,92]. For example, a number of studies reported that optimal drug-to-lipid molar
ratio of paclitaxel-encapsulated conventional liposome formulation was below 4%
[93,94,95,96], thwarting the practical applications of liposomes as drug carriers.
91
Moreover, fine-controlling of the comparative loading yield and release kinetics of
multiple drugs via conventional liposomes remains an unmet need. Thus, a stable
liposomal formulation enabling improved drug loading and drug release from the
carrier in a controlled and sustained manner is necessary for combinatorial drug
delivery.
To address such a need, we have previously reported the development of crosslinked
multilamellar liposomal vesicles (cMLVs) and demonstrated their efficacy in achieving
sustained delivery of doxorubicin both in vitro and in vivo [191]. Herein, we extend the
potential of cMLVs to facilitate synergistic combinatorial delivery of hydrophobic and
hydrophilic drugs in a precisely controllable manner. Dox and PTX, as model
hydrophilic and hydrophobic drugs, respectively, were co-encapsulated into the same
cMLVs with predefined stoichiometric ratios. We show that the combination effects
(antagonistic, additive, or synergistic effect) could be determined by controlling drug
ratios of Dox and PTX in cMLVs. We also demonstrate that the drug ratio-dependent
synergistic combination effect could be achieved via the cMLV codelivery system in a
breast tumor model without significant cardiac toxicity. Moreover, cMLV particles are
capable of prolonging maintenance of the synergistic ratios of combined drugs in vivo
and, in turn, providing a significantly enhanced antitumor efficacy compared to free-
drug cocktail administration. The results demonstrate the great potential of cMLVs as
combinatorial drug delivery vesicles to induce synergy of antitumor therapeutics both
in vitro and in vivo, thus setting a new paradigm in nanomedicine for combination
therapies.
92
4.3. MATERIALS AND METHODS
Cell lines, antibodies, reagents, and mice
B16-F10 (ATCC number: CRL-6475) and 4T1 tumor cells (ATCC number: CRL-2539)
were maintained in a 5% CO 2 environment with Dulbecco’s modified Eagle medium
(Mediatech, Inc., Manassas, VA) supplemented with 10% FBS (Sigma-Aldrich, St.
Louis, MO) and 2 mM of L-glutamine (Hyclone Laboratories, Inc., Omaha, NE).
Mouse anti-β-Actin and rabbit antibody against phospho-specific protein p44/42
MAPK (Erk 1/2) were purchased from Cell Signaling Technology (Danvers, MA).
Goat anti-Rabbit IR dye®680RD and Goat anti-Mouse IR Dye®800CW were obtained
from LI-COR BioSciences (Lincoln, Nebraska). Doxorubicin, Paclitaxel, Daunorubicin
and Doxetaxel were purchased from Sigma-Aldrich (St. Louis, MO).
All lipids were obtained from NOF Corporation (Japan): 1,2-dioleoyl-sn-glycero-3-
phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phospho-(10-rac-glycerol)
(DOPG), and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[4-(p-
maleimidophenyl) but- yramide (maleimide-headgroup lipid, MPB-PE).
Female 6–10 weeks-old BALB/c mice were purchased from Charles River Breeding
Laboratories (Wilmington, MA). All mice were held under specific pathogen-reduced
conditions in the Animal Facility of the University of Southern California (Los
Angeles, CA, USA). All experiments were performed in accordance with the guidelines
set by the National Institute of Health and the University of Southern California on the
Care and Use of Animals.
93
Synthesis of cMLVs
Liposomes were prepared based on the conventional dehydration-rehydration method.
All lipids were obtained from NOF Corporation (Japan). DOPC, DOPG and MPB-PE
were combined in chloroform, at a molar lipid ratio of DOPC:DOPG:MPB = 4:1:5, and
the organic solvent in the lipid mixture was evaporated under argon gas. The lipid
mixture was further dried under vacuum overnight to form dried thin lipid films. To
prepare cMLV (Dox+PTX), paclitaxel in organic solvent was mixed with the lipid
mixture to form dried thin lipid films. The resultant dried film was hydrated in 10 mM
Bis-Tris propane at pH 7.0 with doxorubicin by vigorous vortexing every 10 min for 1
h, and then applied with 4 cycles of 15-s sonication (Misonix Microson XL2000,
Farmingdale, NY) on ice in 1 min intervals for each cycle. To induce divalent-triggered
vesicle fusion, MgCl 2 was added at a final concentration of 10 mM. The resulting
multilamellar vesicles were further crosslinked by addition of Dithiothreitol (DTT,
Sigma-Aldrich) at a final concentration of 1.5 mM for 1 h at 37°C. The resulting
vesicles were collected by centrifugation at 14,000 g for 4 min and then washed twice
with PBS. For pegylation of cMLVs, the particles were incubated with 1 µmol of 2 kDa
PEG-SH (Laysan Bio Inc. Arab, AL) for 1 h at 37°C. The particles were then
centrifuged and washed twice with PBS. The final products were stored in PBS at 4°C.
Characterization of physical properties
The hydrodynamic size and size distribution of cMLVs were measured by dynamic
light scattering (Wyatt Technology, Santa Barbara, CA).
94
In vitro drug encapsulation and release
To study the loading capacity of Dox, cMLV (Dox) and cMLV (Dox+PTX) were
collected and washed twice with PBS, followed by lipid extraction of vesicles with 1%
Triton X-100 treatment. Dox fluorescence (excitation 480 nm, emission 590 nm) was
then measured by Shimadzu RF-5301PC spectrofluorometer (Japan). The amount of
paclitaxel incorporated in the cMLV(PTX) and cMLV(Dox+PTX) was determined by
C-18 RP-HPLC chromatography (Beckman Coulter, Brea, CA). The cMLV(PTX) and
cMLV(Dox+PTX) suspensions were diluted by adding water and acetonitrile to a total
volume of 0.5 ml. Extraction of paclitaxel was accomplished by adding 5 ml of tert-
butyl methyl ether and votex-mixing the sample for 1 min. The mixtures were then
centrifuged and the organic layer was transferred into a glass tube and evaporated to
dryness under argon. Buffer A (95% water, 5% acetonitrile) was used to rehydrate the
glass tube. To test PTX concentration, 1 ml of the solution was injected into a C18
column, and the paclitaxel was detected at 227 nm (flow rate 1 ml/min). To obtain the
release kinetics of Dox and PTX from liposomes, the releasing media was removed
from cMLVs incubated in 10% fetal bovine serum (FBS)-containing media at 37°C and
replaced with fresh media daily. The removed media was quantified for Dox
fluorescence (by spectrofluorometer) and PTX fluorescence (by HPLC) every day.
In vitro drug loading efficiency
Loading efficiency was determined by the ratio of encapsulated drug to total
phospholipid mass. Phospholipid phosphate assay was carried out to calculate the
95
phospholipid mass. cMLVs were centrifuged, and 100 µl chloroform was added to the
pellets to break down the lipid bilayers. The samples were transferred to glass tubes
and evaporated to dryness. After adding 100 µl perchloric acid, the samples were
boiled at 190 °C for 25 min. Samples will turn brown then clear as the lipids are
digested. Samples were cooled to room temperature and diluted to 1 ml with distilled
water. The amount of phospholipid phosphate was determined by the malachite green
phosphate detection kit (R&D systems, Minneapolis, MN).
In vitro cytotoxicity and data analysis
B16-F10 and 4T1 cells were plated at a density of 5 × 10
3
cells per well in 10% fetal
bovine serum (FBS)-containing media in 96-well plates and grown for 6 h. The cells
were then exposed to a series of concentrations of cMLV (single drug) or cMLV (drug
combinations), at different weight ratios of combined drugs, for 48 h. The cell viability
was assessed using the Cell Proliferation Kit II (XTT assay) from Roche Applied
Science according to the manufacturer's instructions. Cell viability percentage was
determined by subtracting absorbance values obtained from media-only wells from
drug-treated wells and then normalizing to the control cells without drugs. The
fraction of cells affected (f a) at each drug concentration was subsequently determined
for each well. The data was analyzed by nonlinear regression to get the IC 50 value. The
Combination Index (CI) values were calculated by the equation: CI=C A,X/IC X,A +
C B,X/IC X,B [192]. Using this analysis method, a CI=0.9-1.1 reflects additive activity, a CI
>1.1 indicates antagonism, while a CI <0.9 suggests synergy.
96
Western blot analysis
Cells were collected 24 h after treatment and lysed in lysis buffer supplemented with
protease inhibitors, incubated on ice for 15 min, and then cleared by centrifugation at
10000×g at 4°C for 10 min. Protein concentration was determined using Micro BCA
Protein Assay Kit (Thermo Scientific). Lysates (20 µg) were separated by reducing 12%
polyacrylamide gel and then transferred to polyvinylidene difluoride membranes.
Immunodetection of ERK was carried out with antibodies specific to rabbit phospho-
specific protein p44/42 MAPK (Erk 1/2) and goat anti-rabbit IR dye®680RD.
Immunodetection of β-actin was carried out with antibodies against β-actin and goat
anti-mouse IR Dye®800CW. Membranes were developed using Odyssey Infrared
Fluorescent Imager (LI-COR BioSciences, Lincoln, Nebraska).
3.8. Determination of doxorubicin and paclitaxel levels in tumor
BALB/c female mice (6–10 weeks-old) were inoculated subcutaneously with 0.2 × 10
6
4T1 tumor cells. The tumors were allowed to grow for 20 days to a volume of ~500
mm
3
before treatment. On day 20, the mice were injected intravenously through the
tail vein with 8.33mg/kg Dox+1.66mg/kgPTX, 5mg/kg Dox+ 5mg/kg PTX, 1.66 mg/kg
Dox + 8.33mg/kg PTX either in solution or in cMLVs. Three days after injection,
tumors were excised and frozen at -20 °C. Docetaxel (10 µl, 100 µg/ml) as an internal
standard (IS) for paclitaxel, or 10 µl of Daunorubicin (100 µg/ml) as an internal
standard for Doxorubicin were added to the weighted tumor tissues. In order to
97
extract paclitaxel and the internal standard (Docetaxel), tumor tissue was
homogenized in 1 ml ethyl acetate and then centrifuged at 5000 rpm for 10 min. In
order to extract doxorubicin and its internal standard (Daunorubicin), tumor tissue
was homogenized in 1 ml methanol and then centrifuged at 5000 rpm for 10 min.
Then the organic layer was transferred to a clean glass tube and evaporated to dryness
under a stream of argon. Buffer A (95% water, 5% acetonitrile) was used to rehydrate
the sample in the glass tube. 1 ml of the solution was injected into C18 column, and the
paclitaxel was detected at 227 nm (flow rate 1 ml/min), and doxorubicin was detected
at 482 nm (flow rate 1 ml/min). Stock solutions of Dox and PTX (100, 10, and 1 µg/ml)
and IS were prepared as calibration samples. Then 500 µl of tumor homogenates were
spiked with 500 µl calibration samples with the internal standard at fixed
concentration of 1 µg/ml. Calibration curves of doxorubicin and paclitaxel were
constructed using the ratio of peak height of doxorubicin or paclitaxel and internal
standard by weighted (1/y) linear regression analysis.
In vivo antitumor activity study
BALB/c female mice (6–10 weeks-old) were inoculated subcutaneously with 0.2 × 10
6
4T1 breast tumor cells. The tumors were allowed to grow for 8 days to a volume of ~50
mm
3
before treatment. On day 8, the mice were injected intravenously through the tail
vein with 3.33 mg/kg Dox + 0.67mg/kg PTX, 2mg/kg Dox + 2mg/kg PTX, 0.67mg/kg
Dox+ 3.33mg/kg PTX, either in cMLVs or in solution every three days (six mice per
group). Tumor growth and body weight were monitored until the end of an
98
experiment. The length and width of the tumor masses were measured with a fine
caliper every three days after injection. Tumor volume was expressed as 1/2 × (length ×
width
2
). Survival end point was set when the tumor volume reached 1000 mm
3
. The
survival rates are presented as Kaplan-Meier curves. The survival curves of individual
groups were compared by a log-rank test.
Immunohistochemistry of tumors, cardiac toxicity and confocal imaging
BALB/c female mice (6–10 weeks-old) were inoculated subcutaneously with 0.2 × 10
6
4T1 tumor cells. The tumors were allowed to grow for 20 days to a volume of ~500
mm
3
before treatment. On day 20, the mice were injected intravenously through tail
vein with 8.33mg/kg Dox+1.66mg/kgPTX, 5mg/kg Dox+ 5mg/kg PTX, 1.66 mg/kg
Dox + 8.33mg/kg PTX in solution or cMLVs. Three days after injection, tumors were
excised, fixed, frozen, cryo-sectioned, and mounted onto glass slides. Frozen sections
were fixed, and rinsed with cold PBS. After blocking and permealization, the slides
were washed by PBS and incubated with TUNEL reaction mixture (Roche,
Indianapolis, Indiana) for 1 h and counterstained with DAPI (Invitrogen, Carlsbad,
CA). Fluorescence images were acquired by a Yokogawa spinning-disk confocal
scanner system (Solamere Technology Group, Salt Lake City, UT) using a Nikon
Eclipse Ti-E microscope. Illumination powers at 405, 491, 561, and 640 nm solid-state
laser lines were provided by an AOTF (acousto-optical tunable filter)-controlled laser-
merge system with 50mW for each laser. All images were analyzed using Nikon NIS-
99
Elements software. For quantifying TUNEL positive cells, 4 regions of interest (ROI)
were randomly chosen per image at ×2 magnification. Within one region, area of
TUNEL-positive nuclei and area of nuclear staining were counted by Nikon NIS-
Element software, with data expressed as % total nuclear area stained by TUNEL in the
region.
For cardiac toxicity, heart tissues were harvested 3 days after injection, and were fixed
in 4% formaldehyde. The tissues were frozen and then cut into sections and mounted
onto glass slides. The frozen sections were stained with hematoxylin and eosin.
Histopathologic specimens were examined by light microscopy.
Statistics
The differences between two groups were determined with Student’s t test. The
differences among three or more groups were determined with a one-way ANOVA.
4.4. RESULTS AND DISCUSSIONS
4.4.1 Characteristics of combinatorial drug delivery via cMLVs
Our strategy of combination drug delivery via crosslinked multilayer liposomal
vesicles was to incorporate hydrophobic drug paclitaxel (PTX) into the lipid
membranes and encapsulate hydrophilic drug doxorubicin (Dox) in the aqueous core
of liposomal vesicles. The crosslinked multilamellar liposomal vesicles (cMLVs) were
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formed by adding MgCl 2 to trigger vesicle fusion, and then stabilized by dithiothreitol
(DTT) to form crosslinkers between adjacent liposomal vesicles [191,193]. The surface
of crosslinked multilayer liposomes was further PEGylated with thiol-termineated PEG,
which is known to enhance the vesicle stability and elongate blood circulation half-life
[194,195]. First, we characterized the physical properties of dual drug-loaded cMLVs
compared to single drug-loaded cMLVs to determine whether drug combinations
could change the physical properties of liposomal formulation. Dynamic light
scattering (DLS) measurements showed that the resulting dual drug-loaded cMLVs
had a similar average hydrodynamic diameter as single drug-loaded cMLVs (Figure
26A-C). We further found that there was no significant aggregation of particles during
the crosslinking process in these three liposomal formulations, as a narrow size
distribution with similar polydispersity was observed in both dual drug-loaded and
single drug-loaded cMLVs. This suggests that the combination of Dox and PTX in a
single nanoparticle has a negligible effect on the formation of cMLV particles.
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Figure 26. Characteristics of cMLV (Dox+PTX). (A-C). The hydrodynamic size distribution of
cMLV(Dox), cMLV(PTX), and cMLV(Dox+PTX) measured by dynamic light scattering. The
mean hydrodynamic diameter (HD) and polydispersity index (PI) of cMLV(Dox),
cMLV(PTX), and cMLV(Dox+PTX) were indicated in the graph. (D, E) Effects of co-
encapsulation of Dox and PTX on loading capability and drug release kinetics profiles of
cMLVs. The encapsulation efficiency (D) and loading efficacy (E) of drugs in cMLV(combined
drugs) and cMLV (single drug). (F-H) In vitro release kinetics of doxorubicin and paclitaxel
from dual-drug loaded cMLVs and single-drug loaded cMLVs. Error bars represent the
standard deviation of the mean from triplicate experiments.
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We next determined whether the encapsulation efficiency and loading yield of cMLVs
were affected by loading multiple therapeutics. Single drug-loaded and dual drug-
loaded cMLVs were dissolved in organic solvents to free all the encapsulated drugs
(Dox and/or PTX), which were quantified by spectrofluorometer and/or HPLC,
respectively. As shown in Figure 26D, the drug encapsulation efficiency of Dox and
PTX in cMLV (Dox+PTX) was not significantly different from that in either cMLV
(Dox) or cMLV (PTX). It was also showed that cMLV (Dox+PTX) had a comparable
drug loading yield (~270 mg drug per g of phospholipids) to single drug-loaded
cMLVs (Figure 26E). Furthermore, the drug release profiles of Dox and PTX were
evaluated in dual drug-loaded cMLVs to investigate whether the cMLVs are able to
release the individual drugs in a controlled manner. The results of in vitro drug release
assay showed that cMLV (Dox+PTX) had slow and linearly sustained release kinetics
of both Dox and PTX (up to 2 weeks), similar to that of single drug-loaded cMLVs
(Figure 26F-H). These results confirm that this approach enables the loading of drugs
with different hydrophobicity into the same nanoparticles with efficient drug loading
yield and sustained drug release profiles.
4.4.2 In vitro analysis of doxorubicin: paclitaxel for drug ratio-dependent synergy
Combinatorial drug delivery is known to induce synergistic effect; however, it has been
reported that the combination effect, synergy, additivity or antagonism, was
determined by combination ratios [89,196]. To test the hypothesis, the cytotoxicity of
cMLV (Dox+PTX) with three combination ratios (5:1, 1:1 and 1:5) against B16 and
4T1 cell lines was examined in comparison with the same three ratio combinations in
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cocktail solution. Figure 27A summarizes the results of IC 50 measurements of the dual
drug-loaded cMLVs with three combination ratios for 48 h of incubation with B16 and
4T1 cells. It was found that IC 50 value of cMLV (Dox+PTX) at Dox:PTX weight ratios
of 1:1 and 5:1 in B16 and 4T1 cells was significantly smaller than that of 1:5 combined
ratios. A similar trend of IC 50 was observed in free Dox and PTX combinations with
different ratios (Figure 27B).
Figure 27. Determination of the ratio of drug combinations to induce synergy. (A, B) In vitro
cytotoxicity of three combination ratios (5:1, 1:1 and 1:5) of Dox and PTX in cMLV
formulations (A) or solution (B) in B16 melanoma tumor or 4T1 breast tumor cell lines. The
cytotoxicity was measured by a standard XTT assay. (C) Combination Index (CI) histogram
for cMLV (different drug combinations) exposed to cultured B16 and 4T1tumor cells. (D)
Combination Index histogram for different ratios of drug combination in solution exposed to
culture B16 and 4T1tumor cells. The surviving cell fraction from three replicates was averaged
and analyzed by nonlinear regression. The histogram presents the CI values obtained at a
fraction of 0.5. Error bars represent the standard deviation of the mean from triplicate
experiments. (E) Immunoblot analysis of phosphorylated ERK in B16 cells treated by
cMLV(Dox+PTX) with three combination ratios: 5:1, 1:1 and 1:5. β-actin was used as control.
(F) Quantification of phosphorylated ERK shown in (E). Protein amounts were estimated by
densitometry of immunoblots. Error bars represent SD.
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Figure 28. IC50 values of Dox and PTX in cMLV formulation or free drug solution in B16
melanoma or 4T1 breast tumor cells.
Moreover, combination index (CI) values were analyzed from in vitro cytotoxicity
curves for doxorubicin and paclitaxel combinations either in cMLVs or cocktail
solutions to assess the effects of combination. The IC 50 values of individual drug either
in cMLVs or in solution are shown in Figure 28. A CI of less than, equal to, and greater
than 1 is known to indicate synergy, additivity, and antagonism, respectively
[89,192,197,198]. Although combination indexes were only shown at a fraction of
affected cells (fa) of 0.5 (50% cell growth inhibition relative to control cells) in Figure
27, the profile of synergy/antagonism was similar for other fa values. As shown in
Figure 27C, at fa=0.5, synergistic effects were observed in both B16 and 4T1 tumor
cells at Dox:PTX weight ratios of 5:1 and 1:1 (Dox:PTX) when they were coloaded into
cMLVs, while the combination at a 1:5 ratio was additive or antagonistic in B16 and
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4T1 cells. In contrast, no synergistic effect was observed in both B16 and 4T1 cells
treated with three ratios of Dox and PTX in cocktails, as shown in Figure 27D, further
confirming the potential of cMLVs to induce synergy by controlling combination
ratios.
Our data indicated that combinatorial delivery via cMLVs with high ratio of PTX
induced additivity or antagonism. In fact, some studies have shown that low
concentration of PTX can induce cell apoptosis more effectively than high
concentration, but the mechanism remains elusive [199,200]. Further studies suggested
that PTX could activate the extracellular signal regulated kinase (ERK), leading to cell
proliferation and building drug resistance [201,202,203]. It was also shown that
inhibiting ERK pathway dramatically enhanced cell apoptosis induced by PTX
[201,203]. These studies indicate that potential factor to antagonism/additivity of drug
combinations are due to the activation of ERK by high PTX concentration in cancer
cells. To investigate whether there is an difference in activation of ERK on melanoma
cells treated by cMLV(Dox+PTX) with three combination ratios, phosphorylated ERK
expression was detected by western blot. As shown in Figure 2E, the combination of
Dox and PTX at a 1:5 ratio showed a significant expression of phosphorylation of ERK
compared to the combinations at a ratio of 1:1 and 5:1. The quantification data (Figure
27F) also demonstrated a 30-fold enhancement in phosphorylation of ERK in cells
treated by cMLV(Dox+PTX) with a combination ratio of 1:5. These data suggested
that ratio-dependent combination effects are likely linked to the ERK activation caused
by high concentration of PTX.
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4.4.3 Drug ratio-dependent efficacy of cMLV(Dox+PTX) in tumor treatment
In order to assess whether the drug ratio-dependent in vitro cytotoxicity was also
manifested in vivo, doxorubicin and paclitaxel were co-encapsulated in cMLV particles
at a weight ratios ranging from 5:1 to 1:5, while keeping the total drug mass
encapsulated in cMLVs constant. This panel of fixed ratio cMLV formulations and the
same fixed ratio combination in cocktail solutions were evaluated for their antitumor
efficacy in 4T1 breast tumor models. As shown in Figure 29A, tumor volume in the
groups treated with drug combinations in solution ranging from 5:1 to 1:5 decreased
significantly compared to that in the control group (p < 0.01). Tumor volume between
the groups treated with different ratios of drug combinations in solution did not show
a significant difference (p > 0.05), consistent with the in vitro finding that free drug
combinations did not show synergistic effect. In comparison, administration of the 5:1
and 1:1 weight ratio of Dox to PTX in cMLV resulted in significantly enhanced
antitumor activity compared to the 1:5 ratio, indicating the ability of cMLVs to induce
the ratio-dependent synergistic effect in vivo. Moreover, no weight loss was observed
for all treated groups during the whole experiment shown in Figure 3B, indicating that
there is no significant toxicity from these dose combinations.
The dose-dependent antitumor activity was further confirmed by survival test as
shown in Figure 29C. Treatment with three ratios of drug combinations in cocktail
solution resulted in an increased survival time (35 days) compared to PBS treatment
(28 days, p < 0.05). Administration of the 5:1 and 1:1 weight ratio in cMLV
formulations resulted in a significant increased life span compared to 1:5 ratio in
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cMLVs (p < 0.05). These results confirmed a dose-dependent synergy of drug
combinations in cMLV formulations and provide a positive correlation linking the
combination effects in vitro to the degree of antitumor efficacy in vivo.
Figure 29. Drug ratio-dependent efficacy of cMLV(Dox+PTX) in tumor treatment. (A) Tumor
growth was measured after treatment with PBS, 3.33 mg/kg Dox + 0.67mg/kg PTX, 2mg/kg
Dox + 2mg/kg PTX, 0.67mg/kg Dox + 3.33mg/kg PTX, either in cMLVs or in solution every
three days. Tumor growth and body weights were monitored until the end of the experiment.
Error bars represent standard error of the mean, n = 6 for each treatment group (*p<0.05,
**p<0.01). (B) Average mouse weight loss over the duration of the experiment. (C) Survival
curves for 4T1 bearing mice treated with PBS, 3.33 mg/kg Dox + 0.67mg/kg PTX, 2mg/kg Dox
+ 2mg/kg PTX, 0.67mg/kg Dox+ 3.33mg/kg PTX either in cMLVs or in solution every three
days. The survival rates are presented as Kaplan-Meier curves. The survival curves of
individual groups were compared by a log-rank test.
4.4.4 Drug ratio-dependent efficacy of co-encapsulated Dox:PTX on tumor apoptosis
To investigate the ratio-dependent antitumor mechanism in vivo, TUNEL assay was
performed to detect apoptotic cells in 4T1 tumors treated with different ratios of Dox
and PTX in cocktail and in cMLV formulations for 3 days. As shown in Figure 30A,
4T1 tumors treated with three different ratios (5:1, 1:1 and 1:5) of Dox and PTX in
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solution induced cell apoptosis by a significant amount compared to controls. The
apoptosis index was not remarkably different among different ratios of drug
combination cocktails (p > 0.05), consistent with the similar effect on tumor growth
between the cocktail treatments. Moreover, the 5:1 and 1:1 ratio of Dox and PTX in
cMLVs promoted tumor cell apoptosis compared to the antagonistic ratio (1:5). The
quantified data (Figure 30B) further confirms that drug ratio-dependent antitumor
efficacy via cMLVs can contribute to the different levels of tumor apoptosis.
4.4.5 In vivo cardiac toxicity evaluation of drug combinations in cMLV formulations
An unexpected clinical outcome of increased cardiotoxicity after combined treatments
of Dox and PTX has been reported, thus limiting their clinical applications [113,204].
To investigate whether the synergistic therapies could induce synergistic cardiac
toxicity, three combination ratios of doxorubicin and paclitaxel in both cMLV
formulations and cocktail solutions were evaluated. Mice bearing 4T1 tumors were
injected intravenously through tail vein with 8.33mg/kg Dox + 1.66mg/kg PTX,
5mg/kg Dox + 5mg/kg PTX, 1.66 mg/kg Dox + 8.33mg/kg PTX in solution or in
cMLVs. Hematoxylin and eosin staining of cardiac tissue sections from each treatment
group were examined. As shown in Figure 31, all three combination ratios of Dox and
PTX in cocktail solutions caused damage of cardiac tissue indicated by myofibrillary
loss, disarray, and cytoplasmic vacuolization. No significant histopathologic changes
in cardiac tissue were observed in three combination ratios of Dox and PTX in cMLV
formulations compared to that in control group, indicating that a reduction in
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systemic toxicity can be achieved when drugs are coencapsulated in cMLVs. Moreover,
no synergistic toxicity was observed in the synergistic ratios (5:1 and 1:1) of Dox and
PTX in cMLVs.
Figure 30. Drug ratio-dependent efficacy of co-encapsulated Dox:PTX on tumor cell apoptosis.
(A) 4T1 Tumor bearing mice were treated with PBS, 8.333mg/kg Dox + 1.667mg/kg PTX,
5mg/kg Dox + 5mg/kg PTX, 1.667mg/kg Dox+ 8.33mg/kg PTX, either in cMLVs or in
solution. Three days after injection, tumors were excised. Apoptotic cells were detected by
TUNEL assay (green), and co-stained by nuclear staining DAPI (blue). Scale bar represents 50
µm. (B) Quantification of apoptotic positive cells in 4T1 tumor. To quantify TUNEL positive
cells, 4 regions of interest (ROI) were randomly chosen per image at ×2 magnification. Within
one region, area of TUNEL positive nuclei, and area of nuclear staining were counted by
software. The data is expressed as % total nuclear area stained by TUNEL in the region. Data
represented as mean ± SD (n=3).
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Figure 31. In vivo toxicity. Histologic appearance of cardiac tissues obtained from C57/BL6
mice with no drug treatment or administered a single intravenous injection with three
combination ratios of Dox and PTX (5:1, 1:1 and 1:5) in solutions or cMLV formulations at 10
mg/kg total drug equivalent.
4.4.6 In vivo maintenance of drug ratios in cMLV formulations
In order to determine if combination ratios of drugs delivered via cMLVs were well
maintained in vivo, and to correlate the in vivo effects to the in vitro combination effect,
the drug concentrations in tumor tissues were measured. Doxorubicin and paclitaxel
were co-encapsulated at the 5:1, 1:1, and 1:5 weight ratios inside cMLVs, and
administered i.v. to mice, while the same ratios of drug combinations in cocktail
solutions were administrated as controls. Twenty-four hours after injection, tumors
were excised and homogenized, and Dox and PTX were extracted and detected by
HPLC analysis, as illustrated in Figure 32A. The HPLC results show that cMLVs
maintain the doxorubicin:paclitaxel weight ratio at 5:1, 1:1 and 1:5, respectively, in
tumors for over 24 h (Figure 32B). In comparison, the Dox:PTX weight ratio changed
dramatically after administration of free-drug cocktails, shown in Figure 32C. In
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addition, remarkedly more doxorubicin and paclitaxel were accumulated in tumors
when administered via cMLV formulations compared to free-drug cocktails with
equivalent amounts of Dox and PTX, thus maximizing their combinatorial effect.
These results indicate that cMLVs can efficiently maintain the combination ratios in
vivo, thus translating the combination effects (synergy, additivity and antagonism)
from in vitro to in vivo.
Figure 32. In vivo maintenance of Dox:PTX ratios in cMLV formulations. (A, B) Tumor
bearing mice were treated with PBS, 8.333mg/kg Dox + 1.667mg/kg PTX, 5mg/kg Dox +
5mg/kg PTX, 1.667mg/kg Dox + 8.33mg/kg PTX, either in cMLVs (A) or in solution (B). 24 h
after injection, tumors were excised and drug concentrations of Dox and PTX were measured
by HPLC. All data are shown as the means of triplicate experiments.
To summarize, a robust approach for combinatorial chemotherapy was presented by
encapsulating two different types of antitumor therapeutics, with ratiometric control
over drug loading, into a crosslinked multilamellar liposomal formulation. Previously,
we have demonstrated the superior ability of cMLVs as drug carriers to offer
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controllable and sustainable drug release profiles of doxorubicin with increased vesicle
stability, enabling an improved antitumor activity. In the present study, we further
explore the potential of cMLVs in combinatorial delivery of Dox and PTX, which have
been widely used as a combined anthracycline-taxane regimen in metastatic breast
cancer [205], to achieve synergistic antitumor activity. A number of studies suggested a
non-coordinate biodistribution profiles of this combination when administered in
cocktail solutions, limiting the efficacy of combination in tumor treatment [83,84].
However, the versatile crosslinked multilamellar liposomes enabled codelivery of Dox
and PTX via a single vesicle to the cancer site, thus coordinating the plasma
elimination and tissue distribution of the combined drugs.
Recent studies revealed that the activity of antitumor drug combinations is determined
by the ratio of the combined drugs exposed to cells [196,206,207,208]. Therefore, it is
highly desirable to maintain the synergistic ratio of combined drugs in vivo to optimize
the antitumor activity. Here, we demonstrate that stability of cMLVs enables us to
coload Dox and PTX with predefined ratios and induce a ratio-dependent synergy in
tumor cells. It was previously reported by a number of studies that only 3%-4% drug-
to-lipid molar ratio could be achieved in paclitaxel-containing liposomes to maintain
the stability of liposomes. For instance, it was reported that at higher than 8% PTX-to-
lipid ratio, liposome formulations (PG: PC 3:7 molar ratio) were stable for less one day
[94]. It is noteworthy that cMLV could achieve up to 30% paclitaxel-to-lipid molar
ratio while maintaining high stability. This is most likely due to the crosslinked
multilamellar structure of cMLVs, allowing co-delivery of Dox and PTX with high
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loading efficiency. In addition, enhanced vesicle stability of cMLVs enables these
nanoparticles to maintain the combination ratios of Dox and PTX at tumor sites,
translating the ratio-dependent synergy from in vitro to in vivo. This would be
beneficial for predicting the efficacy of treatment in clinical trials and the optimal
design of combination therapy based on in vitro cellular experiment. Our in vivo
results also reveals that the enhanced combinatorial efficacy of cMLVs compared to
cocktail combination is due to the augmented accumulation of drugs at tumor sties,
maximizing the combination effects.
An increased cardiac toxicity of Dox and PTX combination in cocktail was reported in
clinical studies [113,204], raising the concern that a significant side effect is associated
with the synergistic therapeutic efficacy. However, we previously demonstrated that
the robust cMLV formulation greatly reduced systemic toxicity of Dox, most likely due
to the sustained drug release profile of Dox. Here, we further demonstrate the potential
of cMLVs in combinatorial delivery by inducing the synergistic effect of combinations
without causing cardiac toxicity. These results, taken together, indicated that the
superior ability of cMLVs in combination therapy is not only attributed to the
prolonged exposure of drugs to tumor cells, but also to the maintenance of synergistic
combination ratios at the site of action with no significant systemic toxicity.
4.5. CONCLUSION
In conclusion, we have demonstrated that the ratio-dependent synergy of drug
combinations shown in vitro can be translated into the synergistic antitumor efficacy
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in vivo by coloading two types of drugs into crosslinked multilamellar liposomal
formulations. Unlike the free-drug cocktail, cMLVs maintain combination ratios for
prolonged times after administration in vivo due to the ability of cMLVs to co-
encapsulate and retain the combined drugs in a manner that coordinates their
pharmacokinetics. In the present study two drugs (Dox and PTX) were chosen to
demonstrate the advantage of this combination drug delivery system by cMLVs. In this
regard, we believe this delivery system can offer the clinical possibility for an improved
synergistic delivery of multiple chemotherapeutics with a ratiometric control over drug
encapsulation for combination cancer treatment.
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Chapter 5
Co-delivery of Doxorubicin and Paclitaxel via Crosslinked Liposomal Formulations
To Overcome Multidrug Resistance in Tumor
Yarong Liu, Jinxu Fang, and Pin Wang
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5.1 ABSTRACT
Multidrug resistance (MDR) is a significant challenge to the effective cancer
chemotherapy treatment. Through the development of a drug delivery system that
allows for sustained release of combined drugs with improved vesicle stability, we have
demonstrated that multidrug resistance in cancer cells can be overcome by codelivery
of doxorubicin (Dox) and paclitaxel (PTX) via a crosslinked multilamellar vesicle
(cMLV). This combinatorial delivery system (cMLV(Dox+PTX)) achieves enhanced
drug accumulation and retention, resulting in improved cytotoxicity in tumor cells,
including drug resistant cells. Moreover, cMLV(Dox+PTX) significantly overcomes
MDR by reducing the expression of P-glycoprotein (P-gp) in cancer cells, thus
improving antitumor activity in vivo. With a combined therapeutic ability that
enhances drug delivery efficacy to tumors and lowers the apoptotic threshold of
individual drugs, cMLV(Dox+PTX) represents a potential multimodal therapeutic
strategy to overcome MDR in cancer therapy.
Keywords: multidrug resistance, combinatorial delivery, doxorubicin, paclitaxel, P-
glycoprotein, crosslinked multilamellar vesicle, cancer therapy
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5.2 INTRODUCTION
The development of multidrug resistance (MDR) against a variety of conventional and
novel chemotherapeutic agents has been a major impediment to the success of cancer
therapy [97,98]. One of the most important mechanisms involved in MDR is P-
glycoprotein (P-gp), an active drug efflux transporter, overexpression in the plasma
membrane of various cancer cells. P-gp is capable of effluxing a broad range of
anticancer agents such as taxanes and anthracyclines [99,100]. For example, the
efficacy of doxorubicin (Dox) and paclitaxel (PTX), two of the most widely used agents
for the treatment of various cancers, is often compromised by P-gp mediated MDR
[101,102]. Therefore, a strategy of P-gp expression inhibition and bypass of P-gp-
mediated drug efflux have been developed to overcome MDR. For instance, a large
number of P-gp inhibitors and siRNAs targeting the gene encoding P-gp have been
delivered in combination with anticancer agents to downregulate P-gp expression,
enabling drugs to reach sufficient concentrations to induce cytotoxicity [103,104,105].
However, P-gp inhibitors (functional inhibitors or siRNA) yielded disappointing
clinical trial result due to high systemic toxicities and enhanced side effects of
chemotherapy in normal cells [106,107].
Combination therapy with multiple chemotherapeutics provides a promising strategy
to suppress MDR. Different drugs may attack cancer cells at varying stages of their
growth cycles, thus decreasing the concentration threshold for individual drugs that is
required for cytotoxicity [108]. It has been reported that various drug combinations
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successfully induced synergistic antitumor activities and prevented disease recurrence
[109,110]. For example, co-administration of Dox and PTX in cocktail, is considered
standard anthracycline-taxane combination treatment for various tumors, as they are
able to overcome drug resistance [111,112,113]. However, one major challenge of
combinatorial therapy is coordinating the pharmacokinetics and cellular uptake of
combined therapeutics, which has limited the clinical success of combination therapy
[83,114].
To overcome this challenge, novel strategies that allow loading of multiple therapeutics
into a single drug-delivery vehicle for concurrent delivery at the site of action have
been extensively explored [115,116]. Several drug-delivery systems have shown the
ability to co-deliver multiple drugs to tumor sites and improve antitumor activities,
potentially overcoming drug resistance, shown by a reduction in the dosage of
individual drugs [117,118]. Indeed, nanoparticle delivery systems are known to
efficiently deliver therapeutics to the tumor sites through the enhanced permeability
and retention (EPR) effect, enhancing the concentration of therapeutics in tumors
[85,86,88]. Moreover, these nanoparticles can enter cancer cells by an endocytosis
pathway, which is independent from the P-gp pathway, thereby enhancing the cell
entry of therapeutics [119,120,121]. Thus, a nanoparticle delivery system which shows
high efficiency of cellular entry and subsequently triggers intracellular release of
multiple anticancer drugs to overcome MDR is highly desirable.
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Liposomes are one of the most popular nanoparticle delivery system for combinatorial
delivery of multiple drugs due to their ability to coload efficiently hydrophilic and
hydrophobic drugs [86,194]. Previously, we have developed a robust crosslinked
multilamellar liposomal vesicle (cMLV), with enhanced vesicle stability, to efficiently
codeliver hydrophilic (doxorubicin) and hydrophobic (paclitaxel) and induce a ratio-
dependent synergistic antitumor activity both in vitro and in vivo (Yarong). Moreover,
it was shown that the cMLVs are mainly internalized by cells through a caveolin-
dependent endocytosis and are trafficked through the endosome-lysosome network for
release of drugs [191]. In this study, we have examined the potential of cMLVs as
combinatorial delivery systems to overcome P-gp-mediated drug resistance both in
vitro and in vivo. We have demonstrated that combination of Dox and PTX, when
administered in cMLV formulations, shows significant enhancement of cytotoxicity
and antitumor activities. cMLV formulations contribute to these antitumor activities
by enhancing systemic delivery efficiency and lowering tumor apoptotic threshold with
their combination drugs.
5.3 MATERIALS AND METHODS
Mice. Female 6–10 weeks-old BALB/c mice were purchased from Charles River
Breeding Laboratories (Wilmington, MA). All mice were held under specific
pathogen-reduced conditions in the Animal Facility of the University of Southern
California (USA). All experiments were performed in accordance with the guidelines
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set by the National Institute of Health and the University of Southern California on the
Care and Use of Animals.
Cell culture. B16 tumor cells (B16-F10, ATCC number: CRL-6475) and 4T1 tumor
cells (ATCC number: CRL-2539) were maintained in a 5% CO2 environment with
Dulbecco’s modified Eagle medium (Mediatech, Inc., Manassas, VA) supplemented
with 10% FBS (Sigma-Aldrich, St. Louis, MO) and 2 mM of L-glutamine (Hyclone
Laboratories, Inc., Omaha, NE). B16-R and 4T1-R cells were produced by
continuously treating B16 and 4T1 cells with 5 µg/ml PTX for 4 days. Then the cells
were recovered by replacing medium with fresh medium without drugs for 7 days. The
remaining cells formed drug resistance for PTX. JC cells (ATCC number: CRL-2116)
were used as a model drug- resistant tumor cell line because it has been showed that JC
cells overexpress P-gp and exhibit drug-resistant phenotype both in vitro and in vivo
[209].
Synthesis of cMLVs. Liposomes were prepared based on the conventional
dehydration-rehydration method. All lipids were obtained from NOF Corporation
(Japan). 1.5 µmol of lipids 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-
dioleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (DOPG), and maleimide-
headgrouplipid1,2-dioleoyl-sn-glycero-3-phosphoeth-anolamine-N-[4-(p-
maleimidophenyl) butyramide (MPB-PE) were combined in chloroform, at a molar
lipid ratio of DOPC:DOPG:MPB = 4:1:5, and the organic solvent in the lipid mixture
was evaporated under argon gas. The lipid mixture was further dried under vacuum
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overnight to form dried thin lipid films. To prepare cMLV (PTX) and cMLV
(Dox+PTX) at a molar ratio of 0.2:1 (drugs:lipids), paclitaxel in organic solvent was
mixed with the lipid mixture to form dried thin lipid films. The resultant dried film
was hydrated in 10 mM Bis-Tris propane at pH 7.0 with (cMLV (Dox) or cMLV
(Dox+PTX)) or without doxorubicin (cMLV (PTX)) at a molar ratio of 0.2:1
(drugs:lipids) with vigorous vortexing every 10 min for 1 h, and then applied with 4
cycles of 15-s sonication (Misonix Microson XL2000, Farmingdale, NY) on ice in 1
min intervals of each cycles. To induce divalent-triggered vesicle fusion, MgCl 2 was
added at a final concentration of 10 mM. The resulting multilamellar vesicles were
further crosslinked by addition of Dithiothreitol (DTT, Sigma-Aldrich) at a final
concentration of 1.5 mM for 1 h at 37°C. The resulting vesicles were collected by
centrifugation at 14,000 g for 4 min and then washed twice with PBS. For pegylation of
cMLVs, the particles were incubated with 1 µmol of 2 kDa PEG-SH (Laysan Bio Inc.
Arab, AL) for 1 h at 37°C. The particles were then centrifuged and washed twice with
PBS. The final products were stored in PBS at 4°C.
In vitro cytotoxicity and data analysis. B16-F10, 4T1, B16-R, 4T1-R, JC cells were
plated at a density of 5 x 10
3
cells per well in D10 media in 96-well plates and grown
for 6 h. The cells were then exposed to a series of concentrations of cMLV (single drug)
or cMLV (drug combinations) for 48 h. The cell viability was assessed using the Cell
Proliferation Kit II (XTT assay) from Roche Applied Science according to the
manufacturer's instructions. Slope m and IC 50 were obtained from median effect
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model, and IIP Cmax was calculated via the equation: IIP Cmax =log (1+(Cmax/IC 50)
m
).
Cmax is maximum plasma drug concentrations for the commonly recommended dose
for each drug.
Cellular uptake of doxorubicin and paclitaxel in cells. 4T1 cells were seeded in 24-well
plates at a density of 2 x 10
5
cells per well and grown overnight. The cells were then
exposed to cMLV(Dox), cMLV(PTX), cMLV(Dox+PTX), and Dox+PTX. The final
concentrations of Dox and PTX are 1 µg/ml for each group. JC cells were seeded at a
density of 10
5
cells per well in D10 media in 96-well plates. The cells were exposed to
cMLV(Dox), cMLV(PTX), cMLV(Dox+PTX), and Dox+PTX. The final
concentrations of Dox and PTX are 5 µg/ml for each group. 48h after treatment, the
cells were washed twice with PBS and lysed with PBS containing 1% Triton X-100.
Doxorubicin and paclitaxel in cell lysates were extracted by 1:1 (v/v)
Chloroform/isopropyl alcohol or ethyl acetate, respectively. Paclitaxel concentrations
in cell lysates were measured by HPLC C18 column and detected at 227 nm (flow rate
1ml/min), and doxorubicin was detected by fluorescence with 480/550 nm
excitation/emission. The concentrations of Dox and PTX were normalized for protein
content as measured with BCA assay (Pierce).
In vivo antitumor activity study. BALB/c female mice (6–10 weeks-old) were
inoculated subcutaneously with 0.2 x 10
6
4T1 breast tumor cells. The tumors were
allowed to grow for 8 days to a volume of ~50 mm
3
before treatment. After 8 days, the
mice were injected intravenously through tail vein with cMLV (2mg/kg Dox), cMLV
123
(2mg/kg PTX), cMLV (2mg/kg Dox: 2mg/kg PTX), cMLV (1mg/kg Dox: 1mg/kg PTX)
every three days (six mice per group). Tumors growth and body weight were
monitored for 40 days or to the end of the experiment. The length and width of the
tumor masses were measured with a fine caliper every three days after injection.
Tumor volume was expressed as 1/2 x (length x width
2
). Survival end point was set
when the tumor volume reached 1000 mm
3
. The survival rates are presented as
Kaplan-Meier curves. The survival curves of individual groups were compared by a
log-rank test.
Immunohistochemistry of tumors and confocal imaging. BALB/c female mice (6–10
weeks-old) were inoculated subcutaneously with 0.2 x 10
6
4T1 or JC tumor cells. The
tumors were allowed to grow for 20 days to a volume of ~500 mm
3
before treatment.
On day 20, the mice were injected intravenously through tail vein with cMLV (5mg/kg
Dox), cMLV (5mg/kg PTX), 5mg/kg Dox + 5mg/kg PTX, cMLV (5mg/kg
Dox+5mg/kg PTX). 3 days after injection, tumors were excised, fixed, frozen, cryo-
sectioned, and mountered onto glass slides. Frozen sections were fixed, and rinsed with
cold PBS. After blocking and permealization, the slides were washed by PBS and then
incubated with TUNEL reaction mixture (Roche, Indianapolis, Indiana) for 1h. For P-
gp expression, after permealization, the slides were stained with mouse monoclonal
anti-P-gp antibody (Abcam, Cambridge, MA) for 1h, followed by stained with
Alexa488-conjugated goat anti-mouse immunoglobulin G (IgG) antibody (Invitrogen,
Carlsbad, CA) and counterstained with DAPI (Invitrogen, Carlsbad, CA).
124
Fluorescence images were acquired by a Yokogawa spinning-disk confocal scanner
system (Solamere Technology Group, Salt Lake City, UT) using a Nikon Eclipse Ti-E
microscope. Illumination powers at 405, 491, 561, and 640 nm solid-state laser lines
were provided by an AOTF (acousto-optical tunable filter)-controlled laser-merge
system with 50mW for each laser. All images were analyzed using Nikon NIS-Elements
software. For quantifying TUNEL and P-gp positive cells, 4 regions of interest (ROI)
were randomly chosen per image at x2 magnification. Within one region, area of
TUNEL or P-gp positive nuclei, and area of nuclear staining were counted by Nikon
NIS-Element software. The data expressed as % total nuclear area stained by TUNEL
or P-gp in the region.
Hematoxylin and Eosin staining of hear sections. The mice bearing 4T1 tumors were
i.v. injected with 5mg/kg Dox + 5mg/kg PTX, cMLV (5mg/kg Dox+5mg/kg PTX). 3
days after injection, heart tissues were harvested, and were fixed in 4% formaldehyde.
The tissues were frozen, cut into sections, and mountered onto glass slides. The frozen
sections were stained with hematoxylin and eosin. Histopathologic specimens were
examined by light microscopy.
Statistics. The differences between two groups were determined with Student’s t test.
The differences among three or more groups were determined with a one-way
ANOVA.
125
5.4 RESULTS
5.4.1 In vitro efficacy study by XTT assay
To achieve combination delivery of doxorubicin (Dox) and paclitaxel (PTX), a
previously developed crosslinked multilamellar liposomal vesicle (cMLV) was used to
incorporate PTX in the lipid membrane and encapsulate Dox in the aqueous core at a
1:1 ratio to form cMLV(Dox+PTX) [191](Yarong). It has been reported that drug
combinations can overcome drug resistance that limits the potential application of
various therapeutics [108]. To determine whether codelivery of Dox and PTX could
overcome drug resistance, an in vitro cytotoxicity assay was performed at a wide range
of concentrations of single drug-loaded cMLVs or cMLV(Dox+PTX)s. As shown in
Figure 33A and 33B, both B16 cells and 4T1 cells developed drug resistance to single
drug-loaded cMLVs, but this resistance can be inhibited by applying cMLV(Dox+
PTX). The maximal cytotoxicity of cMLVs(single drug) observed in these two tumor
cells was between 60%-80%, while cells treated with cMLV (Dox+PTX) showed
significantly more growth inhibition.
To further confirm the efficiency of dual drug-loaded cMLVs in overcoming drug
resistance, drug resistant cell lines B16-R and 4T1-R were generated by continuously
treating parental B16 or 4T1 with high concentration of paclitaxel (5µg/ml). Various
concentrations of cMLV (single drug) and cMLV (Dox+PTX) were incubated with
these two drug resistant cell lines for 48h, and the cytotoxicity was measured by a
standard XTT assay. As shown in Figure 33D and 33E, both B16-R and 4T1-R cells
126
showed a high tolerance when treated with cMLV (PTX) and cMLV (Dox), indicating
that multidrug resistance was developed in these cells. cMLV (Dox+PTX) triggered
significantly more cell death (90-100%) compared to single drug-loaded cMLVs,
confirming that a co-delivery system can overcome drug resistance induced by high
concentration of single drug. Furthermore, in vitro cytotoxicity studies demonstrated
therapeutic efficacy of cMLV(Dox+PTX) in JC cells, a model drug resistant tumor cell
line, corroborating that single drug-loaded cMLVs were less potent than dual drug-
loaded cMLVs. As shown in Figure 33C, the maximal cytotoxicity of cMLV(Dox) and
cMLV(PTX) was in the range of 60-70%, while peak cMLV (Dox+PTX) cytotoxicity
was about 90% in JC cells.
4T1$R&
Drug concentration (µg/ml)
10
-6
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
MLV(Dox)
MLV(Taxol)
MLV(Dox+Taxol)
Cell Viability (%)
Drug concentration (µg/ml)
A
10
-6
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2 MLV(Dox+Taxol)
MLV(Dox)
MLV(Taxol)
Cell Viability (%)
B
10
-6
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
MLV(Dox+Taxol)
MLV(Taxol)
MLV(Dox)
Drug concentration (µg/ml)
Cell Viability (%)
C
E
B16$R&
10
-6
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
MLV(Dox+Taxol)
MLV(Taxol)
MLV(Dox)
Drug concentration (µg/ml)
Cell Viability (%)
IIPmax 0.4
0.5
0.6
0.7
0.8
M(D) M(D+T)
0.4
0.6
0.8
1
1.2
M(T) M(D+T)
IIPmax
0.4
0.6
0.8
1
1.2
M(D) M(D+T)
IIPmax
0.2
0.4
0.6
0.8
1
1.2
1.4
M(D) M(D+T)
IIPmax
0.2
0.4
0.6
0.8
1
M(T) M(D+T)
IIPmax
0.4
0.6
0.8
1
1.2
1.4
1.6
1.8
M(T) M(D+T)
IIPmax
Drug concentration (µg/ml)
10
-6
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
MLV(Dox+Taxol)
MLV(Taxol)
MLV(Dox)
D
Cell Viability (%)
E
cMLV (Dox+PTX)
cMLV (PTX)
cMLV (Dox)
cMLV (Dox+PTX)
cMLV (PTX)
cMLV (Dox)
cMLV (Dox+PTX)
cMLV (PTX)
cMLV (Dox)
cMLV (Dox+PTX)
cMLV (PTX)
cMLV (Dox)
cMLV (Dox+PTX)
cMLV (PTX)
cMLV (Dox)
cMLV(D) cMLV(D+T)
cMLV(D) cMLV(D+T)
cMLV(D) cMLV(D+T) cMLV(T) cMLV(D+T)
cMLV(T) cMLV(D+T)
cMLV(T) cMLV(D+T)
Figure 33. Overcoming drug resistance by codelivery of Dox and Taxol via MLVs. (A, B) In
vitro cytotoxicity of cMLV (single drug) and cMLV (drug combinations) in B16 melanoma
tumor cells (A) and 4T1 breast tumor cells (B). (C, D,E) In vitro cytotoxicity of cMLV (single
drug) and cMLV (drug combinations) in drug resistant JC cells (C), B16-R cells (D) and 4T1-R
cells (E). IIP Cmax was determined by incorporating three parameters (IC 50, D and m) in the
median effect model into the equation: IIP Cmax =log (1+(Cmax/IC 50)
m
). Data represented as
mean ± SD (n=3).
127
Although IC 50, drug concentration that causes 50% inhibitory effect on cell
proliferation, can provide information on the efficacy of drugs, it has been reported
that slope m, parameter mathematically analogous to the Hill coefficient, may also
have a significant effect on the cytotoxicity [210,211]. A new model has been
developed to evaluate drug activety by incorporating three parameters (IC 50, D and m)
from the median effect model into a single value IIP (potential inhibition) with an
intuitive meaning, the log reduction in inhibitory effect [211]. Here, a similar analysis
method was used to evaluate the efficiency of dual drug-loaded cMLVs on the cell
viability. As shown in Figure 33A to 33C (middle and bottom panel), Dox and PTX in
the dual drug-loaded cMLVs have a significantly larger IIP Cmax value in the cells lines
studied compared to the single drug-loaded cMLVs, indicating that combinatorial
cMLVs were more potent in cancer treatment than single drug loaded cMLVs.
5.4.2 Cellular uptake study of doxorubicin and paclitaxel
To investigate the mechanism of enhanced cytotoxicity observed with the cMLV
combination therapy, we evaluated the effect of dual drug-loaded cMLVs on rates of
drug influx/efflux in cells. The intracellular accumulation of doxorubicin and
paclitaxel were examined by HPLC in 4T1 cells following exposure to Dox (1µg/ml)
and PTX (1µg/ml) in cMLVs both individually and in combination, and in JC cells
with higher dose of Dox and PTX (5µg/ml). 3h after incubation, the extracellular
medium was discarded and intracellular drug (Dox or PTX) accumulation was
128
Figure 34. Celluar uptake of doxorubicin and paclitaxel. (A, B) Total cellular uptake of Dox
(A) and PTX (B) in 4T1 cells. 4T1 cells were exposed to cMLV(Dox), cMLV(PTX),
cMLV(Dox+PTX), and Dox+PTX. The final concentrations of Dox and PTX are 1 µg/ml for
each group. (C, D) Total cellular uptake of Dox (C) and PTX (D) in JC cells. JC cells were
exposed to cMLV(Dox), cMLV(PTX), cMLV(Dox+PTX), and Dox+PTX. The final
concentrations of Dox and PTX are 5 µg/ml for each group. The uptake of Dox and PTX were
normalized to protein content measured with BCA assay. All data are shown as the means of
triplicate experiments from three different nanoparticle preparations.
quantitatively determined by analyses of drug concentration in the cell lysates,
normalized by total cellular protein content of the cells. As seen in Figure 34A and 34B,
cMLV(Dox+PTX) significantly increased both Dox and PTX accumulation in 4T1 cells
compared to single drug-loaded cMLVs, suggesting that combination treatments may
overcome drug resistance. In addition, the cMLV combination treatment resulted in
higher cellular accumulation of Dox and PTX than when drug were administrated in
129
solution, probably due to the fact that cMLVs are internalized by cells through
endocytosis [191], bypassing the efflux P-gp pumps. The enhanced cellular
accumulation of drugs in dual drug-loaded cMLVs was also observed in drug resistant
JC cells (Figure 34C and 34D) compared to single drug-loaded cMLVs and
combination solution. These data suggest that cMLV(Dox+PTX) significantly
enhanced the intracellular accumulation of anticancer drugs, through mechanisms
involving both combination treatment and the nanoparticular delivery system.
5.4.3 Effect of codelivery nanoparticles on P-gp expression
Having shown that dual drug-loaded cMLVs enhance cellular accumulation of drugs,
we next sought to verify that this was indeed due to modulation of membrane pumps,
which are responsible for multidrug resistance. We first measured the expression of P-
gp by flow cytometry in 4T1 cells treated with various nanoparticle formulations for 48
h, to test if P-gp involvement in multidrug resistance and decreased drug accumulation
in cells is changed with cMLV formulations [100,212,213]. As shown in Figure 35A,
with single drug loaded cMLVs treatment, the expression of P-gp (in terms of
integrated mean fluorescence intensity) increased significantly in 4T1 cells. This may
lead to development of drug resistance in 4T1 cells. However, dual drug-loaded cMLVs
inhibited expression of P-gp significantly compared to the single drug-loaded cMLVs
and drug combination in solution, suggesting that the combinatorial delivery of Dox
and PTX via cMLVs could efficiently suppress P-gp expression, overcoming MDR.
We next demonstrated whether cMLV (Dox+PTX) could inhibit multidrug resistance
130
in JC cells, which exhibit drug-resistant phenotype by overexpression of P-gp [209].
Figure 35B depicted, that in single drug-loaded cMLVs treatment, expression of P-gp
decreased after 48 h of incubation with JC cells, indicating that the nanoparticular
drug delivery system can at least partially suppress MDR. Moreover, the codelivery
formulation cMLV(Dox+PTX) inhibited P-gp expression significantly compared to
single drug-loaded cMLVs and drug combination in solution. These results, taken
together, indicated that the codelivery of Dox and PTX via cMLVs can inhibit the
expression of P-gp and increase cellular accumulation of drugs, leading to enhanced
drug action in cells, including drug resistant cells.
200
250
300
350
400
450
500
Ctrl cM(D) cM(T) D+T cM(D+T)
350
450
550
650
750
850
Ctrl cM(D) cM(T) D+T cM(D+T)
A
B
Integrated MFI
Integrated MFI
cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T
cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T
Figure 35. Effect of codelivery nanoparticles on P-gp expression. (A) 4T1 cells were exposed to
cMLV(Dox), cMLV(PTX), cMLV(Dox+PTX), and Dox+PTX with the same concentration of
Dox and PTX (1 µg/ml). (B) JC cells were exposed to cMLV(Dox), cMLV(PTX),
cMLV(Dox+PTX), and Dox+PTX with the same concentration of Dox and PTX (5 µg/ml). P-
gp expression was detected by P-gp antibody via flow cytometry. Data represented as mean ±
SD (n=3).
131
5.4.4 Efficacy of dual drug-loaded cMLVs against a breast cancer model
It has been demonstrated that codelivery of Dox and PTX via cMLVs can overcome
drug resistance in vitro, however, due to the complicated in vivo environment, whether
this effect can be translational in vivo has to be determined. Here, a mouse breast
tumor model was used to evaluate the therapeutic efficacy of dual drug-loaded cMLVs
compared with that of single drug liposomal formulations. At day 0, BALB/c mice
were inoculated subcutaneously with 4T1 breast tumor cells. On day 8, mice bearing
tumors were randomly sorted into six groups and each group was treated with one of
the following: PBS (control), cMLV(2 mg/kg Dox), cMLV(2 mg/kg PTX), and
cMLV(2mg/kg Dox+2mg/kg PTX) every three days. Tumor growth and body weights
were monitored untill the end of an experiment (Figure 36A).
As shown in Figure 36B, mice in groups receiving cMLV(2mg/kg Dox) or cMLV
(2mg/kg PTX) showed tumor inhibition compared to those in control group (p < 0.01).
Significantly, cMLV (2mg/kg Dox+2mg/kg Taxol) treatment induced a greater
inhibition than cMLV encapsulating a single drug (p < 0.01). No weight loss was seen
over the duration of the experiment, even at the highest dose of cMLV (2 mg/kg
Dox+2mg/kg PTX) (Figure 36C), indicating that there were no signs of systemic
toxicity from this co-delivery system. The in vivo efficacy of dual drug-loaded cMLVs
against 4T1 tumor model was further confirmed by survival test. As shown in Figure
36D, the groups treated with cMLV(Dox) or cMLV(PTX) had a prolonged life span
compared to control group, while the mice in the group treated with cMLV(Dox+PTX)
132
had a significantly increase life span compared to the groups treated with single drug
loaded cMLVs (p < 0.01).
20 25 30 35 40 45 50
0
20
40
60
80
100
120
control
MLV (Dox)
MLV (Taxol)
MLV (D+T)
Tumor size (mm
3
)
Days after tumor inoculation
**
**
**
Body weight (g)
Days after tumor inoculation
Days after tumor inoculation
B C
D
Challenge with
4T1 cells (0.2 x10
6
)
i.v. injection with
indicated liposomal formulation
every three days (5 times)
8 days
measure tumor size and body weight
every three days
A
% survival
0"
100"
200"
300"
400"
500"
600"
700"
800"
900"
1000"
8" 11" 14" 17" 20" 23"
control
cMLV(Dox)
cMLV(PTX)
cMLV(Dox+PTX)
10
15
20
25
30
8 11 14 17 20 23
control
cMLV(Dox)
cMLV(PTX)
cMLV(Dox+PTX)
control
cMLV(Dox)
cMLV(PTX)
cMLV(Dox
+PTX)
Figure 36. In vivo efficacy of drug combinations via cMLVs against 4T1 tumor model. (A)
Schematic diagram of the experimental protocol for in vivo 4T1 tumor study in BALB/c mice.
(B) Tumor growth was measured after treatment with no injection (black line), cMLV (2 mg/kg
Dox) (red dashed line), cMLV (2 mg/kg PTX) (green line), or cMLV (2 mg/kg Dox+2 mg/kg
PTX) (blue line). Error bars represent standard error of the mean, n = 6 for each treatment
group (**p<0.01). (C) Average mouse weight loss over the duration of the experiment. (D)
Survival curves for 4T1 bearing mice treated with no injection (black line), cMLV 2 mg/kg Dox)
(red line), cMLV (2 mg/kg PTX) (green line), or cMLV (2 mg/kg Dox+2 mg/kg PTX) (blue
line). Survival end point was set when the tumor volume reached 1000 mm
3
. The survival rates
were presented as Kaplan-Meier curves. The survival curves of individual groups were
compared by a log-rank test.
133
5.4.5 Histology study
To study the antitumor mechanism in vivo, TUNEL assay was carried out to detect
tumor cell apoptosis in tumors treated with Dox (5mg/kg) and/or PTX (5mg/kg) in
various formulations for 3 days. As shown in Figure 37A, 4T1 tumors treated by
cMLV(Dox), cMLV(PTX), and Dox+PTX in solution showed significantly more
apoptotic cells compared with the controls. The apoptosis index was significantly
higher in cMLV(Dox+PTX)-treated group compared with other groups. The
quantification data (Figure 37C) further indicated that the increased tumor cell
apoptosis by cMLV(Dox+PTX) treatments in vivo may explain its efficacy as an
antitumor treatment. To further confirm the induction of cell apoptosis in treated
groups, TUNEL assay was performed in drug resistant JC tumors treated with various
formulations for 3 days. As shown in Figure 37B and 37D, cMLV(Dox), cMLV(PTX),
and Dox+PTX induced more apoptotic cells compared to control JC tumors (p < 0.05).
Dual drug loaded cMLVs-treated JC tumors showed remarkably higher apoptosis
index compared to other groups (p < 0.01), confirming the enhanced antitumor
activity of cMLV(Dox+PTX).
To further investigate the innate characteristics of treated tumors, both 4T1 and JC
tumor sections from each treatment group were analyzed for the expression of P-gp
protein. As shown in Figure 36A, P-gp expression level was moderate in the control
group. There appeared to be a significant enhancement of P-gp expression in
cMLV(Dox) and cMLV(PTX) groups, and more remarkable in Dox+PTX group
134
compared to controls. However, there is a remarkable decrease in the
cMLV(Dox+PTX)-treated group as compared to cMLV(Dox), cMLV(PTX), and
Dox+PTX groups, further confirmed by the quantification data (Figure 36C). It is
possible that dual drug-loaded cMLVs treatment may reverse drug resistance induced
by single drug-loaded cMLVs or free drug combination. Interestingly, P-gp was very
high in JC tumor control group shown in Figure 36B. There appeared to be a slight
decrease in cMLV(Dox), cMLV(PTX), and Dox+PTX group, further confirmed by the
quantification data in Figure 36D. Furthermore, there is a more remarkable decrease of
P-gp expression in the cMLV(Dox+PTX) group, indicating that dual drug-loaded
cMLVs may indeed alter the innate characteristics of the multidrug resistant tumors.
Taken this data together, we show that drug-loaded nanoparticles can partially bypass
the efflux pumps P-gp to increase cellular uptake of Dox and PTX, sufficiently induce
cytotoxicity in cancer cells.
135
Ctrl cMLV (D) cMLV (T) D+T cMLV (D+T)
0
5
10
15
20
25
control M (D) M (T) D+T M (D+T)
% positive staining
% positive staining
A
B
C D
0
5
10
15
20
25
30
35
control M(D) M(T) D+T M(D+T)
cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T
Figure 37. Effect of co-delivery cMLVs on tumor apoptosis. (A, B) 4T1 Tumor (A) and
multidrug resistant JC tumor (B) bearing mice were injected intravenously through tail vein
with cMLV (5mg/kg Dox), cMLV (5mg/kg PTX), 5mg/kg Dox + 5mg/kg PTX, cMLV (5mg/kg
Dox+5mg/kg PTX). 3 days after injection, tumors were excised. Apoptosis cells were detected
by TUNEL assay (green), and then costained with nuclear staining DAPI (blue). Scale bar
represents 50 µm. (C, D) Quantification of apoptotic positive cells in 4T1 (C) and JC (D)
tumors. For quantifying TUNEL positive cells, 4 regions of interest (ROI) were randomly
chosen per image at x2 magnification. Within one region, area of TUNEL positive nuclei, and
area of nuclear staining were counted by software. The data expressed as % total nuclear area
stained by TUNEL in the region. Data represented as mean ± SD (n=3).
136
A
B
0
10
20
30
40
50
60
70
control M (D) M (T) D+T M (D+T)
0
20
40
60
80
100
120
control M(D) M(T) D+T M(D+T)
% positive staining of P-gP
% positive staining of P-gP
C D
cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T cMLV (D) cMLV (T) cMLV (D+T) Ctrl D+T
Ctrl cMLV (D) cMLV (T) D+T cMLV (D+T)
Figure 38. Effect of co-delivery cMLVs on P-gp expression in tumros. (A, B) 4T1 Tumor (A)
and multidrug resistant tumor JC (B) bearing mice were injected intravenously through tail
vein with cMLV (5mg/kg Dox), cMLV (5mg/kg PTX), 5mg/kg Dox + 5mg/kg PTX, cMLV
(5mg/kg Dox+5mg/kg PTX). 3 days after injection, tumors were excised, and stained by P-gp
antibody (Green) and costainded by nuclear staining DAPI (blue). Scale bar represents 50 µm.
(C, D) Quantification of P-gp positive cells in 4T1 (C) and JC (D) tumors. For quantifying P-
gp positive cells, 4 regions of interest (ROI) were randomly chosen per image at x2
magnification. Within one region, area of P-gp positive nuclei, and area of nuclear staining
were counted by software. The data expressed as % total nuclear area stained by P-gp in the
region. Data represented as mean ± SD (n=3).
I
t has been reported that doxorubicin treatment results in severe irreversible
cardiotoxicity, leading to myocyte apoptosis [214]. In addition, an unexpected clinical
outcome of combined Dox and PTX was reported due to an increased cardiac toxicity
[113,204,215]. Therefore, systemic toxicity of free Dox+PTX, and cMLV(DOx+PTX)
137
were evaluated to determine whether codelivery cMLVs could decrease the side effect
of combination drug treatment. Treatment with free Doxorubicin (5mg/kg) and PTX
(5mg/kg) in solution caused cadiac toxicity indicated by myofibrillary loss, disarray,
and cytoplasmic vacuolization. However, administration of Dox (5mg/kg) and PTX
(5mg/kg) via cMLVs resulted in no visible loss of myocardial tissue.
5.5 DISCUSSION
Chemotherapeutics are crucial to combating a variety of cancers; however, clinical
outcomes are always very poor, as cancer cells develop a multidrug resistance (MDR)
phenotype after first exposure to the chemotherapeutics. Many efforts have been made
to develop a therapeutic strategy to overcome tumor MDR by enhancing systemic
delivery efficiency of therapeutics to the tumor site and lowering the apoptotic
threshold by combining multiple therapeutics. In this study, we have examined
augmentation of therapeutic efficacy upon coadministration of Dox and PTX using a
crosslinked multilamellar liposomal vesicle (cMLV) in breast cancer cells and drug
resistant JC cells. We demonstrated that combination therapy of Dox and PTX,
especially when codelivered in the cMLV formulations, was very effective in enhancing
the cytotoxicity in both wild-type and drug resistant cells by improving the cellular
accumulation and retention of the drugs. We also showed that the dual therapeutic
strategy efficiently suppressed tumor growth by enhancing the apoptotic response.
P-glycoprotein (P-gp), a membrane-bound active drug efflux pump, is considered one
of the most important mechanisms involved in MDR [100,213]. There has been
138
growing interest in the development of nanoparticle drug delivery systems to
overcome MDR due to their several advantages over conventional chemotherapy.
Nanoparticles, with their unique properties, are able to passively target to the tumor
mass through the enhanced permeability and retention (EPR) effect, enhancing the
accumulation of chemotherapeutics at target sites [85,86,88]. In addition,
nanoparticles can enter cells through the endocytosis pathway, which is thought to be
independent of the P-gp pathway, thus increasing the cellular uptake and retention of
therapeutics in resistant cancer cells [120,121]. A crosslinked multilamellar liposomal
vesicles (cMLV), previously developed in our lab, has shown advantages in cancer
therapy over conventional liposomal formulations due to its sustained drug release,
enhanced vesicle stability and improved drug release, resulting in improved
therapeutic activity with reduced systemic toxicity [191]. Moreover, cMLVs are
internalized by tumor cells through a clathrin-mediated endocytosis [191], suggesting
that cMLVs would be a efficient drug carrier to overcome MDR. In this study, our in
vitro and in vivo results demonstrated that the coadministration of Dox and PTX via
cMLVs efficiently suppressed the P-gp expression in both wild-type and drug resistant
cancer cells.
In addition to nanodelivery, combination of multiple therapeutics has been considered
as another potential strategy to overcome MDR. Combination of Dox and PTX in
cocktail, a standard anthracycline-taxane treatment regimen, was found to be
efficacious in treating a variety of tumors by reducing the individual drug
139
concentration that is required for cytotoxicity, thus overcoming drug resistance
[111,112,113]. However, its clinical outcome was limited by an non-cooridinated
biodistribution of combined drugs [83,114] and an increased cardiac cytotoxicity
[216]. In this study, pharmacokinetics of Dox and PTX were unified by co-
encapsulated into a single cMLV particle. We have demonstrated the therapeutic
efficacy of dual drug-loaded cMLVs in reducing the P-gp expression, increasing
cellular accumulation of drugs, and enhancing the cytotoxicity in cancer cells
including drug resistant cells, relative to single drug-loaded cMLVs. Moreover,
combination therapy of Dox and PTX administered in cMLV formulations has shown
increased efficacy over cMLV monotherapy in suppression of tumor growth by
promoting the apoptotic response in vivo.
5.6 CONCLUSION
In summary, we have developed a multimodal therapeutic strategy to overcome tumor
MDR by codelivery of Dox and PTX via a crosslinked multilamellar liposomal vesicle.
We demonstrated that such a combinatorial delivery system induced an increase of
therapeutic efficacy by enhancing the delivery efficiency to tumors and lowering the
apoptotic threshold of individual drugs, thus overcoming drug resistance. The
properties of cMLVs such as improved stability and sustained release of drugs enable
the nanoaprticles to sufficiently accumulate in tumor sites, subsequently enter tumor
cells via endocytosis to release therapeutics, thus potentially bypassing the P-gp
pathway to enhance cellular retention of therapeutics. Moreover, cMLVs enable the
140
combinatorial delivery of multiple therapeutics to the same action site which lowers
tumor apoptotic threshold of individual therapeutics, potentially inhibiting the MDR.
Taken together, this dual therapeutic strategy can have significant potential in cancer
therapy.
141
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Abstract (if available)
Abstract
Virus‐based nanoparticles have shown promise for mediating gene delivery due to their well‐defined nanostructure and intrinsic bioactive functionality. Adeno‐associated virus (AAV) has been considered as a promising vehicle for human gene therapy based on its ability to infect both dividing and nondividing cells, as well as establish long‐term gene expression in vivo without known pathological consequence of infection. However, because of their native tropisms, the applicability of AAV nanoparticles is often limited to the restricted ranges of cells or tissues. Studies proposed that low expression of receptors/coreceptors on cell surface and an impaired intracellular trafficking pathway of vectors could be the rate-limiting steps of AAV‐mediated transduction. In this study, we have developed two strategies for enhancing AAV‐mediated gene delivery by overcoming these two biological barriers. The first strategy we used is generating a targeted AAV2 vector by genetically encoding an aldehyde tag on viral capsids. Such a tag can be exploited for site‐specific attachment of targeting molecules and allows for further introduction of targeting antibodies or ligands. The results showed that the site‐specific conjugation of targeting antibodies could significantly enhance viral transduction to those target cells that have otherwise exhibited very low permissiveness to AAV2 infection. This method also allows the functional incorporation of RGD peptides onto AAV2 for enhanced delivery with implications for cancer gene therapy. Another strategy we used to enhance AAV transduction, both in vitro and in vivo, is incubating AAV vectors with cell‐permeable peptides (CPPs). We show that CPPs increase internalization of viral particles into cells by facilitating both energy‐independent and energy‐dependent endocytosis. Moreover, CPPs can significantly enhance the endosomal escape process of viral particles, thus enhancing viral transduction to those cells that have exhibited very low permissiveness to AAV2 infection as a result of impaired intracellular viral processing. We also demonstrated that this approach could be applicable to other AAV serotypes. ❧ Non‐viral based nanoparticles offers new hope for cancer detection, prevention and treatment due to their potentials to deliver drugs to tumors. Liposomes constitute one of the most popular nanocarriers for the delivery of cancer therapeutics. However, since their potency is limited by incomplete drug release and inherent instability in the presence of serum components, their poor delivery occurs in certain circumstances. In this study, we address these shortcomings and demonstrate an alternative liposomal formulation, termed crosslinked multilamellar liposomal vesicles (cMLVs). With its properties of improved sustainable drug release kinetics and enhanced vesicle stability, cMLVs can achieve controlled delivery of cancer therapeutics. We further showed that the cMLVs are potential in combination therapy by loading drugs with different hydrophilicities into the same cMLV in a precisely controllable manner over drug ratios. The stability of cMLVs allows an improved loading efficiency and sustained release of doxorubicin (Dox) and paclitaxel (PTX), maximizing the combination therapeutic effect and minimizing the systemic toxicity. Furthermore, in vivo experiments showed that the robust cMLV formulation maintains drug ratios for prolonged times, enabling the ratio‐dependent combination synergy translating from in vitro to in vivo antitumor activity. We also demonstrated that this combinatorial delivery system (cMLV(Dox+PTX)) achieves enhanced drug accumulation and retention, resulting in improved cytotoxicity in tumor cells, including drug resistant cells. Moreover, cMLV(Dox+PTX) significantly overcomes MDR by reducing the expression of P-glycoprotein (P-gp) in cancer cells, thus improving antitumor activity in vivo. With a combined therapeutic ability that enhances drug delivery efficacy to tumors and lowers the apoptotic threshold of individual drugs, cMLV(Dox+PTX) represents a potential multimodal therapeutic strategy to overcome MDR in cancer therapy.
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Liu, Yarong
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Core Title
Engineering nanoparticles for gene therapy and cancer therapy
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Viterbi School of Engineering
Degree
Doctor of Philosophy
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Chemical Engineering
Publication Date
04/02/2014
Defense Date
02/11/2014
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adeno‐associated virus,Combination Therapy,crosslinked multilamellar liposomes,drug ratio,OAI-PMH Harvest,overcome drug resistance,site‐specific,synergy,target gene delivery
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Wang, Pin (
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Tags
adeno‐associated virus
crosslinked multilamellar liposomes
drug ratio
overcome drug resistance
site‐specific
target gene delivery