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University of Iowa
Iowa Research Online
Theses and Dissertations
Spring 2012
Gene therapy for hereditary hearing loss: lessons
from a mouse model
Abraham Matthias Sheffield
University of Iowa
Copyright 2012 Abraham Matthias Sheffield
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/2984
Recommended Citation
Sheffield, Abraham Matthias. "Gene therapy for hereditary hearing loss: lessons from a mouse model." PhD (Doctor of Philosophy)
thesis, University of Iowa, 2012.
http://ir.uiowa.edu/etd/2984.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Genetics Commons
GENE THERAPY FOR HEREDITARY HEARING LOSS: LESSONS FROM A
MOUSE MODEL
by
Abraham Matthias Sheffield
An Abstract
Of a thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Genetics
in the Graduate College of
The University of Iowa
May 2012
Thesis Supervisor: Professor Richard J.H. Smith
1
ABSTRACT
Hearing impairment is the most common sensory deficit worldwide, affecting at
least one child in every one thousand born. Gene therapy targeting the inner ear offers
promise for treatment of genetic forms of hearing loss. Many genetic forms of deafness
are congenital and gene therapies in these cases would require treatment prior to inner ear
maturation. Included in this category is the dominant-negative R75W mutation in GJB2
which encodes connexin 26, a gap junction protein expressed in the supporting cells of
the organ of Corti. RNA interference (RNAi)-based therapeutics offer promise for
treating dominant-negative diseases. Our goal has been the in vivo application of RNAitherapy to the GJB2-R75W transgenic mouse, a model of severe-to-profound dominantnegative hearing loss. Here we describe our efforts to identify a therapeutic, a suitable
delivery route, and an optimal delivery vector. We have designed and optimized siRNA
to achieve robust silencing of the mutant transgene in vitro and have prepared artificial
miRNA constructs for in vivo application. We have determined to use the embryonic
otocyst microinjection technique as the route for therapeutic delivery and have
successfully utilized this technique to study the tropism and safety of several viral vector
(adeno-associated virus 2/1, early- and late-generation adenoviruses, and bovine adenoassociated virus). For the first time we have characterized viral tropism for cochlear
supporting cells following in utero delivery to their progenitor cells in the developing
cochlea and identified bovine adeno-associated virus as a safe vector for gene delivery to
the supporting cells of the cochlea. We have also described two previously unreported
phenotypes in the GJB2-R75W transgenic mouse model: skin disease and cataracts. Both
can be caused by dominant connexin mutations in humans. Our work shows that although
gene therapy is not simple, powerful tools are in place for treating dominant forms of
hereditary hearing loss.
2
Abstract Approved: ____________________________________
Thesis Supervisor
____________________________________
Title and Department
____________________________________
Date
GENE THERAPY FOR HEREDITARY HEARING LOSS: LESSONS FROM A
MOUSE MODEL
by
Abraham Matthias Sheffield
A thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Genetics
in the Graduate College of
The University of Iowa
May 2012
Thesis Supervisor: Professor Richard J.H. Smith
Copyright by
ABRAHAM MATTHIAS SHEFFIELD
2012
All Rights Reserved
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Abraham Matthias Sheffield
has been approved by the Examining Committee
for the thesis requirement for the Doctor of Philosophy
degree in Genetics at the May 2012 graduation.
Thesis Committee: ___________________________________
Richard J.H. Smith, Thesis Supervisor
___________________________________
Michael G. Anderson
___________________________________
Terry A. Braun
___________________________________
Michael D. Henry
___________________________________
Michael J. Welsh
To Erin
ii
He that hath ears to hear,
Let him hear.
Matthew 13:17, New Testament
The woods are lovely, dark, and deep,
But I have promises to keep,
And miles to go before I sleep,
And miles to go before I sleep.
Robert Frost, “Stopping by woods on a snowy evening”
iii
ACKNOWLEDGMENTS
A special Thank You to Dr. Richard Smith for his mentorship, leadership, and
support; to my Thesis Committee (Drs. Michael Anderson, Terry Braun, Michael Henry,
and Michael Welsh) for their encouragement and important role in my education; to
members of the Molecular Otolaryngology Research Laboratories for being there each
day; to Val Sheffield for his counsel and insights; and also to the Animal Caretakers who
made my research possible. I would also like to express my appreciation to the many
people who have assisted in this research: Dr. Michael Hildebrand provided supervision
and assistance throughout my time in the lab; Dr. Sam Gubbels (University of Wisconsin)
taught us the embryonic otocyst injection and continued to provide extensive help with
injections; Dr. Marlan Hansen and members of his lab for assistance with microscopy and
cochlear dissection; Dr. Frederic Venail for his assistance with viral vectors and cochlear
explants; Charles Searby and Qihong Zhang of the Val Sheffield lab for generous and
ongoing help with methods and protocols; Dr. Douglas Brough (GenVec Inc.,
Gaithersburg, MD, USA) for providing the Adf.11D vector as well as valuable
discussion; Drs. John Chiorini and Giovanni DiPasquale (National Institutes of Health)
for provided the BAAV vectors; The University of Iowa Gene Transfer Vector Core for
preparation of Ad5.CMV.GFP; Penny Harding for cryosectioning cochleae; Dr. Mark
Behlke, Dr. Scott Rose and many others (Michael Collingwood, Garret Rettig, Bernardo
Moreira, Kristin Long) at IDT for assistance with siRNA, artificial miRNA, QRT-PCR,
and cloning; Dr. Geoff Lively, Dr. Demelza Koehn, and Adam Hedberg-Buenz of the
Michael Anderson lab for help with eye phenotyping and analysis; Michael Miller of the
Michael Henry lab for assistance with the Odyssey System.
iv
ABSTRACT
Hearing impairment is the most common sensory deficit worldwide, affecting at
least one child in every one thousand born. Gene therapy targeting the inner ear offers
promise for treatment of genetic forms of hearing loss. Many genetic forms of deafness
are congenital and gene therapies in these cases would require treatment prior to inner ear
maturation. Included in this category is the dominant-negative R75W mutation in GJB2
which encodes connexin 26, a gap junction protein expressed in the supporting cells of
the organ of Corti. RNA interference (RNAi)-based therapeutics offer promise for
treating dominant-negative diseases. Our goal has been the in vivo application of RNAitherapy to the GJB2-R75W transgenic mouse, a model of severe-to-profound dominantnegative hearing loss. Here we describe our efforts to identify a therapeutic, a suitable
delivery route, and an optimal delivery vector. We have designed and optimized siRNA
to achieve robust silencing of the mutant transgene in vitro and have prepared artificial
miRNA constructs for in vivo application. We have determined to use the embryonic
otocyst microinjection technique as the route for therapeutic delivery and have
successfully utilized this technique to study the tropism and safety of several viral vectors
(adeno-associated virus 2/1, early- and late-generation adenoviruses, and bovine adenoassociated virus). For the first time we have characterized viral tropism for cochlear
supporting cells following in utero delivery to their progenitor cells in the developing
cochlea and identified bovine adeno-associated virus as a safe vector for gene delivery to
the supporting cells of the cochlea. We have also described two previously unreported
phenotypes in the GJB2-R75W transgenic mouse model: skin disease and cataracts. Both
can be caused by dominant connexin mutations in humans. Our work shows that although
gene therapy is not simple, powerful tools are in place for treating dominant forms of
hereditary hearing loss.
v
TABLE OF CONTENTS
LIST OF TABLES ............................................................................................................. ix
LIST OF FIGURES .............................................................................................................x
LIST OF ABBREVIATIONS ........................................................................................... xii
CHAPTER 1: AN OVERVIEW OF GENE THERAPY .....................................................1
CHAPTER 2: INTRODUCTION ........................................................................................4
Structure and Function of the Auditory System ...............................................4
Development of the Mammalian Ear ........................................................5
Anatomy of the Mammalian Ear ...............................................................6
Hereditary Hearing Loss ...................................................................................8
GJB2 (CX26) and Other Connexins ...............................................................10
Connexin Expression in the Inner Ear .....................................................11
Connexin Function in the Inner Ear ........................................................12
Connexin Mouse Models .........................................................................13
GJB2 Mutations .......................................................................................14
Goal and Organization of Thesis ....................................................................15
CHAPTER 3: MOUSE MODEL OF DOMINANT-NEGATIVE DEAFNESS ...............27
Introduction.....................................................................................................27
GJB2-R75W Transgenic Mouse..............................................................27
Materials and Methods ...................................................................................29
Animal Care and Breeding ......................................................................29
Tissue Samples ........................................................................................30
Tissue Fixation and Paraffin Embedding ................................................30
DNA Extraction .......................................................................................31
RNA Extraction .......................................................................................31
Quantitative Real-Time PCR (QPCR).....................................................32
Genotyping ..............................................................................................32
Results.............................................................................................................34
Hearing Loss ............................................................................................34
Transgene Copy Number Analysis ..........................................................35
Tissue Expression of GJB2-R75W Transgene ........................................36
Skin Disease ............................................................................................37
Cataracts ..................................................................................................39
Discussion .......................................................................................................40
Skin ..........................................................................................................43
Cataracts ..................................................................................................47
CHAPTER 4: DESIGN AND VALIDATION OF RNA INTERFERENCE ....................61
Introduction.....................................................................................................61
siRNA Design ..........................................................................................62
Materials and Methods ...................................................................................66
siRNA Design ..........................................................................................66
Plasmid Constructs ..................................................................................66
vi
Cell Culture and Transfection .................................................................67
Reverse Transcription ..............................................................................68
Quantitative Real-Rime PCR (QPCR) ....................................................68
Artificial miRNA design and construction ..............................................69
Results.............................................................................................................70
Rationally and Computationally Designed siRNA Candidates ...............70
Potent and Specific siRNA-mediated Knockdown .................................71
Artificial miRNA Design and Validation ................................................72
Discussion .......................................................................................................74
siRNA Design ..........................................................................................75
Design Algorithms...................................................................................76
Artificial miRNA vs. shRNA ..................................................................77
Promoter Selection ..................................................................................77
In Vitro Testing of Artificial miRNA Constructs....................................78
CHAPTER 5: VIRAL VECTOR TROPISM FOR SUPPORTING CELLS IN THE
DEVELOPING MURINE COCHLEA ..........................................................95
Abstract ...........................................................................................................95
Introduction.....................................................................................................96
Methods of Cochlear Delivery ................................................................96
Viral Vectors ...........................................................................................98
Materials and Methods .................................................................................102
Virus Production ....................................................................................102
Animals and Virus Administration ........................................................103
Transuterine Otocyst Microinjection .....................................................103
Auditory Brainstem Response (ABR) Testing ......................................104
Sample Preparation and Immunofluorescent Imaging ..........................104
Results...........................................................................................................105
In utero Delivery of Adenovirus to the Developing Murine Inner
Ear..........................................................................................................105
Effect of in utero Adenoviral Transduction on Hearing .......................106
In utero Delivery of Bovine Adeno-Associated Virus to the
Developing Murine Inner Ear ................................................................107
Discussion .....................................................................................................109
CHAPTER 6: IN VIVO DELIVERY OF BAAV.MIRNA TO GJB2-R75W MICE ......124
Introduction...................................................................................................124
Materials and Methods .................................................................................126
BAAV Vector Production .....................................................................126
Cell Culture and Transfection ...............................................................126
Western Blot ..........................................................................................127
Otocyst Injections and Auditory Brainstem Response Testing .............128
Results...........................................................................................................128
Packaging of Artificial miRNA into BAAV .........................................128
In vitro Assay of Knockdown with BAAV ...........................................129
In vivo Delivery .....................................................................................130
Discussion .....................................................................................................132
Otocyst Injections ..................................................................................134
Hearing Tests .........................................................................................136
CHAPTER 7: CONCLUSIONS AND LESSONS LEARNED ......................................141
vii
Summary .......................................................................................................141
Lessons Learned ...........................................................................................143
RNA Interference ..................................................................................143
Otocyst Injections ..................................................................................145
Future Studies ...............................................................................................149
REFERENCES ................................................................................................................154
viii
LIST OF TABLES
Table 1. Autosomal dominant non-syndromic (DFNA3) and syndromic GJB2
mutations..............................................................................................................22
Table 2. Primers used for genotyping mice. ......................................................................51
Table 3. QPCR primer and probe sequences. ....................................................................84
Table 4. QPCR reaction conditions. ..................................................................................85
Table 5. Sequences of siRNA and D-siRNAs. ..................................................................87
ix
LIST OF FIGURES
Figure 1. Development of the murine inner ear. ................................................................17
Figure 2. Anatomy of the ear. ............................................................................................18
Figure 3. Diagram of a cross-section of the cochlea. .........................................................19
Figure 4. Supporting cells of the organ of Corti. ...............................................................21
Figure 5. Schematic diagram showing connexin topology and channel variety................23
Figure 6. Heteromeric connexon compositions. ................................................................24
Figure 7. Deafness- and skin disease-causing CX26 mutations. .......................................25
Figure 8. R75W plays a role in inter-connexin interactions and oligomerization. ............26
Figure 9. Construct of the GJB2-R75W transgene. ...........................................................52
Figure 10. Hearing loss associated with the GJB2-R75W transgene. ...............................53
Figure 11. Quantitative results of HsGJB2-R75W in the transgenic mice. .......................54
Figure 12. Phenotype of palmoplantar keratoderma with deafness. ..................................56
Figure 13. PPK-like skin disease in GJB2-R75W transgenic mice. ..................................57
Figure 14. Cataracts in GJB2-R75W transgenic mice. ......................................................58
Figure 15. Lens histology in GJB2-R75W transgenic mice. .............................................59
Figure 16. Expression of GJB2 (CX26) in the cochlea. ....................................................60
Figure 17. RNA interference (RNAi) pathways. ...............................................................83
Figure 18. Sample QPCR output. ......................................................................................86
Figure 19. In vitro siRNA knockdown of GJB2-R75W transgene. ...................................88
Figure 20. Predicted artificial miRNA structures. .............................................................89
Figure 21. Vector maps for pAAV-tVal-CMV-dsRED2 and pAAV-H1-CMVdsRED2. ............................................................................................................90
Figure 22. In vitro artificial miRNA silencing of GJB2-R75W transgene. .......................91
Figure 23. In vitro artificial miRNA silencing by pAAV_tVal-miRNAs. ........................92
Figure 24. Experimental outline for in vitro silencing assays. ..........................................93
Figure 25. HsGJB2-MmGjb2 sequence alignment and siRNA target sites. ......................94
x
Figure 26. Microinjection into the developing mouse otocyst. .......................................114
Figure 27. Surgical room setup. .......................................................................................115
Figure 28. Adenovector transduction of inner ear. ..........................................................116
Figure 29. Adenoviral transduction of cochlear cells in the adult mouse inner ear. ........117
Figure 30. Auditory assessment of mice injected with viral vectors. ..............................119
Figure 31. Pure tone ABR thresholds of mice injected with BAAV. ..............................120
Figure 32. BAAV transduction of cochlear cells in the adult mouse inner ear. ..............121
Figure 33. BAAV transduction in the cochlear apex and base. .......................................122
Figure 34. Gradient of BAAV transduction from cochlear apex to base. ........................123
Figure 35. BAAV.miGJB2-D expression construct. .......................................................137
Figure 36. In vitro silencing of GJB2-R75W transgene with BAAV.miGJB2-D. ..........138
Figure 37. Auditory brainstem response results for mice injected with BAAV
constructs. .......................................................................................................139
Figure 38. Progressively diminishing returns in the otocyst injection procedure............140
Figure 39. Predicted connexon channel composition based on stoichiometric ratios. ....153
xi
LIST OF ABBREVIATIONS
AAV = Adeno-associated virus
ABR = Auditory brainstem response
ADNSHL = Autosomal dominant non-syndromic hearing loss
ARNSHL = Autosomal recessive non-syndromic hearing loss
AV = Adenovirus
BAAV = Bovine adeno-associated virus
CHO = Chinese hamster ovary cell line
CMV = Cytomegalovirus promoter
CX26 = Homo sapien connexin 26 (GJB2 product)
Cx26 = Mus musculus connexin 26 (Gjb2 product)
CX30 = Homo sapien connexin 30 (GJB6 product)
Cx30 = Mus musculus connexin 30 (Gjb6 product)
dB = Decibel
DFNA3 = Autosomal dominant non-syndromic hearing loss locus (GJB2/GJB6)
DFNB1 = Autosomal recessive non-syndromic hearing loss locus (GJB2/GJB6)
D-siRNA = Dicer substrate small interfering RNA
gDNA = Genomic DNA
GFP = Green fluorescent protein
H1 = H1 RNase P RNA promoter (weak pol III)
HsGJB2 = Homo sapien gap junction beta 2 (GJB2)
HsGJB6 = Homo sapien gap junction beta 6 (GJB6)
IHC = Inner hair cell
miRNA = MicroRNA
MmGJB2 = Mus musculus gap junction beta 2 (GJB2)
MmGJB6 = Mus musculus gap junction beta 6 (GJB6)
xii
mRNA = Messenger RNA
OHC = Outer hair cell
pol III = RNA polymerase III
PPK = Palmoplantar keratoderma
QPCR = Quantitative real-time PCR
QRT-PCR = Quantitative reverse transcription PCR
R75W = p.Arg75Trp substitution in HsGJB2 (c.223C>T)
RISC = RNA-induced silencing complex
RNAi = RNA interference
SGN = Spiral ganglion neuron
shRNA = Short hairpin RNA
siRNA = Small interfering RNA
SNHL = Sensorineural hearing loss
tVal = Transfer RNA (tRNA)-valine promoter (weak pol III)
xiii
1
CHAPTER 1: AN OVERVIEW OF GENE THERAPY
Physicians, researchers, and patients have anticipated the time when effective
therapeutics would be developed for Mendelian disorders. Encouraged by dramatic
Hollywood depictions of novel therapies (e.g. Lorenzo’s Oil, Extraordinary Measures)
and frequent discussion of a future in which “individualized” or “personalized” medicine
is routinely practiced, the public’s high expectations for elegant new cures have, so far,
gone largely unfulfilled. In fact, early gene therapy trials had disastrous consequences. In
1999, 18-year-old Jesse Gelsinger was injected with an adenoviral vector designed to
treat ornithine transcarbamylase deficiency. He died four days after the injection, having
suffered a fatal immune response to the treatment. About 10 years ago, gene therapy for
severe-combined immune deficiency (SCID) was administered to ten children in Europe.
Clinical trials were halted when two of the ten children developed leukemia. Despite
these setbacks, the public’s expectations for gene therapy remain high.
2011 marks the 10th anniversary of the completed draft of the Human Genome
Project.[1, 2] The announcement of the draft sequence was announced by President Bill
Clinton predicting that it would “revolutionize the diagnosis, prevention and treatment of
most, if not all, human diseases.”[3] In time this statement will likely become reality,
however the hype surrounding the third promise (“treatment”) too often overshadows the
first two promises (“diagnosis” and “prevention”). A recent newspaper headline reads,
“A decade later, genetic map yields few new cures,” and suggests a disappointing lack of
progress in the past ten years.[3] The timeline of expectation can be much shorter than
the timeline of reality, especially when it comes to treatment of disease.
Regardless of the perceived “disappointment” in the eyes of some, geneticists are
well aware that the completion of the Human Genome Project represented a
commencement, not a conclusion, to our understanding of human genetics. It lifted us to
2
a new elevation, providing a clearer view of the bright future—and the work required to
get there.
Genetics has made great strides in the past decade, including in therapeutics. For
example, eye researchers recently used sub-retinal injections of an adeno-associated viral
vector to deliver a corrective gene to patients with a form of blindness called Leber’s
congenital amaurosis (LCA).[4] The patients receiving the treatment have reported
dramatic results. Children who were using Braille prior to treatment are now reading
books. An elementary school student who received the treatment is now able to see the
chalk board and participate in class without the assistance of a computer or an aide.[5]
Therapeutic progress has also been recently reported for other diseases, including
adrenaleukodystrophy (ALK) and β-thalassemia.[6, 7] These exciting advancements have
prompted some to scientists to observe that gene therapy is making a “comeback”.[8]
The field of hearing and deafness research has seen great progress in the
identification of mutant genes, understanding the molecular mechanisms of hearing, and
providing patients with a genetic diagnosis.[9] For example, our lab recently utilized
massively parallel sequencing technologies to develop a sensitive and specific genetic
test (OtoSCOPE) to screen for mutations in all known deafness genes.[10] This genetic
test will provide accurate diagnosis, prognostic information, and help determine best
treatment options for an individual patient. The next step in the field of hearing loss
genetics is to develop and test treatments to well-defined forms of hereditary hearing loss.
The recent successes with gene therapy in the eye have been encouraging to us.
Like the eye, the inner ear is a surgically accessible, immune privileged organ that lends
itself well to therapeutic interventions.[11, 12] The molecular genetics of many forms of
hereditary hearing loss are well-described and a number of studies involving gene
transfer to the rodent cochlea have been published in the last decade.[13] Despite these
advantages, the inner ear is an exquisitely complex structure and presents a number of
challenges which must be overcome prior to therapeutic success. Success is most likely to
3
be realized when the therapeutic strategies are informed by basic science research and
understanding of the disease process. In this Thesis I discuss my work to design gene
therapy for a specific dominant-negative deafness mutation and strategies to overcome
various roadblocks to success.
A decade ago Francis Collins, then the director of the public Human Genome
Project, promised that “over the longer term…you will see a complete transformation in
therapeutic medicine.”[3] The realization of such therapies must be founded upon
knowledge gained from basic genetic research.[14] Given the difficult lessons taught by
early gene therapy trials, “Proceed with Caution” would be a good motto for physicians
and scientists who seek gene-based treatments to disease. But given the explosion of
knowledge and reported successes in the past decade, it is important that we proceed.
With the power of gene-based diagnostics and detailed molecular genetic understanding
of disease processes, new therapies for Mendelian disorders will continue to roll forth. To
paraphrase Robert Frost, geneticists have promises to keep and miles to go before we
sleep.[15]
4
CHAPTER 2: INTRODUCTION
Hearing loss is the most common sensory disorder in developed countries. It is an
etiologically heterogeneous trait with many known genetic and environmental (infections,
noise exposure, premature birth, exposure to ototoxic drugs) causes. Hearing loss is
usually classified as conductive hearing loss or sensorineural hearing loss (SNHL) and
can occur in isolation (nonsyndromic) or be associated with other symptoms (syndromic).
Conductive impairment can be caused by otitis media, otosclerosis, or obstruction by a
tumor or other foreign body and is often reversible. Bilateral congenital SNHL affects
one in every 500 newborns and may lead to significant problems in speech development,
educational attainment, employment prospects, and life expectancy.[13, 16] Presbycusis
(age-related hearing loss) is also highly prevalent in developed societies, affecting 2540% of individuals over the age of 65. Hearing aids can provide significant amplification
to patients, although for severe-to-profound hearing loss cochlear implantation is usually
the better habilitation option. Despite the marked benefits that can be obtained through
these interventions, prevention of cochlear damage or restoration of normal function is
the ideal outcome. Genetic-based therapies offer potential for intervention to prevent
hearing loss or restore inner ear function.
Structure and Function of the Auditory System
In 1961 Dr. Georg von Békésy accepted the Nobel Prize in Physiology or
Medicine for his work in describing how sound waves are discriminated in the inner ear.
During his Nobel Lecture, Dr. von Békésy said that he was drawn to studying the ear
because “I am always ready…to look at beautiful things.”[17] The ear is an organ of
beautiful complexity, containing multiple compartments and a diverse variety of cell
types. As Nature’s solution to interpreting pressure waves as sound, the ear is an
exquisite model of form fitting function.
5
Development of the Mammalian Ear
The vertebrate ear is derived from all three germ layers of the embryo and the
neural crest. The middle and outer ear develop from the first pharyngeal pouch and the
surrounding wall of the pharynx.[18] The inner ear develops from otic placode tissue,
bilateral thickenings within the ectoderm adjacent to the hindbrain. In mice, otic placodes
first appear at embryonic day 8.5 (E8.5). During the next 24 hours, the otic placodes
invaginate to form the otocyst (also called the otic vesicle). The otocyst will eventually
give rise to all of the cells within the organ of Corti, as well as most other cell types
within the membranous portion of the inner ear.[19] Soon after formation of the otocyst,
neuroblasts delaminate from its ventral region and migrate adjacent to the developing
inner ear to form the statoacoustic ganglion (SAG). The SAG becomes the ganglion of
cranial nerve VIII. At E10.5 the spherical otocyst begins to undergo an elaborate series of
changes which result in the formation of distinct dorsal and ventral regions. The dorsal
protrusion will eventually develop into the endolymphatic duct (ED), and the ventral
cochlear duct (CD) will eventually form the cochlea (Figure 1). By E12.5, the developing
semicircular canals (anterior, posterior and lateral) can be identified. Each canal forms
from an outgrowth of the otocyst, the central portion of which undergoes resorption. Also
at approximately E12.5, the CD begins to extend and form a spiral. As the CD extends, a
subset of cells within its ventral region begins to develop as the sensory epithelium which
will become the organ of Corti. At E13.5, the spiraling cochlea has extended to a threequarter turn and neurons from the developing SAG begin to extend dendrites which will
form contacts with developing hair cells throughout the sensory epithelia. The inner ear
continues to grow between E15.5 and E17.5 until the cochlea reaches its mature length of
one and three-quarter turns.[20] The cells of the organ of Corti will continue to develop
until they eventually mature and differentiate into inner and outer hair cells and a variety
of supporting cells. In contrast with human auditory development, organ of Corti
6
maturation in mice continues into the postnatal period, with hearing onset occurring at
approximately postnatal day 12 (P12).[19]
Anatomy of the Mammalian Ear
The ear functions to facilitate conversion of mechanical (acoustic) stimuli into
electrical (nerve) impulses. The anatomy of the ear can be divided into the external,
middle, and inner ear (Figure 2). The external ear is comprised of the pinna (auricle) and
the external auditory canal from the meatus to the tympanic membrane (ear drum). The
external ear channels sound waves to the tympanic membrane. The tympanic membrane
vibrates in response to the sound waves and relays this mechanical energy to the middle
ear, an air-filled compartment within the temporal bone. The middle ear contains three
ossicles—the malleus (hammer), incus (anvil) and stapes (stirrup)—linking acoustic
energy from the air-filled external auditory canal to the fluid-filled cochlea of the inner
ear.
The inner ear can be divided into a bony labyrinth and a membranous labyrinth.
The bony labyrinth consists of three cavities encased by dense bone—the semicircular
canals (superior, posterior, lateral), the vestibule, and the cochlea. These cavities are
filled with perilymph fluid. The membranous labyrinth is suspended within the bony
labyrinth. The major components of the membranous labyrinth include the utricle and
saccule in the vestibule, the three semicircular ducts located in the semicircular canals
and the membranous duct of the cochlea. The structures of the membranous labyrinth are
filled with a specialized fluid called endolymph.
The two major sensory components of the inner ear are the cochlea (responsible
for hearing) and the vestibular system (responsible for balance). The vestibular system
detects both the position and motion of the head. The three semicircular ducts detect
angular acceleration and deceleration in all three planes. The utricle and saccule detect
position of the head relative to gravity as well as changes in position. The cochlea is a
7
coiled, bony tube divided into three separate compartments: the scala vestibule, scala
media and scala tympani (Figure 3). The scala vestibule and the scala tympani are filled
with perilymph, an extracellular- like fluid with high sodium concentration (139 mEq/L)
and low potassium concentration (4 mEq/L). The scala media is filled with endolymph,
an intracellular-like fluid with a low sodium concentration (13 mEq/L) and a high
potassium concentration (144 mEq/L). The scala media has a resting potential of
approximately +80 mV, referred to as the endocochlear potential (EP). Strict maintenance
of the EP is required for normal auditory function. The scala media is divided from the
perilymphatic compartments by the basilar membrane (scala tympani boundary) and
Reissner’s membrane (scala vestibuli boundary) (Figure 3). The entire endolymphatic
space is protected from surrounding structures by a layer of luminal cells connected by
tight junctions.[21]
The sensory cells of the cochlea reside in the organ of Corti which rests on the
basilar membrane and is comprised of a number of specialized cell types which can be
broadly categorized as sensory or supporting cells. The two types of sensory cells, the
outer (OHC) and inner (IHC) hair cells, are arranged in ordered rows. The tectorial
membrane, an acellular membrane connected to the spiral limbus, overlies the organ of
Corti and is connected to the stereocilia tips of the OHC. A single row of IHCs is located
on the medial edge of the organ of Corti, while three rows of OHCs are located on the
lateral edge. IHCs and OHCs are functionally and morphologically distinct. IHCs are
goblet-shaped cells topped with stereocilia. IHCs are the true sensory receptors of the
cochlea. They convert the mechanical energy of sound waves into electrical energy when
mechanosensitive transduction ion channels open on the surface of the stereocilia
projecting from their apical surface.[19] The positive EP causes K+ influx when the ion
channels open, leading to intracellular depolarization and a signaling cascade which
culminates in activation of afferent nerve fibers. The cylindrically-shaped OHCs, also
with apical stereocilia projections, have been identified as the cochlear amplifier.[22] The
8
stereocilia of the OHCs are connected to the tectorial membrane and the OHCs can
actively modify the motion of the basilar membrane to enhance frequency selection and
detection of low intensity sound.[23]
In addition to the sensory hair cells, non-sensory supporting cells are the other
major cell type in the organ of Corti. The mammalian organ of Corti contains at least
seven distinct types of supporting cells (Figure 4): inner sulcus cells, pillar cells,
border/inner phalangeal cells, Deiters cells, Hensen cells, and Claudius cells. These cells
rest on the basilar membrane and support the sensory hair cells. They contribute to
maintenance of cochlear homeostasis and loss of supporting cells eventually results in
death of hair cells. The specific role of supporting cells will be further discussed
below.[19]
Hereditary Hearing Loss
Evident from its anatomy and cellular composition, the cochlea is a finely tuned
structure which is susceptible to damage, either through environmental insult or through
genetic mutation. 50-60% of congenital and early-onset SNHL cases have a Mendelian
basis. Of these, the majority (~70% of cases) are non-syndromic hearing loss (NSHL)
and the remainder (~30% of cases) are associated with syndromes such as Pendred and
Usher syndrome.[16] Genetic hearing loss is highly heterogeneous with autosomal
recessive inheritance (ARNSHL) in ~80% of cases, autosomal dominant (ADNSHL)
inheritance in ~20% of cases, and fewer instances of X-linked (<1%) and mitochondrial
(<<1%) inheritance.[24] Recessive deafness is most commonly congenital and profound,
while dominant deafness tends to be later-onset and progressive.
Underscoring the complexity of the inner ear and the array of proteins required
for hearing, it is estimated that the mutation of any of several hundred genes may result in
deafness. To date a total of 36 ARNSHL genes and 24 ADNSHL genes have been
identified. A further 45 loci for recessive and 34 loci for dominant deafness have been
9
mapped to chromosomal regions.[9] Known deafness mutations affect a variety of inner
ear cells and tissues including: sensory hair cells (e.g. MYO7A, KCNQ4), non-sensory
supporting cells (e.g. GJB2, GJB6), and the tectorial membrane (e.g. COL11A2,
TECTA).[9, 16, 25]
Despite the significant locus heterogeneity in hereditary deafness, mutations in a
single gene, Gap Junction Beta-2 (GJB2), are responsible for at least 50% of prelingual
ARNSHL (DFNB1; MIM#220290) cases in some populations.[26] The DFNB1 locus
contains the deafness genes GJB2 and GJB6 that encode connexin 26 (CX26) and CX30
proteins, respectively [27, 28]. Approximately 100 GJB2 mutations have been associated
with ARNSHL. The c.35delG variant is the single most common cause of ARNSHL in
Caucasian individuals, with a carrier rate of 2-4%.[29, 30] A founder mutation arising
approximately 10,000 years ago in southern Europe is thought responsible for the high
rates of this mutation in these populations.[31, 32]
A total of over one hundred different deafness-causing mutations in GJB2 have
been reported.[9, 33] Although not nearly as prevalent as the recessive mutations, a
number of mutations in GJB2 are transmitted via autosomal dominant inheritance
(DFNA3 deafness (MIM#601544)). This type of deafness has childhood onset, is
progressive, moderate-to-severe and preferentially affects the high-frequencies, though
the audioprofiles may vary from individual to individual.[33] The majority of DFNA3
mutations cause prelingual, progressive hearing loss (W44C, P58A, R75Q, R75W, and
R143Q). Two mutations (D179N and C202F), however, result in post-lingual hearing
loss, ranging in onset from the first to third decade of life.[34] The prevalence of DFNA3
is not known, but it is relatively rare. Ten DFNA3 mutations have been described and
most of these mutations have only been identified in single families or sporadic cases
(Table 1).[34] Two of the DFNA3 mutations (R75Q and R75W) are also reported to
cause autosomal dominant syndromic hearing loss associated with a skin disorder called
palmoplantar keratoderma (PPK).[35, 36] In addition to the R75Q and R75W mutations,
10
close to twenty other dominant mutations in GJB2 cause syndromic hearing loss
associated with distinct skin manifestations such as Keratitis-Ichthyosis-Deafness (KID)
syndrome (MIM#148210), Vohwinkel syndrome (MIM 124500), and palmoplantar
keratoderma (PPK) with deafness (MIM#148350).[37-39]
GJB2 (CX26) and Other Connexins
The coding sequence of GJB2 (exon 2) is 681 base pairs (including the stop
codon) and is translated into a 226-amino acid protein, connexin 26 (CX26). Connexin 26
is a beta-2 gap junction protein. Intercellular signaling is fundamental to maintenance of
homeostasis as well cell-cell communication in multicellular organisms. Gap junctions
are highly specialized organelles consisting of clustered channels that permit direct
intercellular exchange of ions and molecules through central aqueous pores. A gap
junction contains clusters of tens to thousands of individual “gap junction channels.”
Each gap junction channel is formed by end-to-end docking of two gap junction hemichannels in adjacent cells. Each hemi-channel, or “connexon,” is composed of six
connexin proteins surrounding a central pore.[40] Each individual connexin protein
contains two extracellular (E1-E2), four transmembrane (M1-M4), and three cytoplasmic
domains (N-terminus, C-terminus, and a cytoplasmic loop located between M2 and M3).
Each extracellular domain contains three cysteine residues joined between the E1 and E2
loops by at least one disulfide bond.[41, 42] The third transmembrane domain (M3) is
amphipathic and lines the putative wall of the pore.[41] E1 and E2 domains are the most
highly conserved of the connexin protein domains and are thought to be involved in the
docking of two connexons between cells. The C-terminus and the cytoplasmic loop (CL)
display the most variability between connexin isoforms and are thought to govern
channel properties.[43]
21 connexins have been identified in humans and twenty in mice. Although they
all have similar structure they also exhibit differences in ionic selectivity and gating
11
mechanisms.[44, 45] In general, gap junction channels are permeable to ions and small
metabolites with relative molecular masses up to approximately 1 kilodalton (kDa), but
the size and charge of molecules able to pass through a gap junction is dependent on the
composition of the gap junction channel. The unique permeability properties of different
gap junction channels is termed “permselectivity.”[43] Different connexin isoforms can
interact with each other to form channels with unique permselectivity. A connexon
composed of a single connexin isoform is termed “homomeric” and a connexon
composed of at least two different connexin isoforms is termed “heteromeric”; likewise, a
“homotypic” gap junction channel is formed by two identical connexons and a
“heterotypic” gap junction channel is formed by at least two different connexons (Figure
5).[43]
Connexin Expression in the Inner Ear
Connexins are expressed in most tissues and tissues can express multiple
connexin isoforms at one time. Although not all isoforms are compatible for
oligomerization with one another, in general, connexin assembly is thought to be a
stochastic process. Thus, in a cell that expresses two connexin isoforms, there are 14
possible different connexon possibilities with seven different stoichiometric ratios
(Figure 6).[46] In some cases the gap junctional channel reflects the permselectivity of its
most abundant member, but there are also cases in which the channel displays
permselectivity unique to either of its members.[43] Thus, channels in a single cell can
display a range of properties, especially in tissues expressing more than one or two
connexins.
The inner ear expresses multiple connexin isoforms including CX26, CX29,
CX30, CX31, and CX43.[43] The predominant connexin isoforms of the cochlea are
CX26 and CX30. They are expressed between interdental cells of the spiral limbus, the
supporting cells of the organ of Corti, and cells within the spiral ligament. CX26 and
12
CX30 are co-expressed and form homo- and heterotypic gap junction channels. These
channels form two gap junction networks within the cochlea: the epithelial network and
the connective tissue network. The epithelial gap junction network connects the
supporting cells of the sensory epithelia and bordering epithelial cells; the connective
tissue network is found in the lateral wall between basal and intermediate cells of the stria
vascularis and fibrocytes of the spiral ligament.[47, 48]
Connexin Function in the Inner Ear
The function of gap junctions in the cochlea is less clearly defined than their
expression. Gap junction networks are thought to be involved in buffering and recycling
of K+ following mechanotransduction by the sensory hair cells.[49] This “potassium
recycling model” proposes that K+ ions exit sensory hair cells after depolarization and
immediately enter adjacent supporting cells. The ions then move from cell to cell via the
epithelial gap junction network to the extracellular space of the lateral wall, where they
are then taken up by the Na+/K+-ATPase and Na+/2Cl-/K+-cotransporter of spiral ligament
fibrocytes where the ions enter the connective tissue network of the lateral wall. The
connective tissue gap junction network shuttles the K+ ions to the stria vascularis and
they are eventually transported back into the endolymph via another Na+/K+-ATPase and
Na+/2Cl-/K+-cotransporter.[50] Recently, a more complex picture of the roles of gap
junctions in cochlear physiology has emerged. Gap junctions have been implicated in
intercellular signaling within the cochlea. In addition to K+, gap junctions have been
shown to transport signaling molecules such as ATP, cAMP, IP3.[51, 52] It has been
suggested that connexons may be involved in Ca2+-activated ATP release leading to
propagation of calcium waves in the sensory epithelium. A role for connexons in
maintaining osmotic balance in the stria vascularis has also been proposed.[52] Although
the many possible functions of connexin proteins in the cochlea continue to be debated, it
13
is clear from human patients and mouse models that CX26 and CX30 play a crucial role
in cochlear development, maintenance of homeostasis and, ultimately, function.
Connexin Mouse Models
Mice are important model organisms for studying hearing and deafness, but there
has never been a mouse reported with spontaneous deafness-causing mutations in Gjb2 or
Gjb6. However, the function of Cx26 has been extensively studied through the creation
of knockout and transgenic mice. In contrast to humans where homozygous loss-offunction mutations in GJB2 cause non-syndromic hearing loss, a complete knockout of
Gjb2 in mice results in embryonic lethality. Cx26 is involved in transplacental uptake of
glucose in mice, a role played by Cx43 in humans. Due to embryonic lethality caused by
Gjb2 deletion, targeted knockouts are required to assess the role of Cx26 in the inner
ear.[53]
The first inner ear-specific knockout of Gjb2 was the Cx26OtogCre mouse. It was
made by crossing Cx26loxP/loxP mice with mice in which Cre expression is driven by
the Otogelin promoter.[54] This resulted in specific Cx26 deletion in the cells of the
epithelial gap junction network. Cx26OtogCre mice have ~30 dB hearing loss by 3 weeks of
age. The inner ear and the endocochlear potential develop normally up until ~P14. At P14
apoptosis is observed in the supporting cells of the organ of Corti, beginning with
supporting cells surrounding the inner hair cells and then extending to those around the
outer hair cells, eventually leading to loss of outer hair cells. Surviving inner hair cells
and supporting cells exhibit dysmorphology. Although the cochlea developed normally
up until 2 weeks of age, the endocochlear potential in adult Cx26OtogCre mice was
significantly diminished, as was the potassium concentration of the endolymph. This loss
of endolymphatic homeostasis is due to breaches in the epithelial barrier of the
endolymphatic compartment.[55] This indicates that Cx26 expression in the epithelial
14
gap junction network is not necessary for establishing the endocochlear potential, but is
important for maintaining the integrity of the sensory epithelium.
A group of researchers at Emory University made three different conditional
knockout mice (cCx26 null) to study the role of Cx26 in the cochlea: foxg1-Cre, pax2Cre, and TMX-inducible.[56, 57] These mice had 40-50 dB loss in hearing and impaired
postnatal organ of Corti development. The cCx26 null mice failed to form the tunnel of
Corti or space of Nuel. Degeneration was first observed at P8 in the Claudius cells, with
death of outer hair cells and the surrounding supporting cells beginning at ~P13. Cell
death begins in the middle cochlear turn and gradually spreads to the basal and apical
turns. By one month of age the organ of Corti in the middle turn is completely
degenerated. These mice support a critical role for Cx26 in the postnatal maturation of
the organ of Corti prior to onset of hearing and indicate that cell death occurs secondary
to developmental anomalies.[57]
GJB2 Mutations
Dominant mutations in GJB2 cause deafness via a dominant-negative mechanism.
A number of the dominant mutations in GJB2 have been functionally tested for dominant
negative effects in recombinant expression systems (W44C, W44S, R75W, R75Q, and
M163L). The ability of mutant GJB2 connexins to prevent formation of functional gap
junction channels was first demonstrated with the R75W mutation in a Xenopus oocyte
model.[58, 59] The W44C, W44S, and R75Q mutations have been shown to prevent
functional channel formation in vitro.[60, 61] In addition to dominant-negative inhibition
of wild-type CX26, the W44S and R75W mutations show a transdominant-negative
effect on wild-type Cx30 channel formation. It is particularly interesting that the R75W
mutant is also able to exert a transdominant-negative effect on CX43, a connexin with
which wild-type CX26 is not known to interact. CX43 is expressed in the skin and the
gain-of-function transdominant interaction with CX43 suggests a mechanism whereby
15
the R75W mutant causes skin disease.[46] A GJB2-R75W transgenic mouse was created
by Kudo and colleagues.[47] This is a mouse model of a dominant-negative mutation in
CX26. The mouse has significant hearing loss from 2 weeks of age. We have obtained
this mouse for our research and it will be discussed in greater detail in the next chapter.
The GJB2 mutations associated with syndromic deafness cluster in the E1 domain
of CX26 (Figure 7). This is one of the most conserved regions of all connexins. Recently,
Maeda and colleagues produced a high resolution crystal structure at a resolution of 3.5Å.[62] This structure has provided insight into the function of connexin domains as well
as specific amino acids within the protein. A number of dominant mutations in GJB2 that
are associated with deafness and skin disease were shown to be involved in interconnexin interactions or connexon docking. The highly conserved E1 and E2 domains are
involved in connexon-connexon docking. The R75 residue (mutated in R75W and R75Q)
is located at the E1/TM2 periphery. Interactions with this residue are predicted to
contribute to appropriate folding and positioning of extracellular loops for docking as
well as stabilization of inter-connexin interactions. Replacing the Arg75 with a Trp
residue would disrupt two salt bridges that R75 participates in (intramolecular to E47 and
intermolecular to E187) (Figure 8). This structural evidence confirms in vitro evidence
that the R75W mutation exerts a dominant-negative effect on connexin oligomerization
and connexon docking.[40] It is likely that the syndromic CX26 mutants co-oligomerize
not only with wild-type CX26 but also with other connexin proteins co-expressed in the
skin and exert a dominant- or transdominant-negative effect.[46, 63]
Goal and Organization of Thesis
Much has been learned about the physiology of normal hearing and the
pathophysiology of inherited hearing loss. Despite this progress, hearing aids and
cochlear implants remain the only habilitation options for hearing loss. Physicians,
researchers, and patients expect more effective therapeutics to be developed for hearing
16
loss. Gene therapy offers the potential to prevent or treat the unique pathologies
associated with different types of hereditary hearing loss. There are three essential
components to effective gene therapy: (1) a therapeutic substrate; (2) a route of delivery;
and (3) a vehicle for delivery. Potential therapeutic substrates include RNAi molecules,
genes, zinc finger nucleases, enzyme replacements, and drugs that alter biochemical
pathways. As researches come to better understand the molecular basis of inherited
disorders more effective therapeutic options will become available. Likewise, as new
methods to manipulate gene expression become available, new therapeutic strategies will
emerge for the inner ear.
The overall goal of my Thesis has been to apply the latest knowledge in molecular
pathology, gene therapeutics, and delivery methods to a mouse model of dominant
hereditary hearing loss. To develop gene therapies for humans it is necessary to first test
them in animal models. Understanding the animal model and its pathophysiology enables
a more rational design of therapeutic strategies. The specific aims of my Thesis are
described in the following five chapters:
Specific Aim 1: To further characterize the GJB2-R75W transgenic mouse model
of dominant-negative deafness. (Chapter 2)
Specific Aim 2: To design and validate siRNA for potent and specific knockdown
of the GJB2-R75W transgene (Chapter 3)
Specific Aim 3: To identify a viral vector suitable for safe and efficient gene
delivery to the supporting cells of the developing murine cochlea (Chapter 4)
Specific Aim 4: To deliver and assess efficacy of in vivo gene therapy for
dominant-negative GJB2-R75W deafness (Chapter 5)
17
Figure 1. Development of the murine inner ear. Lateral view of paintfilled-membranous
labyrinths ranging from E10.5 to E17.5 are shown. The spherical otocyst
develops at E9.5 from invagination of the otic placode. By E10.5, the otocyst
begins to change shape as a result of the formation of dorsal and ventral
protrusions that will eventually develop into the endolymphatic duct (ED) and
the cochlear duct (CD). By E12.5, the developing cochlear duct begins to form a
spiral and the proximal cochlea continues to lengthen. At E13.5, the developing
cochlea has completed three-quarters of a turn and by E17.5 has reached its
mature length of one and three-quarter turns. Otocyst microinjections are
performed between E11.5 and E12.5. White arrows indicate growth of the
proximal cochlea from E13.5 to E17.5. Adapted from Morsli et al. 1998 and
Kelley 2006.
18
Figure 2. Anatomy of the ear. The major structures of the eternal, middle, and inner ear
are depicted.
19
Figure 3. Diagram of a cross-section of the cochlea. Scala media (or cochlear duct, filled
with endolymph) lies between the scala tympani and scala vestibuli (filled with
perilymph). The base of scala media is formed by the osseous spiral lamina
(OSL) and the basilar membrane (BM). Resting on the basilar membrane is the
organ of Corti, which contains sensory inner hair cells (IH) and outer hair cells
(OH), separated by pillar cells (P) forming the tunnel of Corti. The inner hair
cell is synaptically connected to afferent dendrites (AD) of type I spiral
ganglion neurons. Efferent axons (EA) form contacts with outer hair cells. Each
outer hair cell is supported at its base by an outer phalangeal cell (OPh), or
Deiters cell. From each Deiters’ cell, a phalangeal projection extends upward to
the the reticular lamina, that forms the upper layer of the organ of Corti. The
apices of the outer hair cells are firmLy held by the reticular lamina, but the cell
bodies are suspended in fluid that fills the spaceof Nuel and the tunnel of Corti.
The composition of this fluid, called cortilymph, is thought to be similar to
perilymph. The inner hair cells are supported by the inner phalangeal cells
(IPh). The inner border cell (B) and cuboidal epithelial cells line the spiral
limbus on the inner sulcus (IS) side of the organ of Corti. The tectorial
membrane (TM) covers the hair cells and reaches the cells of Hensen (H). Two
other types of epithelial cells, the cells of Claudius (C) and Böttcher (B), cover
the outer sulcus. At the upper margin of the outer sulcus is the spiral
prominence (SP) followed by the stria vascularis (SV). Reissner’s membrane
(RM) stretches from the stria vascularis to the medial margin of the spiral
limbus. Endocochlear potential (EP) is the electrical potential difference
between endolymph and perilymph. Adapted from Schutz et al. 2010.
20
21
Figure 4. Supporting cells of the organ of Corti. Cross-sectional illustration showing the
cell types within the organ of Corti. A single row of inner hair cells (IHC) is
located on the medial side of the organ while three rows of outer hair cells
(OHCs) are located laterally. The sensory hair cells are surrounded and
supported by non-sensory supporting cells. The sub-types and location of
supporting cells are indicated by color. The fluid-filled tunnel of Corti and space
of Nuel are also indicated. Adapted from Driver and Kelley 2009.
22
Table 1. Autosomal dominant non-syndromic (DFNA3) and syndromic GJB2 mutations.
Mutation
cDNA level
Protein Domain
Disease
Protein level
c.131G>C
W44S
E1
DFNA3
c.132G>C
W44C
E1
DFNA3
c.172C>G
P58A
E1
DFNA3
c.223C>T
R75W
M2
DFNA3
c.224G>A
R75Q
M2
DFNA3
c.428G>A
R143Q
M3
DFNA3
c.487A>C
M163L
E2
DFNA3
c.535G>A
D179N
E2
DFNA3
c.551G>A
R184Q
E2
DFNA3
c.605G>T
C202F
M4
DFNA3
c.34G>C
G12R
NT
KID
c.42C>G
N14K
NT
Hypotrichosis-deafness
c.40A>C
N14Y
NT
KID
c.50C>T
S17F
NT
KID
c.119C>T
A40V
M1
KID & the follicular occlusion triad
125_127delAGG
E42del
M1
PPK w/deafness
c.134G>A
G45E
E1
KID
c.148G>A
D50N
E1
KID; hystrix-like ichthyosis-deafness
c.148G>T
D50Y
E1
KID
c.160A>C
N54H
E1
Bart-Pumphrey
N54K
E1
Bart-Pumphrey
c.175G>C
G59R
E1
PPK w/deafness
c.175G>A
G59S
E1
PPK w/deafness
c.176C>G
G59A
E1
PPK w/deafness
c.196G>C
D66H
E1
Vohwinkel
c.219A>G
H73R
M2
PPK w/deafness
c.223C>T
R75W
M2
PPK w/deafness
c.224G>A
R75Q
M2
PPK w/deafness
c.389G>T
G130V
CL
Vohwinkel
c.424T>C
F142L
M3
Mucositis-deafness
c.548C>T
S183F
E2
PPK w/deafness
23
Figure 5. Schematic diagram showing connexin topology and channel variety. (Left)
Model showing membrane topology for a connexin protein. M1-M4 represent
the four transmembrane domains, E1 and E the two extracellular loops, and CL
the cytoplasmic loop. (Right) Schematic drawing of possible arrangements of
connexons to form heterotypic and heteromeric channels. Adapted from Mese et
al. 2007.
24
Figure 6. Heteromeric connexon compositions. Schematic representation of the 14
possible connexon conformations in a cell co-expressing two connexin
isoforms.
25
Figure 7. Deafness- and skin disease-causing CX26 mutations. Topological map of point
mutations associated with non-syndromic deafness (blue) and syndromic
deafness with skin disease (red). Adapted from Laird 2006.
26
Figure 8. R75W plays a role in inter-connexin interactions and oligomerization. Cartoon
transparency side view (left) of two protomers with R75 side chains depicted in
magenta. Salt bridges around the R75W mutation site are indicted (right). The
R75W mutation is predicted to disrupt the salt bridges. Images courtesy of
Aaron Ver Heul (Department of Biochemistry, University of Iowa) and based
on crystal structure by Maeda et al. 2009.
27
CHAPTER 3: MOUSE MODEL OF DOMINANT-NEGATIVE
DEAFNESS
Introduction
The first step in designing gene therapy is selecting the appropriate animal model
for the disease of interest. The mouse has long been recognized as an important model
organism for studying the biology of hearing and deafness.[64] Unlike humans, cochlear
maturation and onset of hearing in rodents occurs postnatally. For example, hearing onset
in mice occurs on postnatal day 12 (P12).[65] This makes the mouse an excellent
resource for studying cochlear development and the physiologic, anatomic, and genetic
changes involved in hearing onset. In addition, the ability to target and manipulate the
mouse genome has brought the mouse to the forefront of model organisms for studying
deafness.
GJB2-R75W Transgenic Mouse
Kudo and colleagues created a mouse that expresses the dominant-negative
HsGJB2-R75W allele.[47] The GJB2-R75W transgenic mouse constitutively expresses
human GJB2-R75W and EGFP under the control of the CMV early enhancer/chicken β
actin (CAG) promoter (Table 2).[66] This mouse has severe-to-profound hearing loss by
two weeks of age, the age of hearing onset in mice. This phenotype makes the GJB2R75W transgenic mouse a suitable model for dominant congenital hearing loss.
Although the precise mechanism by which the GJB2-R75W allele prevents normal
hearing in humans is not definitively established, studies of the GJB2-R75W transgenic
mouse have provided valuable information about the pathologic consequences of this
mutation. The GJB2-R75W transgenic mouse never develops normal hearing. Auditory
brainstem response (ABR) testing shows that as wild-type mice mature and begin to
exhibit auditory responses approaching normal thresholds, the GJB2-R75W mice hearing
thresholds remain profoundly elevated at approximately 100 dB.[65] Inoshita and
28
colleagues studied the developmental morphology of GJB2-R75W mouse cochleae
compared to wild-type mouse cochleae and noted several important differences
including: (1) the tunnel of Corti, which is first seen at P5 in wild-type mice and is
required for normal hearing, fails to develop; (2) reduced number and organization of
microtubules in inner pillar cells; (3) failure of the organ of Corti to increase in height
during development as compared to wild-type cochleae; (4) swelling of the organ of Corti
supporting cells.[65] However, the inner and outer hair cells in GJB2-R75W mice appear
to develop normally through P12, as does Reissner’s membrane, stria vascularis, spiral
ligament, and spiral ganglion neurons. The endocochlear potential (EP), which begins to
be established by the stria vascularis at P5 and reaches mature levels by P17-18, was
within the normal range (80-100 mV) and did not significantly differ from wild-type
mice.[67] Despite morphologically normal inner and outer hair cells at two-weeks of age
and a normal EP, these transgenic mice are profoundly deaf.[68]
In summary, there appear to be two processes at play in the pathophysiology of
the GJB2-R75W transgenic mouse: First, the dominant-negative mutation prevents
normal development of the organ of Corti supporting cells. This may be the result of
failed intercellular transport of important signaling molecules and other growth factors
vital to early organ of Corti development. This transgenic mouse provides evidence that
proper supporting cell development is necessary for normal hearing. Second, although
inner and outer sensory hair cells appear to develop normally through P12, by 7 weeks of
age they have largely degenerated and disappeared.[47] The loss of sensory hair cells is
thought to result from disrupted cortilymph homeostasis, either through failure of the
endothelial gap junction network to properly recycle potassium (K+) and other ions or
from absence of important fluid compartments (e.g. tunnel of Corti and space of Nuel)
that are crowded out by swollen supporting cells.[47, 65]
In the paper first describing the GJB2-R75W transgenic mouse, Kudo and
colleagues suggested that it “would provide a valuable tool for investigating therapeutic
29
means to prevent the degeneration of supporting cells in the organ of Corti at an early
stage of development.”[47] Through collaboration, we obtained the GJB2-R75W
transgenic mouse with the ultimate goal of designing gene therapy for this deafness
model.
In order to design gene therapy with the highest likelihood of success, we
undertook further characterization of the transgenic mouse. This chapter discusses our
work to understand the genetic context of the disease. We investigated the genomic copy
number and tissue expression levels of the GJB2-R75W transgene to assess whether this
mouse appropriately reflects the human disease and is a suitable model for in vivo
application of RNAi-based therapy.
Materials and Methods
Animal Care and Breeding
All mice were maintained on a 12-hour light/dark cycle with access to food and
water ad libitum in the Medical Laboratories Animal Facility at the University of Iowa.
All animal-related procedures were performed in adherence with institutional and
national guidelines and were approved by the Institutional Animal Care and Use
Committee at the University of Iowa.
Upon initial receipt of the GJB2-R75W transgenic mice, we performed crosses
with the CBA strain. We subsequently backcrossed with the CBA/J strain (Jackson
Laboratory, Bar Harbor, ME) and selected for G/G homozygotes at the age-related
hearing loss locus (Cdh23). Mice homozygous for the non-susceptibility locus (G/G)
were crossed with each other, continuing with littermate intercrosses. Subsets of mice
continue to be backcrossed with CBA/J mice. These mice are currently at the F7
generation. GJB2-R75W transgenic mice have also been crossed with SKH1 (hairless)
mice (Charles River Laboratories, Kingston, NY), currently at F8 generation. We have
30
also crossed the GJB2-R75W transgenic mice with BALB/cJ mice (Jackson Laboratory),
currently at F2 generation.
Tissue Samples
Deeply anesthetized mice were sacrificed via cervical dislocation or decapitation
and desired tissue was removed for downstream applications: histological analysis, DNA
extraction, or RNA extraction.
Tissue Fixation and Paraffin Embedding
Cochleae
Inner ears were dissected and, after making a hole in the cochlear apex,
immediately fixed for two hours by immersion in 4% PFA, rinsed in PBS, decalcified in
EDTA and prepared for cryosectioning as previously described [69]. Briefly, decalcified
cochleae were washed with PBS and put through serial immersion with increasing
concentrations of sucrose in PBS, with final overnight immersion in 30% sucrose at 4°C.
Cochleae were then infiltrated with OCT and oriented for sectioning. 10 μm midmodiolar cryosections were sliced, mounted on slides, and stored at -80°C in preparation
for immunohistochemistry.
Skin
Skin was removed with scissors, fixed for two hours in 4% PFA, rinsed and stored
at 4°C in PBS overnight. Skin samples for cryosectioning were prepared in a similar
manner as the cochleae without the decalcification step. Skin samples for paraffinembedding and sectioning were dehydrated through graded alcohol and xylene followed
by embedding in paraffin. Paraffin-embedded samples were manually sectioned with a
microtome to sections of 6 µm thickness and mounted on glass microscope slides. After
de-paraffinization, sections were stained with hematoxylin and eosin (H&E) for
examination with light microscope.
31
Eyes
Eyes were removed and fixed in 2.5% gluteraldehyde in 0.1 M Na cacodylate for
16 hours, and post-fixed with 1% osmium tetroxide in 0.1 M Na cacodylate buffer at
room temperature for 1 hour. A series of acetone dehydrations were performed followed
by infiltration with Embed-812/DDSA/NMA/DMP-30 for 24 hours. 0.5-mm sections
were cut (EM UC6 ultramicrotome; Leica, Wetzler, Germany), and stained with 1%
toluidine blue. Sections were visualized and imaged using a light microscope (BX52;
Olympus, Tokyo, Japan) equipped with a digital camera (DP72; Olympus, Tokyo, Japan).
DNA Extraction
DNA for genotyping was extracted from mouse tail clips taken just prior to
weaning (P21). Genomic DNA of mice was extracted from tissue samples using DNeasy
Tissue kit (Qiagen Inc., Valencia, CA) according to manufacturer protocol. DNA was
eluted in 100µL of water, concentration measured by NanoDrop 2000 Spectrophotometer
(Thermo Scientific NanoDrop Products, Wilmington, DE), and diluted in water to a final
concentration of [20 ng/µL] in preparation for PCR. DNA for QPCR was subsequently
diluted by measuring concentration as the average of three NanoDrop readings and
diluting in water to [5 ng/µL].
RNA Extraction
Immediately following tissue collection total RNA was extracted using RNeasy
Fibrous Tissue Mini Kit (QIAGEN Inc., Valencia, CA) with on-column Dnase I digestion
(QIAGEN Inc., Valencia, CA) according to manufacturer protocol. Briefly, samples are
treated with proteinase K and debris is pelleted by centrifugation. Ethanol is then added
to the cleared lysate, and RNA is bound to the RNeasy silica membrane. Traces of DNA
that may copurify are removed by a DNase treatment on the RNeasy spin column. DNase
and any contaminants are washed away. RNA is eluted in an appropriate volume of
RNase-free water (30 to 100 µL, depending on tissue) and stored at -80°C.
32
Quantitative Real-Time PCR (QPCR)
RNA from mouse tissue was reverse transcribed in triplicate to yield [10ng/µL]
cDNA (heart and testis samples yielded [5ng/µL] cDNA). 20ng of cDNA (10ng for heart
and kidney) from each tissue was assayed in triplicate with QRT-PCR probing for
HsGJB2-R75W and MmGjb2. Serial dilutions of known quantities of linearized plasmids
(ranging from 101 molecules to 107 molecules per µL) were used as standards to
quantitate the experimental samples. The reaction conditions and quantitative analysis
methods are described in the next chapter. In total, nine separate quantities were
generated for each tissue (three reverse transcription replicates each assayed in triplicate
QRT-PCR reactions) and the averages and standard deviations calculated for each tissue
were calculated for copy number per nanogram of cDNA.
Genotyping
GJB2-R75W Transgenic Mice
The GJB2-R75W transgenic construct is composed of the CAG promoter
followed by the cDNA sequence (681 bases) for human GJB2 containing the single base
substitution c.223C>T which results in the p.Arg75Trp amino acid substitution. This
sequence is followed immediately by an internal ribosomal entry site (IRES) sequence
and the enhanced green fluorescent protein (EGFP) sequence.[47] To identify mice
containing this transgene, we designed primers to amplify a 204 base segment within the
CAG promoter (Table 2). Each 25 µL PCR reaction contained: 2.5 µL of 10X reaction
buffer (Bioline, Taunton, MA), 2.0 µL dNTPs, 0.5 µL forward primer (20 µM), 0.5 µL
reverse primer (20 µM), 0.75 µL MgCl2, 5.0 µL 5x betaine (Charles Searby, Sheffield
Lab, University of Iowa), 12.6 µL dH2O, 0.15 µL Taq DNA polymerase (BIOLASE™;
Bioline, Taunton, MA), and 1.0 µL gDNA (20 ng/µL). PCR conditions were: 94°C for 4
minutes, 35 cycles of (94°C for 30 seconds, 55°C for 30 seconds, 72°C for 30 seconds),
and 72°C for 7 minutes. PCR products were analyzed on a 1% agarose gel using EtBr
33
detection. The 204 base product is only present in reactions that carry the transgene
DNA.
We use Sanger sequencing to verify the presence of the c.223C>T mutation in the
GJB2-R75W transgenic mice. For sequence confirmation we selected primers that
amplify a 279 base region of human GJB2 that includes the c.223C>T mutation (Table
2). Each 25 µL PCR reaction contained: 2.5 µL of 10X reaction buffer (Bioline, Taunton,
MA), 2.0 µL dNTPs, 0.5 µL forward primer (20 µM), 0.5 µL reverse primer (20 µM),
0.75 µL MgCl2, 17.6 µL dH2O, 0.15 µL Taq DNA polymerase (BIOLASE™; Bioline,
Taunton, MA), and 1.0 µL gDNA (20 ng/µL). PCR conditions were: 94°C for 4 minutes,
35 cycles of (94°C for 30 seconds, 56°C for 30 seconds, 72°C for 30 seconds), and 72°C
for 7 minutes. Because the primers also amplify some endogenous mouse Gjb2, the PCR
products are sequenced using the forward and reverse primers in separate sequencing
reactions to verify the presence of the c.223C>T mutation.
Age-Related Hearing Loss (Cdh23) Locus
An allelic variant of the Cdh23 gene confers susceptibility to age-related hearing
loss. A restriction digest assay allows accurate discriminate between the two alleles. Each
25 µL PCR reaction contained: 2.5 µL of 10X reaction buffer (Bioline, Taunton, MA),
2.5 µL dNTPs, 0.5 µL forward primer (20 µM), 0.5 µL reverse primer (20 µM), 0.5 µL
MgCl2, 17.0 µL dH2O, 0.5 µL Taq DNA polymerase (BIOLASE™; Bioline, Taunton,
MA), and 1.0 µL gDNA (20 ng/µL). PCR conditions were: 95°C for 3 minutes, 35 cycles
of (94°C for 30 seconds, 55°C for 30 seconds, 72°C for 30 seconds), and 72°C for 10
minutes. 10 µL of PCR product is analyzed on a 1% agarose gel using EtBr detection to
confirm target amplification (230 base product). After confirmation of amplification, the
remaining 10-15ul of PCR product was directly digested with the addition of 0.5 µL
MspI restriction enzyme (New England Biolabs, Ipswich, MA) and incubation for 1 hour
at 37°C. Digestion product is then analyzed on a 2% agarose gel using EtBr detection.
34
Mice homozygous for the age-related hearing loss allele (A/A) will show a single 230
base band; mice homozygous for the non-age-related hearing loss allele (G/G) will show
two bands—166 bases and 62 bases (indicated complete cleavage of the original);
heterozygous mice will show all three bands—230, 166, and 62 bases.
Results
Hearing Loss
Upon receipt of the “mixed C57BL” background GJB2-R75W mice, we crossed
them with CBA/J mice and maintained them in the hemizygous state through backcrosses
according to standard laboratory protocol. The CBA/J inbred strain is commonly used in
hearing loss research because they do not carry the age-related hearing loss variant of
Cdh23.[70] I designed a PCR and restriction digest assay to determine genotype at this
locus. While age-related hearing loss is a confounding factor for mouse models of lateonset or progressive hearing loss, it is not a consideration for mice with congenital and
non-progressive hearing loss. Therefore, in studies which will be described later, I was
not concerned with the Cdh23 genotype of GJB2-R75W mutant mice produced by
intercrosses with other strains (e.g. SKH1, C57BL/6, and BALB/c). In addition, I control
for effects of age on hearing loss in this study by auditory testing before mice reach 4
months of age, an age prior to onset of age-related hearing loss in most C57BL/6 and
BALB/c mice.[70] Most of the hearing tests on the GJB2-R75W transgenic mice were
performed between 3 and 10 weeks of age.
Using auditory brainstem response (ABR) testing, I verified the previously
reported hearing loss in the GJB2-R75W mice.[47] Hearing onset in mice occurs
approximately at age P12 and we are able to perform ABR at 2 weeks of age. The clickstimulus ABR threshold for mice with the GJB2-R75W transgene (Tg+) is 88.6 ± 10.7
dB compared to 45.3 ± 5.6 dB for non-transgenic (Tg-) littermates (n = 60; p = 1.4x10-53)
(Figure 10). The hearing thresholds ranged from a low of 75 dB to a high of 130 dB, the
35
highest sound level testable by our system. Through sibling crosses we also generated
litters containing mice homozygous for the transgene, but we did not observe a
correlation between genotype and degree of hearing loss (data not shown). Therefore, we
conclude that GJB2-R75W transgenic mice have profound hearing loss at the earliest age
at which testing is possible. The hearing loss phenotype is fully penetrant. Variability in
hearing thresholds exists, but the degree of hearing loss is independent of whether the
mice are hemizygous or homozygous. The hearing loss is not progressive during the first
4 months of life (data not shown).
Transgene Copy Number Analysis
In transgenic mice created by pronuclear microinjection, the transgene integrates
randomLy into the genome, often as multiple head-to-tail concatamers.[71] Kudo and
colleagues did not report the integration site or the transgene copy number in the GJB2R75W mouse. Since transgene copy number influences the expression level of the
transgene which has implications for therapeutics we sought to determine the number of
copies integrated into the transgenic mouse genome. We used quantitative real-time PCR
(QPCR) on genomic DNA (gDNA) extracted from transgenic mouse tails to estimate the
copy number of the GJB2-R75W transgene.[72]
gDNA extracted from tails of 5 female and 5 male mice hemizygous for the
GJB2-R75W transgene was amplified in triplicate by qPCR. The qPCR assay is validated
to specifically amplify and quantify endogenous murine Gjb2 and the GJB2-R75W
transgene. The average number of copies of MmGjb2 per nanogram of DNA is 205.9 ±
70.9 and the average number of copies of HsGJB2-R75W transgene is 72.1 ± 26.7
copies/ng of gDNA. The ratio of endogenous MmGjb2 to transgene is 2.9 ± 0.3 to 1
(Figure 11). This approximates a single copy of the transgene per diploid genome.
To validate our QPCR assay for determining relative copy number, I quantified
copies of endogenous Gjb2 relative to the murine androgen receptor gene (Ar). Ar is
36
located on the X chromosome and so female mice have twice as many copies of the gene
as male mice (two copies versus one copy). Therefore, we would expect the results of a
QPCR assay to show that the ratio of Gjb2:Ar in male mice is twice the ratio of Gjb2:Ar
in female mice. The actual QPCR assay showed a Gjb2:Ar ratio of 4.0 ± 0.5 : 1 for male
mice (n=27) and 2.1 ± 0.7 : 1 for female mice (n=26) (Figure 11). These data are
consistent with the expectation that male mice, with half the number of X chromosomes
compared with females, would have twice the number of Ar gene copies relative to Gjb2
gene copies as the female mice. This provided validity to our assay. Our data is consistent
with the presence of a single copy of the transgene per diploid genome in the GJB2R75W transgenic mice.
Tissue Expression of GJB2-R75W Transgene
The expression level of the GJB2-R75W transgene compared to endogenous Gjb2
is the critical factor in predicting therapeutic potential. GJB2-R75W transgene expression
is driven by the CAG promoter. This promoter is highly active in pre-implantation
embryos and therefore achieves widespread expression throughout the mouse. The CAG
promoter has been reported to drive strong transgene expression in the brain, heart,
muscle, pancreas, and skin.[73] We used QRT-PCR to determine the relative expression
levels of the GJB2-R75W transgene in comparison with endogenous MmGjb2 expressed
in various tissues of the transgenic mice. GJB2-R75W mouse was sacrificed at age P3
and the following tissues were harvested: bladder, brain, cochlea, eyes, heart, kidney
liver, lung, skeletal muscle, skin and testis. The cochlear tissue was further dissected to
isolate the sensory epithelium and the lateral wall, the regions of the inner ear which
endogenously express Gjb2. RNA was extracted, reverse transcribed and assayed with
QRT-PCR As expected, the transgene was expressed in each tissue assayed: bladder
(4989.5 ± 1006.5), brain (2536.7 ± 1006.5), cochlea (437.8 ± 146.2), eye (2924.5 ±
303.1), heart (77547.1 ± 15803.5), kidney (3198.0 ± 436.9), liver (344.9 ± 51.2), lung
37
(4871.3 ± 449.6), skeletal muscle (4221.2 ± 504.7), skin (2871.6 ± 173.8), and testis
(15240.7 ± 1789.1) (Figure 11). We found that Gjb2 was endogenously expressed in the
brain, cochlea, eye, kidney, liver and skin. Expression of endogenous Gjb2 was
negligible (<10 copies/ng cDNA) in the bladder, heart, lung, skeletal muscle and testis.
Based on the hearing loss and skin disease phenotypes associated with the R75W
mutation in humans, the most relevant data is the level of expression of transgene to
endogenous MmGjb2 expression in the cochlea and in the skin. In one transgenic mouse,
we found the relative expression level of transgene:Gjb2 in the cochlea was 1.0:1 and the
relative expression level in the skin was 3.0:1.We then assayed several more sets of tissue
isolated from additional transgenic mouse cochleae. Results from one mouse showed a
transgene:Gjb2 expression of 1.6:1 and results from a pooled set of three transgenic mice
cochleae showed a ratio of 1.8:1. Combining the data from all QRT-PCR assays of
transgenic mouse cochleae (5 cochleae and 27 QRT-PCR data points), gave an average
expression level of 1710.8 ± 969.5 for the transgene and 1067.8 ± 669.0 for endogenous
Gjb2. Our calculations show that the overall transgene:Gjb2 expression for the inner ear
(sensory epithelium and lateral wall) of GJB2-R75W transgenic mice is 1.6:1, suggesting
that the CAG drives expression three times mores robustly than the endogenous Gjb2
promoter.
Skin Disease
A majority of the known dominant mutations in GJB2 lead to forms of syndromic
deafness with distinct skin manifestations in humans (Table 1). The R75W mutation can
cause palmoplantar keratoderma (PPK) with deafness (MIM#148350). PPK consists of
thickening of the skin on the palms and soles, which leads to cracking and sloughing of
skin and constriction bands around the digits (pseudoainhum). In the most severe cases,
affected persons may lose portions of digits secondary to auto-amputation (Figure 12).
The GJB2-R75W transgenic mice have not been reported to have a skin phenotype;
38
however based on the human phenotype we studied the mice in detail to determine skin
pathology was present.
Presuming that severity of any skin phenotypes would increase with age, we
initially undertook gross examination of the skin of adult mice. The presence of hair
complicates evaluation of any skin defects and no obvious defects were noted in adult
mice. However, mice are born hairless and the skin over the entire body is readily visible
for the first 7 to 10 days of life. I began examining the skin of transgenic mouse litters
from the time of birth and noted that some of the mice had scaly-appearing skin
compared to littermate controls that had smooth, non-scaly skin (Figure 13). Genotyping
showed that scaly-skinned mice carried the transgene, while non-scaly-skinned mice did
not. The most severe scales were generally localized to the backside (lower back and base
of tail) and on the tails. Additionally, the tails of some of the mice had constriction bands
reminiscent of pseudoainhum described in some patients with dominant GJB2 mutations.
In most of the mice, the grossly visible skin manifestations were no longer noticeable
after the animals grew hair. Several mice, however, had extreme skin phenotypes
including several mice with constriction bands so severe that auto-amputation of the tail
occurred (Figure 13). A single mouse had a constriction band which lead to autoamputation of a toe.
Prompted by the gross skin manifestations, we analyzed skin histology in
transgenic mice compared to wild-type littermates. We sacrificed two litters of pups at
age P6 and P7 and removed skin from the most severely affected areas on the back and
the tail of transgenic mice and non-transgenic controls. The skin was fixed, paraffinembedded and H&E stained as described in the Methods section in preparation for
histologic analysis. Dr. David Meyerholz (Department of Pathology, University of Iowa)
analyzed the skin sections for histologic evidence of pathology while blinded to
genotype . The skin of the mutant mice showed signs of acanthosis (epidermal
proliferation), orthokeratotic hyperkeratosis, inflammation, vascular congestion and
39
variable changes consistent with necrosis. The annular constrictions of the tail are
consistent with a non-specific degenerative condition in rodents called “ringtail.” None of
these pathologies was observed in the non-transgenic skin sections. While these skin
manifestations are non-specific and from histology alone do not definitely reflect a
genetic etiology, these changes were found only in mutant mice, consistent with a causeand-effect relationship.
Since the presence of hair makes skin evaluation more difficult, we began
breeding the GJB2-R75W transgene onto a hairless mouse background. We have been
backcrossing GJB2-R75W transgenic mice onto the SKH1 mouse strain (Crl:SKH1-hr,
Charles River Laboratories, Wilmington, MA). These albino, hairless mice are euthymic
and immunocompetent. They grow hair normally as pups but then start to lose it at about
age P14. The hair loss begins at the head and proceeds toward the tail until it is
completely gone by about age P25. The hair loss is due to a hypomorphic mutation called
hr in the hairless gene (Hr).[74] Hr encodes a transcriptional co-repressor protein, Hr.
Homozygosity for hr results in altered transcription of genes that play a role in
keratinocyte differentiation. The result is loss of hair in the SKH1 mouse. We are
currently at the F8 generation of creating a congenic GJB2-R75W mouse on the SKH1
background.
Cataracts
In the course of regularly working with and observing the GJB2-R75W transgenic
mice we noticed another previously unreported phenotype: cataracts (Figure 14). A
cataract is when the normally clear lens becomes cloudy or opaque. Gross examination
showed that some of the mice had a white patch in the center of their eyes, a phenotype
that segregated with the transgene. To further investigate this new phenotype and
determine whether it represented a true cataract, Dr. Geoffrey Lively (Department of
Molecular Physiology and Biophysics, University of Iowa) performed slit-lamp
40
examinations on 56 transgenic mice and 36 non-littermates ranging in age from 3 weeks
to approximately 1 year. 55 of the 56 transgenic mice had cataracts while none of the
non-transgenic mice showed signs of cataracts, confirming that expression of the GJB2R75W transgene in the lens tissue causes cataract formation in mice. Subsequent
observations indicate that cataracts develop as early as P14 in GJB2-R75W transgenic
mice.
To examine the lens pathology, Adam Hedberg-Buenz (Department of Molecular
Physiology and Biophysics, University of Iowa) removed the eyes of transgenic mice and
processed them for histologic analysis as described in the Methods section. The sections
showed severe nuclear cataracts with cellular disorganization and vacuoles or enlarged
intercellular spaces on the periphery (Figure 15). Both the gross and histologic
appearance of the transgenic mouse lenses are consistent with severe cataracts. It is
interesting to note that the transgene is expressed in the lens, but that endogenous Gjb2 is
not expressed in the lens.
Discussion
We obtained GJB2-R75W transgenic mice with the goal of responding to the call
of Kudo and colleagues to use this mouse as “a valuable tool for investigating therapeutic
means to prevent” connexin-related dominant-negative hearing loss.[47] Gene therapy
holds tremendous promise, but there are also many obstacles that must be overcome if
this promise is to be realized.
Deafness due to the GJB2 p.Arg75Trp mutation is caused by a dominant-negative
mechanism. Carriers of null mutations in GJB2 (e.g. 35delG) have normal hearing.
Unfortunately, patients who are heterozygous for the p.Arg75Trp mutation have hearing
loss despite the presence of one normal copy of the gene. GJB2-R75W deafness, then, is
not a problem of connexin quantity, but rather a problem of connexin quality. The
41
dominant-negative CX26 protein prevents the wild-type connexins from forming
functional gap junction channels.
Transgenes integrate randomly into the genome, often as multiple head-to-tail
concatamers.[71] Because transgene copy number can influence the level of expression,
we measured copy number. by quantitative real-time PCR (qPCR).[72] We designed and
validated a TaqMan qPCR assay to discriminate between the GJB2-R75W transgene and
endogenous murine Gjb2.[75], which allowed us to quantify the number of molecules of
both the transgene and the endogenous gene in a single multiplex reaction (see Tables 3
and 4). qPCR results from gDNA of transgenic mice indicate 0.7 ± 0.1 transgene copies
per diploid genome. This value is consistent with a single copy of the transgene in each
GJB2-R75W mouse.
The ideal animal model for a genetic disease would not only mirror the human
phenotype, but would also approximate other important aspects such as expression level
and expression location of the mutant allele. Humans with the R75W mutation are
heterozygotes and are expected to have relatively equal expression of each allele. The
presence of a single transgene copy per mouse approximates human patients, but
transgenes are inherently different from the natural state. The expression of a gene
ultimately determines its biologic influence and expression is determined by several
factors. One of the chief determinants of gene expression is the promoter. The GJB2R75W transgene is driven by the engineered CAG promoter, a strong and ubiquitous
promoter that leads to high levels of transgene expression in many tissues.[73] The
expression level of the transgene compared to the expression level of endogenous Gjb2
will be a key indicator of whether gene therapy is feasible in this mouse model.
Kudo and colleagues assayed expression data by semi-quantitative reverse
transcription PCR and reported a 2.9 to 1 ratio of transgene mRNA to Gjb2 mRNA in the
mouse cochlea.[47] Because expression data is central to our therapeutic strategy, we
decided to complete our own expression analysis using reverse transcription real-time
42
quantitative PCR (qPCR). We found that the transgene is expressed in every tissue
assayed: bladder, brain, cochlea, eye, heart, kidney, liver, lung, skeletal muscle, skin, and
testis. Gjb2 is endogenously expressed in the brain, cochlea, eye, kidney, liver and skin. It
is apparent, therefore, that the transgenic mice express dominant-negative CX26 even in
tissues that do not normally express CX26.
The cochlea is the target tissue for deafness therapy. Despite its small size, it is an
extremely complex structure consisting of a bony capsule enclosing two main fluid-filled
compartments separated by membranes and containing dozens of different cell types (see
Figure 16). CX26 and CX30 proteins are localized to supporting cells in the organ of
Corti and fibrocytes of the lateral wall. We found the ratio of transgene mRNA to
endogenous Gjb2 mRNA to be 1.6 to 1.
Our finding of a 1.6:1 expression ratio is approximately half as much as reported
by Kudo and colleagues.[47] This discrepancy likely reflects the assays used. Kudo and
colleagues used a semi-quantitative method while we used a real-time assay with high
quantitative sensitivity and specificity.[76] In addition, Kudo and colleagues assayed the
whole cochlea, while we studied the organ of Corti and lateral wall, two a relatively small
components of the whole cochlea. Based on the superior reliability of the RT-qPCR
technique and the specific selection of tissue used in our assay, we are confident in our
conclusion that there is less than twice the level of transgene mRNA compared with
endogenous Gjb2 mRNA in the critical regions of the transgenic mouse cochlea.
Although this ratio is greater than the 1:1 ratio expected in human patients, it is within a
level that could reasonably be reached with therapeutics.
The primary phenotype of the GJB2-R75W mouse is severe to profound hearing
loss. Connexin isoforms are expressed in virtually every tissue and connexin mutations
cause an array of pleomorphic diseases including.[43, 77] We have identified two
previously unrecognized aspects of the phenotype in the GJB2-R75W mouse: skin
disease and cataracts.
43
Skin
Although Kudo and colleagues failed to note any phenotypes other than hearing
loss, we undertook a gross examination of the transgenic mouse skin to ascertain whether
there is evidence of skin disease similar to patients with palmoplantar keratoderma (PPK)
and deafness (MIM#148350). The presence of hair makes skin examination difficult and
so we began regularly examining the skin on pups younger than P7. We noticed that
some mice had more scales on their skin than their littermates, and some had constriction
bands on the tail (Figure 13). The most severe-looking skin phenotypes always correlated
with presence of the transgene (data not shown).
There are several explanations for why the skin phenotype has not been
previously reported. First, the GJB2-R75W transgenic mice were specifically created to
exhibit a hearing loss phenotype and hearing analysis cannot be undertaken in mice until
the external auditory canal opens and the inner ear has matured at around P12-14. By this
time the pups are covered with hair and the scaly skin cannot be seen. Second, the skin
phenotype incompletely penetrates and shows variable expressivity. In fact, a majority of
the transgenic mice did not show gross skin abnormalities and, of those with skin
manifestations, the severity ranged from mild scaling to auto-amputation of the tail
(Figure 13). After more than 4 years of work with hundreds of transgenic mice, we have
only observed a total of 5 mice with auto-amputation of the tail. Variations in penetrance
and expressivity suggest the presence of genetic modifiers.[78] A valuable future study
would seek to identify genetic modifiers of the GJB2-R75W transgene in mouse skin.
This could be particularly relevant because genetic modifiers are also likely to play a role
in the incomplete penetrance of the PPK phenotype in humans with the p.Arg75Trp
mutation and in the general variability of phenotypes associated with mutations in
GJB2.[77, 79]
Third, the skin disease is most severe in mice on a mixed background. The GJB2R75W mice that we received from our collaborators were mainly on a C57BL/6
44
background (generation unknown).[47] It was not until we began breeding the mice onto
different backgrounds that we first observed the skin phenotype. Severe skin
manifestations have not been noted in transgenic mice that are beyond the fourth or fifth
generation of backcrossing to an inbred strain. Each of the mice with severe autoamputation of the tail has been within several generations of backcrosses with CBA,
C57BL/6 or SKH1. The idea that a mixed background is most likely to provide for the
widest range of phenotypes in transgenic mice was recently suggested by Dr. Thomas
Doetschman.[80] He suggests the use of mixed background for initial detection of
phenotypes, some of which can be lost on an inbred background. After the phenotypes
have been identified, breeding onto inbred strains can help determine which phenotypes
have modifiers in each of the background strains.[80]
Our observations suggest that the transgenic mice have a PPK-like skin phenotype
and are a model for syndromic PPK with deafness. After consultation with Dr. Mary
Stone (Department of Dermatology, University of Iowa) and Dr. David Meyerholz
(Department of Pathology, University of Iowa) paraffin-embedded H&E sections of skin
were prepared for histologic evaluation. The skin sections were taken from the most
severely affected areas and control skin samples were taken from similar locations on
non-transgenic littermates. Blinded to the mouse genotype, Dr. Meyerholz analyzed the
skin sections and found hyperplastic and hyperkeratotic patterns of dermatitis.
The pathologic skin processes in the mice are non-specific and may be consistent
with a reaction to some environmental stimulus. Ringtail is a general process that rodents
become more susceptible to when exposed to low levels of humidity or high
temperatures. Because we have only observed a small number of animals with this severe
phenotype we cannot rule out that these are sporadic cases unrelated to the genotype.
However, the fact that each mouse observed with auto-amputation have also been
positive for the transgene suggests a cause-and-effect role for the dominant-negative
connexin mutation in the etiology of the skin disease.
45
In addition to genetic modifiers, phenotypic variability may be due to
environmental factors.[43] It is possible, for example, that the dominant-negative
connexin predisposes mice to skin disease and by enhancing sensitivity to environmental
insults such as humidity, temperature, bacteria and other irritants. This idea could be
tested in the future by exposing the skin to a mild irritant and assessing for hyper-reactive
responses in the transgenic mice compared to wild-type. Future histologic analysis of the
mouse skin might also include immunohistochemistry to assess for differences between
transgenic and wild-type mice. Potential investigations might include: (i) proliferation
assays (e.g. Ki-67, PCNA, Cyclin-D, etc.) for epidermal and follicular epithelia; (ii)
inflammation analysis, including mast cell (e.g. ABPY and Giemsa stains),
polymorphonuclear cell (e.g. Myeloperoxidase) and eosinophil (e.g. Congo red)
enumeration; (iii) additional special stains for bacterial or fungal organisms on the
surface of the skin.
To date, two other Cx26 mutant mice have been reported with skin disease.[81,
82] Bakirtzis and colleagues created a transgenic mouse with targeted epidermal
expression (keratin 10 promoter) of the p.Asp66His mutation. This dominant GJB2
mutation causes Vohwinkel syndrome (MIM#124500), a dominant congenital deafness
syndrome with diffuse mutilating keratoderma.[81] The K10Connexin 26(D66H) mouse
manifests Vohwinkel-like epidermal thickening and premature apoptosis of
keratinocytes.[81] Recently, Schutz and colleagues created a conditional knock-in mouse
model of keratitis-ichthyosis-deafness (KID) syndrome (MIM#148210).[82] In humans,
KID syndrome is characterized by keratitis and chronic progressive corneal
neovascularization, skin hyperplasia, sensorineural hearing loss and increased
carcinogenic potential. In the Cx26S17F mouse model Gjb2 is replaced by the
p.Ser17Phe mutation expressed under control of the endogenous Gjb2 promoter. The
phenotypes for this mouse include hearing loss, epidermal hyperplasia, and annular
constriction bands on the tail.[82] The skin phenotypes we observed in the GJB2-R75W
46
transgenic mouse are similar to those reported in these other models of dominant CX26
skin disease, reinforcing the genetic etiology of the skin phenotype.
The precise mechanism for how mutations in CX26 cause skin disease is not
clear. However, pathologic skin manifestations are only associated with dominant gainof-function CX26 mutants and not with recessive loss-of-function mutations. This
suggests that CX26 mutants associated with skin disorders likely interact with other
genes in a manner that results in disease. Since connexin isoforms interact to form
channels and multiple isoforms are expressed in the skin, these genes are good candidates
for trans-pathological effects of CX26 mutations. Rouan and colleagues showed that
wild-type CX26 could form functional heteromeric channels with wild-type CX43, skin
pathology-inducing CX26 mutants exerted a trans-dominant-negative effect on wild-type
CX43, significantly reducing channel activity.[83] The authors also showed colocalization of a dominant CX26 mutant (p.E42del) with CX43 in lesional skin from a
patient.[83] The data support the hypothesis that skin disease-associated mutations in
CX26 alter the function of proteins in the skin that are important for maintenance of
homeostasis and proliferative balance.[77]
Skin manifestations in human PPK with deafness patients are, as the name
indicates, limited to the hands and the feet. The skin phenotypes in the mouse models,
however, are more diffuse. This difference is due to species specific variation in temporal
and spatial connexin expression. In humans, GJB2 is expressed in very low levels in the
basal layer of skin, but at high levels throughout the hairless skin of the palms and the
soles (i.e. glabrous skin).[43] Gjb2 is expressed diffusely in embryonic mouse skin, but is
not strongly expressed in adult mouse skin.[84] This expression pattern may provide an
additional explanation for failure to observe skin disease in adult transgenic mice.
Due to difficulties in effective RNAi delivery to the skin and our initial goal of
treating hearing loss, we decided to focus our therapeutic efforts on the cochlea.
Recently, however, significant progress has been made in the delivery of siRNA to the
47
skin which has led to therapeutic trials for a rare genetic skin disorder, pachyonychia
congenital (MIM#148041), which also manifests with focal PPK.[85-87] When current
techniques for siRNA delivery to the skin are optimized and proven safe and effective,
dominant mutations in CX26 that lead to skin disease will make excellent candidates for
RNAi-based therapy.
Cataracts
Hearing loss and skin disease are the only phenotypes reported in humans with
the p.Arg75Trp mutation in GJB2. However, during our routine work with the GJB2R75W transgenic mice we observed an additional phenotype: cataracts. Cataract refers to
opacity of the lens. We noticed that some of the mice had distinct white spots in the
center of the eye as early as 3 weeks of age. These white spots were bilateral and
segregated with the presence of the transgene. Dr. Geoffrey Lively (Department of
Molecular Physiology and Biophysics, University of Iowa) was masked to the genotype
and performed binocular slit-lamp examination on a total of 92 mice. He found that 55/56
of the transgenic mice had bilateral cataracts while none of the wild-type mice had
cataracts. The lone transgenic mouse without cataracts was 3 weeks old at the time of the
slit-lamp examination and subsequently lost to follow-up so we do not know whether it
developed cataracts at a later date. Our data and continued observations indicate,
however, that the presence of early cataracts segregates perfectly with the presence of the
GJB2-R75W transgene. In fact, we now use cataracts as a visual marker to indicate
presence of the transgene for routine colony maintenance.
With the help of Adam Hedberg-Buenz (Department of Molecular Physiology and
Biophysics, University of Iowa), we analyzed toluidine blue-stained sections of
transgenic lenses. The nuclear cataracts are extremely severe, extending from the center
and filling 75-80% of the entire lens. The differentiating fiber cells on the periphery are
swollen and completely lack their characteristic organization.
48
The lens expresses three different connexins: CX43 (GJA1), CX46 (GJA3), and
CX50 (GJA8). Mutations in CX46 and CX50 have been described in patients affected by
cataract.[88] Likewise, there are a number of mouse models of cataracts caused by
mutations in Cx46 and Cx50.[89] Connexin-mediated communication plays a role in lens
growth and is necessary for maintaining lens transparency.[77] Connexins participate in
the internal circulating current system of the lens by allowing Na+ and Ca2+ ions to move
along their electrochemical gradient from central fibers to the lens surface. This microcirculatory carries nutrients into and removes metabolic waste from the avascular lens
tissue.[43] Loss of this system leads precipitation of crystallin proteins and cataract
formation.
CX26 is not naturally expressed in either mice or human lens tissue.[43]
However, driven by the CAG promoter, the GJB2-R75W transgene is expressed in most
tissues of the transgenic mice, including the lens. As expected, we found high levels of
transgene mRNA in the transgenic lens tissue and confirmed that wild-type Gjb2 is not
expressed in the mouse lens. Cataracts in the GJB2-R75W transgenic mice are most
likely the result of the transgene exerting a trans-dominant effect on endogenous Cx46
and/or Cx50 in the lens, showing that Cx26 interacts with both Cx46 and Cx50 to form
gap junction channels.[90] It is likely that the dominant-negative CX26 mutant interacts
with endogenous lens connexins to prevent the formation of a functional gap junction
system, just as it does with Cx30 and Cx26 in the cochlea. We did not further evaluate
the lens connexins or cataracts in the transgenic mice. The GJB2-R75W mouse is not a
model for any known eye disease because it is the result of a misexpressed mutant gene.
However, this cataract model could be useful for researchers who are interested in further
understanding the roles of connexins in the lens and the development of severe cataracts.
It is also possible that misexpression of CX26 or related genes in humans could play a
role in human disease phenotypes, such as cataracts and dermatological disease. Ectopic
expression of genes is known to result in abnormal phenotypes. For example, the yellow
49
mouse obesity syndrome is due to dominant mutations at the Agouti locus, which is
characterized by obesity, hyperinsulinemia, insulin resistance, hyperglycemia,
hyperleptinemia, increased linear growth, and yellow coat color. This syndrome is caused
by ectopic expression of Agouti in multiple tissues.[91]
The observation of a phenotype (cataracts) in a tissue (lens) where CX26 is not
naturally expressed begs the question: Could the GJB2-R75W transgenic mouse be a
model for widespread connexin dysfunction? This question provides opportunity for
future investigations. Our efforts have been focused on gathering information useful in
the development of gene therapy for hearing loss and so we have not performed rigorous
multi-system phenotyping screens on the mice. Since there are many connexin types that
do not interact with the mutant CX26 it is not likely that the CX26-R75W mutant protein
will have an inhibitory effect on every wild-type connexin type. But, because of the
presence of connexins throughout the body, there is the possibility for dominant-negative
interactions which may lead to pathology in many tissues where gap junctions play an
important role. The GJB2-R75W transgene is expressed in the heart and the brain, two
tissues in which connexin mutants are known to lead to human pathology. CX40, CX43
and CX45 are expressed in the heart and mutations in these have been linked with
arrhythmias, structural defects, and predispose to coronary artery disease.[43] In the
brain, connexin mutations may play a role in certain forms of epilepsy.[43] Thus, we
suggest the brain and the heart as strong candidate tissues for examination of new
phenotypes in the GJB2-R75W transgenic mouse.
In Summary, we have confirmed the previously reported hearing loss phenotype
in the GJB2-R75W transgenic mouse. We have performed quantitative assays to
determine transgene copy number in the mice as well as expression analysis in a variety
of tissues. The 1.5 to 1 (transgene to Gjb2) expression in the cochlea is significantly less
than was previously reported. As such, the molecular pathology in the GJB2-R75W
transgenic mouse cochlea approximates that of humans with the p.Arg75Trp mutation in
50
GJB2. Furthermore, the pathology associated with the GJB2-R75W mutation is located in
the supporting cells and prevents proper postnatal development of the organ of Corti. It
is clear that the loss of sensory hair cells is secondary to supporting cell dysfunction. This
information provides a specific therapeutic target: organ of Corti supporting cells. We
concluded that the GJB2-R75W transgenic mouse is a model of dominant deafness and
skin disease and is a promising target for cochlear genetic therapies. As we design
therapeutics for this mouse, we used the genetic and phenotypic details to guide our
selection of therapeutic, delivery method, and delivery vector.
51
Table 2. Primers used for genotyping mice.
52
Figure 9. Construct of the GJB2-R75W transgene. Adapted from Kudo et al. 2003.
53
Figure 10. Hearing loss associated with the GJB2-R75W transgene. GJB2-R75W
transgenic mice have significantly increased hearing thresholds compared to
non-transgenic littermates when measured by click stimulus auditory
brainstem response testing. The click-stimulus ABR threshold for mice with
the GJB2-R75W transgene (Tg+) is 88.6 ± 10.7 dB compared to 45.3 ± 5.6 dB
for non-transgenic (Tg-) littermates (n = 60; p = 1.4x10-53). This threshold
difference is present as early as age P14, the earliest age at which hearing can
be tested in mice.
54
Figure 11. Quantitative results of HsGJB2-R75W in the transgenic mice. (a) Transgene
copy number was estimated using QPCR analysis of gDNA from transgenic
mice. The average number of copies of MmGjb2 per nanogram of DNA is
205.9 ± 70.9 and the average number of copies of HsGJB2-R75W transgene is
72.1 ± 26.7 copies/ng of gDNA. The ratio of endogenous Gjb2 to transgene is
2.9 ± 0.3 to 1. As a means of validating our qPCR assay for determining
relative copy number, we quantified copies of endogenous Gjb2 relative to the
murine androgen receptor gene (Ar). Ar is located on the X chromosome. The
actual qPCR assay showed a Gjb2:Ar ratio of 4.0 ± 0.5 : 1 for male mice
(n=27) and 2.1 ± 0.7 : 1 for female mice (n=26). These results are consistent
with the expected ratio. (b) mRNA expression levels of the transgene
compared with endogenous Gjb2 in different tissues as measured by QRTPCR. The ratio of transgene to endogenous Gjb2 expression in the cochlea is
1.6 to 1.
55
56
Figure 12. Phenotype of palmoplantar keratoderma with deafness. Sample audiogram
(left) showing down-sloping pattern of severe-to-profound hearing loss in a
patient with dominant CX26-related hearing loss. Palmoplantar keratoderma
(right) manifests as thickening of the skin on the hands and the feet with
sloughing and occasional constriction banding around the digits (c). Adapted
from Feldmann et al. 2005.
57
Figure 13. PPK-like skin disease in GJB2-R75W transgenic mice. Mice ranging in age
from P7 to 4 months of age with severe skin phenotypes are shown. All mice
carry the GJB2-R75W transgene. Skin on the backside and tail is scaly;
constriction bands on the tails lead to auto-amputation. Images (f) and (g) are
of the same mouse taken several days apart.
58
Figure 14. Cataracts in GJB2-R75W transgenic mice. Images of cataracts in mice
carrying the GJB2-R75W transgene (a, c, d) are shown in comparison to an
age-matched wild-type mouse (b). Images (a and b) are of non-dilated pupils;
image (c) is with a dilated pupil. Cataracts evident as white areas indicated
with arrows; arrowheads indicate artifact of photographic flash. Examination
of the mouse lens was performed with a slit lamp and photodocumented with
a digital camera (D100; Nikon, Tokyo, Japan). All images were taken using
identical camera settings and prepared by processing with identical image
software. Magnification is 20x. Images courtesy of Dr. Geoffrey Lively
(Department of Molecular Physiology and Biophysics, University of Iowa).
59
Figure 15. Lens histology in GJB2-R75W transgenic mice. Cross-sectional images of
lens tissue of mice carrying the GJB2-R75W transgene(b, d, f) and wild-type
mice (a, c, e) are shown stained with toluidine blue. The large nuclear
cataracts present in the transgenic mice stand in contract to the clear, wellorganized appearance of the wild-type lens. Images are of the whole lens (a,
b); lens periphery (c, d); center of lens (e, f). Images courtesy of Adam
Hedberg-Buenz (Department of Molecular Physiology and Biophysics,
University of Iowa).
60
Figure 16. Expression of GJB2 (CX26) in the cochlea. Connexin 26 is expressed in the
nonsensory epithelial cells (interdental cells of the spiral limbus, inner and
outer sulcus cells, sensory supporting cells, and cells within the root process
of the spiral ligament) shown in green, and the connective tissue cell system
(fibrocytes of the spiral ligament and spiral limbus, basal and intermediate
cells of the stria vascularis) depicted in brown. Each connexin 26 molecule is
known as a connexin (yellow). Six connexins oligomerize to form a
connexon, and each connexon joins with another to form a gap junction.
Adapted from Hildebrand et al. 2008.
61
CHAPTER 4: DESIGN AND VALIDATION OF RNA INTERFERENCE
Introduction
Having established the validity of the GJB2-R75W transgenic mouse as an animal
model of dominant deafness for our experiments, my next goal was to design an effective
therapeutic agent. Due to its silencing potency and sequence specificity, RNA
interference (RNAi) is considered the method of choice for post-transcriptional silencing
of the dominant-negative mutation. The selective suppression of defective alleles without
affecting wild-type expression offers the potential to ameliorate the effects of a number
of dominant genetic disorders. To date, RNAi-based therapy has shown promising results
in experimental models for neurological diseases,[92-94] cancer[95] and skin
disease.[96]
RNAi is a sequence-specific, post-transcriptional gene-silencing mechanism that
is induced by double-stranded RNA (dsRNA). RNAi was first described in 1998 by Fire
and Mello[97] who observed that dsRNAs can trigger silencing of complementary
mRNA sequences in the nematode Caenorhabditis elegans. The RNAi mechanism was
soon observed to apply to higher organisms, including mammals. dsRNAs from viral
genome, microRNA (miRNA) precursors, or artificial dsRNA are digested into 21- to 23nucleotide fragments of siRNA by a member of the RNase III family of ATP-dependent,
dsRNA-specific ribonucleases called Dicer. These siRNA duplexes bind to a nuclease
complex to form the RNA-induced silencing complex (RISC), with the anti-sense strand
(guide RNA) serving as a template for specific transcript recognition. When the guide
RNA is extensively complementary to the target mRNA, RISC triggers rapid cleavage
and degradation of the mRNA [55]. Argonaute 2 (Ago2) is the slicer enzyme in RISC
that cleaves target mRNA. As few as one or two base-pair mismatches can significantly
abolish the silencing function of siRNAs, especially when these mismatches are at the
middle or 3’ end of the targeted sequence (Figure 17).[98] With the recognition that
62
synthetic small interfering RNAs (siRNA)—21–23 nucleotide RNA duplexes with twonucleotide 3’overhangs at each end—can be directly administered to the cytoplasm for
incorporation into RISC and gene silencing, siRNA has become one of the most
intensively studied tools for possible gene therapy.[99, 100] By targeting a sequence from
hepatitis C virus with siRNA, McCaffrey and colleagues were the first to show that RNAi
could be used in vivo in mice.[101] RNAi-mediated gene silencing has been achieved in
subsequent murine models by using naked synthetic siRNA, liposomes, plasmid and viral
vectors.[99]
Previous work performed in our lab by Dr. Yukihide Maeda showed siRNAmediated in vitro suppression of the R75W allele variant of GJB2 in cultured mammalian
cells, without significantly affecting levels of endogenous murine Gjb2.[102] When this
siRNA was exogenously administered to mouse cochleae along with a plasmid
expressing the GJB2-R75W transgene, the siRNA prevented hearing loss that would
otherwise be caused by the transgene.[102] This study provided proof-of-principle that
RNAi could effectively prevent hearing loss caused GJB2-R75W in vivo. My goal was to
build on this work by designing and validating potent and specific siRNAs that can be
used to silence the endogenous transgene of GJB2-R75W transgenic mice.
siRNA Design
Since the discovery of RNAi, efforts have been made to design potent and
specific siRNAs. The most successful siRNAs are able to achieve >90% gene silencing at
concentrations between 1 and 20 nM.[103] Highly functional siRNAs are those that
achieve 75-80% knockdown or greater.[104] Designing potent and specific siRNAs,
however, is a complex process. Since the majority (between 50-80%) of randomLy
designed siRNAs are poorly functional, effective siRNA design requires understanding
the properties of functional siRNAs compared with less effective siRNAs. In the past
decade researchers have sought to identify rules for siRNA design by looking for
63
consistent features shared by effective siRNAs.[105] The earliest design methodologies
relied on empirical rules related to length and GC content of the oligonucleotides.[106]
Silencing requires a minimum of 19 bases of target binding.[107] The most commonly
recommended length for siRNAs is 21-23 bases, allowing for a 19- to 21-base duplex
region designed with two-base overhangs at each 3’ end.[106, 108] However, Dicer
substrate siRNAs (D-siRNA), which are 27 bases in length, have been shown, in some
cases, to achieve superior knockdown when compared to the corresponding 21mers.[105, 109] Longer RNA duplexes have been shown to trigger an interferon (IFN)
response through activation of Toll-like receptor 3 by double-stranded RNA.[110] The
length of RNA that will trigger this IFN response appears to vary depending on cell type,
but the 27-mer D-siRNAs have not been reported to trigger this response.[105] It is
generally accepted that GC content should be between 30-64%,[104, 105] although some
authors recommend more stringent guidelines of 30-52% or 40-55%.[111, 112] High GC
content inhibits dissociation of the duplex and prevents effective loading into the RISC.
Very low GC content is associated with decreased functionality, likely due to lower
affinity for the target.[104]
As the molecular mechanisms of RNAi became better understood and more
studies that systematically tested multiple siRNAs across a gene were published, socalled Rational Design algorithms were generated. These algorithms lead to an increased
likelihood of designing better functioning siRNAs. When referring to specific base
positions and ends of siRNA as the Rational Design algorithms do, it is important to be
consistent with nomenclature. The anti-sense (AS) strand of siRNA is also referred to as
the guide strand. This is the active strand that is loaded into the RISC and pairs with the
target mRNA, marking it for cleavage. The sense (S) strand of siRNA is the strand that is
not loaded into RISC and it shares an identical sequence with the target mRNA. The S
strand is also referred to as the passenger strand, which is destroyed upon incorporation
of the guide strand into RISC.[108] When base preferences at specific positions are
64
referred to in siRNA design, it is most common to refer to the S strand with the 5’-most
base designated as position 1.
One of the most important features identified by Rational Design studies is
differential end stability. It was found that the stability of the duplex determines which of
the two strands is loaded into the RISC and, therefore, acts as the guide strand for
targeting mRNA knockdown.[113, 114] The strand whose 5’ terminus is on the less
stable end of the duplex is preferentially loaded into the RISC, becoming the guide
strand. Reynolds and colleagues used the number of A/U bases in positions 15-19 of the
sense strand as a approximation of relative 5’ anti-sense instability. They found that
siRNAs with more A/U bases in positions 15-19 were more likely to achieve high levels
of silencing, while siRNAs that completely lacked A/U bases in positions 15-19 were all
non-functional.[112] Other features included in Rational Design algorithms for functional
siRNA prediction are: (i) GC content; (ii) specific base preferences (G/C at position 1, A
at positions 3, 6 and 19, U at position 10, A/U at position 13, no U at position 1, and no
G/C at position 19 of the sense strand); (iii) lack of certain motifs known to trigger an
interferon (IFN) response (e.g. GUCCUUCAA); (iv) absence of more than four of any
single base in a row; (v) absence of stretches of internal complementarity that could lead
to hairpin formation.[104, 106, 112, 115]
Second generation siRNA prediction algorithms have been generated by
combining large-scale experimental results with computer-learning. Many of these
algorithms use artificial neural networks (ANNs) to select the most efficient siRNAs.
ANNs, often referred to as “black box” algorithms, are adaptive networks used to find
patterns in large sets of data and model complex relationships.[106] Beginning in 2005
with the BIOPREDsi algorithm which used 2431 randomLy selected siRNAs targeting 34
genes to train the ANN, a number of publically available second-generations algorithms
have been published (17129386; and also siRNA Scales and i-Score).[116-121] siRNA
design tools can also be accessed through the websites of many companies that
65
manufacture siRNAs. For example, the online IDT SciTools RNAi Design website
(idtdna.com/Scitools/Applications/RNAi/RNAi.aspx, Integrated DNA Technologies,
Coralville, IA) allows users to input target gene sequence and then the program generates
a list of Dicer substrate siRNAs (D-siRNA) to test.[105, 106] The IDT algorithm uses
another machine-learning methodology, support vector machine (SVM), to build its
predictive models of RNAi activity.[122]
While prediction algorithms and calculators can generate lists of strong candidate
siRNA, the functionality cannot be perfectly predicted until tested. Some siRNAs with
very high scores do not perform well while some siRNAs with low scores perform very
well.[106] The specific silencing capacity of an siRNA is the ultimate determinant of its
functionality and so siRNAs must be tested experimentally to determine actual
effectiveness. Published protocols recommend using online siRNA selection tools and
other published selection algorithms to identify at least 4-6 good candidate siRNAs for in
vitro testing.[104, 105, 111]
After potent siRNAs (>75-80% in vitro silencing) are identified and validated,
consideration must be given to the best method for in vivo application. In the previous
study, Dr. Maeda used lipid-complexed siRNA to achieve transient knockdown of an
exogenously delivered transgene. Long-term in vivo silencing of an endogenous gene by
RNAi requires a different strategy. Rather than using lipid-complexed siRNAs, long-term
in vivo expression requires delivery by a viral vector leading to transcription of shRNAs
or artificial miRNAs. In this chapter, I describe how we used Rational Design and a
computer-generated algorithm to design and validate siRNAs for effective silencing of
the GJB2-R75W transgene.[105, 112] I also describe how we re-designed them as
artificial miRNAs to be packaged in a viral vector for in vivo application.
66
Materials and Methods
siRNA Design
siRNA2 was previously designed using Rational Design rules.[102] The other ten
D-siRNAs were new designs. GJB2-A, GJB2-B, GJB2-C, GJB2-D and GJB2-E were
hand-designed using Rational Design guidelines with preference given to regions with
mismatches between the transgene and endogenous MmGjb2 transcript.[112] GJB2-38,
GJB2-289, GJB2-514, GJB2-550, GJB2-596 were designed in silico using a new
proprietary computer algorithm (Integrated DNA Technologies, Coralville, IA). This
algorithm adapts the online IDT SciTools RNAi Design engine with a mismatch site
finder to identify top candidates for potent and specific knockdown of the GJB2-R75W
transgene.[122]
All chemical D-siRNAs described in this study were synthesized and high
performance liquid chromatography-purified by Integrated DNA Technologies. The
identity of each compound was verified using electrospray ionization mass spectrometry
and the purity of the compounds was confirmed, using analytical high performance liquid
chromatography, as being >90%. Duplexes were also QC tested by analytical HPLC and
were >90% pure. Final duplexes were prepared as sodium salts. The sequences are
provided in Table 3.
Plasmid Constructs
Plasmids used for transfection and for QPCR serial dilution standards included
pHsGJB2-R75W-myc-FLAG, pMmGjb2-WT-myc-FLAG, and pMmGjb6-WT-mycFLAG. All three were cloned similarly into the 3pTmod2 vector (Charles Searby,
University of Iowa). The 3pTmod2 vector was created from the pcDNA3.1(-)/myc-His
plasmid (Invitrogen, Carlsbad, CA), with the His tag replaced by a FLAG tag. Sequences
can be readily cloned into this plasmid, reading frame B, using XhoI and BamHI
restriction sites. Forward primers with XhoI sites included and reverse primers with
67
BamHI sites included were designed for each of the three sequences. The primers for
HsGJB2-R75W (restriction sites underlined) are: 5’—
TTCTCGAGATGGATTGGGGCACG—3’ and 5’—
TGGGAAGTCAAAAAAGCCAGTGGATCCTT—3’. The primers for MmGjb2-WT
are: 5’—TTCTCGAGATGGATTGGGGCACA—3’ and 5’—
AGGAAAGTCCAAAAGACCAGTGGATCCTT—3’. The primers for MmGjb6-WT
are: 5’—TTCTCGAGATGGACTGGGGGACC—3’ and 5’—
TGCAATCACAAGTTTCCCAAGTTCGGATCCTT—3’. Primers were used for PCR
amplification of the insert, followed by double digestion (XhoI and BamHI) of the PCR
products and 3pTmod2 plasmid. Linearized plasmid was CIP-treated and ligation
reaction performed overnight. Ligation products were transformed into XL-1 blue
supercompetent cells, grown on ampicillin plates and four colonies selected for plasmid
extraction and purification with DNA mini-prep (QIAGEN). All clones were confirmed
by sequencing.
Cell Culture and Transfection
Cell culture media and supplements were purchased from Gibco (Invitrogen)
unless otherwise specified. CHO cells were cultured in Dulbecco’s Modified Eagle
Medium (DMEM) that was supplemented with 10% FBS (Thermo Fisher Scientific,
Waltham, MA), pen-strep and L-glutamine, 100U/mL penicillin, and 100 µg/mL
streptomycin. Cell cultures were passaged every two or three days and maintained
between 50 and 90% confluency.
For knockdown assays, CHO cells were seeded at a concentration of 1.0 × 105
cells/well in 24-well plate in 1mL of media per well. Plasmid transfections were
performed using FuGene6 (Roche) and siRNA transfections performed using RNAiMax
reagent (Invitrogen), each according to manufacturer’s protocol. Quantities used in
typical transfection reactions were: 500ng of target plasmid DNA per well; 1500ng of
68
artificial miRNA plasmid DNA per well; and final concentration of 10nM siRNA per
well. Transfections were performed in serum-free media and allowed to incubate for at
least 6 hours prior to changing media. Typically, cells were harvested after 24 hours, but
in some cases left for up to 96 hours. For longer experiments, media was changed daily.
Cells are lysed and RNA extracted using RNeasy Tissue Mini Kit (QIAGEN Inc.,
Valencia, CA) according to manufacturer instructions and treated with on-column DNase
I digestion. The resulting RNA samples were subjected to QRT-PCR as described below.
Reverse Transcription
Following RNA extraction, concentration of purified RNA was measured by
NanoDrop 2000 Spectrophotometer (Thermo Scientific NanoDrop Products, Wilmington,
DE), and diluted in RNase-free water to a final concentration of [50 ng/µL] for reverse
transcription. Reverse transcription was performed using 400 ng of total RNA per
sample. Each sample was used for three reverse transcription reactions. Reverse
transcription was performed using Superscript III (Invitrogen) according to manufacturer
protocol. Each reaction mixture consisted of: 8 µL (400ng) RNA, 1 µL Oligo(dT)20
primer, 1 µL Random Hexamer primer, 1 µL 10mM dNTP, 2 µL 5x RT buffer, 2 µL
50mM MgCl2, 2 µL water, 2 µL 0.1M DTT, 1 µL RNaseOUT, and 1 µL SSIII reverse
transcriptase. Following RT reaction (50°C for 50 minutes, followed by termination at
85°C for 5 minutes), the cDNA samples are treated with 0.5 µL 5U/µL RNaseH (New
England Biolabs) at 37°C for 20 minutes, spun down, and diluted in 18.5 µL water for a
total volume of 40 µL cDNA at a final concentration of 10ng/µL.
Quantitative Real-Rime PCR (QPCR)
Quantitative real-time PCR assays (QPCR) were done using 20 ng cDNA per 10
μL reaction. Immolase DNA polymerase (Bioline, Randolph, MA), 200 nM primers, and
200 nM probe. Cycling conditions employed were 95°C for 10 minutes followed by 40
cycles of 2-step PCR with 95°C for 15 seconds and 60°C for 1 minute. Reaction mix and
69
cycling conditions are summarized in Table 4. Primers and TaqMan probes (Applied
Biosystems Inc., Foster City, CA) used in QPCR experiments are listed in Table 3. The
PCR and fluorescence measurements were done using an AB 7900HT Fast Real-Time
PCR System (Applied Biosystems Inc.) in 384-well format. Expression data were
normalized to levels of an internal control gene, either MmHprt, CHO-Hprt, or
MmRpl23. Copy number standards (102 to 107 molecules/µL) were run in parallel using
linearized cloned amplicons corresponding to each of the assays (Figure 18). Unknowns
were extrapolated against standards to establish absolute quantitative measurements using
the Applied Biosystems Sequence Detection software and data were further analyzed by
Excel spreadsheets (Microsoft, Redmond, WA). All data points reported are the mean of
three replicate assays and error is reported as the standard error.
Artificial miRNA design and construction
Artificial miRNAs (miRNA2, miGJB2-C, miGJB2-D and miNC1) were
constructed based on the sequences of siRNA2, GJB2-C, GJB2-D and NC1. The
artificial miRNA design was based upon the backbone of miR-30 as previously
described.[123] The hairpin duplexes for each artificial miRNA were ordered from
Integrated DNA Technologies as 5’-phosphorylated 4 nmole ultramers. They were
hydrated in 40 µL IDTE pH 7.5 to a concentration of 100 µM, combined 100 pmoles of
top strand and 100 pmoles of bottom strand in STE (10 mM Tris pH 7.5, 0.1 mM EDTA,
100 mM NaCl) to final 50ul volume (2 µM each strand). These were heated to 95C for 5
min and cooled slowly. After annealing, I added 50 µL of TE was added to dilute each
duplex to 1 µM.
To create the pAAV-tVal-CMV-dsRED2 and pAAV-H1-CMV-dsRED2 plasmids
for insertion of the artificial miRNA sequence, we digested pUC19+CMV-dsRED2
(Invitrogen), pAAV-tRNA, and pAAV-H1 with KpnI and BamHI. The CMV-dsRED2
fragment from pUC19+CMV-dsRED2 was gel purified, as were the post-digestion
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linearized pAAV-tRNA and pAAV-H1 plasmids. The CMV-dsRED2 fragment was then
ligated into each of the two vectors using T4 DNA ligase to form pAAV-H1-CMVdsRED2 and pAAV-tRNA-CMV-dsRED2. These constructs were then digested with
NheI and BamHI and gel purified. Finally, the artificial miRNA duplexes were ligated
into the linear plasmids with T4 DNA ligase, and transformed into TOP10
cells. Colonies were grown up in a 96-well plate and the plasmid was isolated using
broth purification followed by sequence confirmation.
Results
Rationally and Computationally Designed siRNA
Candidates
siRNA2 had previously been designed and proven effective in vitro and in vivo
by our lab.[102] siRNA2 is a classic 21-mer with two-nucleotide 3’ overhangs. siRNA2
was designed using early published guidelines. It potently knocked down expression of
GJB2-R75W in vitro (~85% knockdown) and only mildly affected the expression of
endogenous Gjb2 (~40% knockdown).[102] In order to have more available options for
future therapeutic use, I designed ten additional Dicer-substrate siRNAs (D-siRNA) for
experimental validation. Five of these were hand-designed (“eyeball selection”) with the
help of Drs. Mark Behlke and Scott Rose (Integrated DNA Technologies, Coralville, IA).
For these five (GJB2-A, GJB2-B, GJB2-C, GJB2-D, GJB2-E), general rational design
guidelines were considered as well as mismatches with the endogenous Gjb2
transcript.[112] The other five D-siRNAs (GJB2-38, GJB2-289, GJB2-514, GJB2-550,
GJB2-596) were designed by a new proprietary computer algorithm created by Kyle
McQuisten (Integrated DNA Technologies, Coralville, IA). This algorithm adapted the
online IDT SciTools RNAi Design engine with a mismatch site finder to identify top
candidates for potent and specific knockdown of the GJB2-R75W transgene.[122] The
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sequences for siRNA2 along with the ten newly-designed candidate D-siRNAs are shown
in Table 5.
Potent and Specific siRNA-mediated Knockdown
We used a Chinese hamster ovary (CHO) cell culture co-transfection strategy to
experimentally determine knockdown efficiency of our rationally designed siRNAs.
Equal numbers of CHO cells (1x105 cells/well) were plated in 24-well plates and
incubated under standard conditions overnight. We then performed a co-transfection with
a plasmid expressing the GJB2-R75W transgene and siRNA targeting the transgene. NosiRNA and scramble sequence siRNA (NC1) negative controls and a validated
endogenous Hprt-targeting siRNA positive control were used. Cell co-transfections were
performed in triplicate wells and then the triplicates combined at time of cell harvesting
24 hours post-transfection. RNA was extracted from the cells and each RNA sample used
for three reverse transcription (RT) reactions. Triplicate RTqPCR reactions were then
performed to quantify target mRNA knockdown by each of the siRNAs. GJB2-R75W (or
Hprt for positive control) expression results were normalized to levels of Rpl23 internal
control and knockdown levels of GJB2-R75W (or Hprt) were normalized to negative
control siRNA (NC1) treated cells = 100% expression (0% knockdown). The four most
effective siRNAs were siRNA2 (89 % ± 0% knockdown), GJB2-C (84% ± 6%), GJB2-D
(85% ± 4%), and GJB2-596 (79% ± 3%). The Hprt knockdown with Hprt-targeting
siRNA was (79% ± 3%), verifying the transfection efficiency and overall integrity of the
assay (Figure 19). siRNA2, GJB2-C, and GJB2-D were the three siRNAs that achieved
our in vitro goal of ~80% knockdown of the GJB2-R75W transgene. In multiple trials,
these three siRNAs consistently achieved the greatest degree of knockdown compared to
the other siRNAs. A similar assay was performed using MmGjb2 as the target instead of
GJB2-R75W. This experiment verified that none of the three highly effective siRNAs
had a silencing effect on endogenous Gjb2 expression (data not shown). Thus, we have
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confirmed siRNA2 and identified GJB2-C and GJB2-D as siRNAs that potently will
silence the GJB2-R75W transgene in vivo.
Artificial miRNA Design and Validation
siRNA2, GJB2-C and GJB2-D were next prepared for in vivo application by
redesigning them as artificial miRNAs. Artificial miRNAs have been shown to be
preferable to shRNA constructs in minimizing in vivo toxicity in the mouse brain.[124]
We incorporated the siRNA sequences into an artificial miRNA scaffold based on human
miR-30, creating miRNA2, miGJB2-C, and miGJB2-D.[124, 125] The same was done
using the scrambled siRNA control (NC1), creating miNC1 (Figure 20).
The next consideration in preparation for in vivo application was choice of
promoter to drive the artificial miRNA expression. The goal is to achieve sufficient levels
of transgene silencing while avoiding the in vivo toxicity that can be associated with in
high levels of exogenous shRNA or artificial miRNA. The U6 small nuclear RNA
(snRNA) promoter (U6) and the H1 RNase P RNA promoter (H1) are the two most
commonly used RNA polymerase III (pol III) promoters for driving expression of
shRNAs.[111, 126] H1 is a weaker promoter than U6,[127] but both promoters have been
associated with toxicity due to their high transcriptional activity.[128, 129] A third
promoter, the transfer RNA (tRNA)-valine promoter (tVal), is weaker than the other two
and has been shown to effectively express shRNAs in vivo without reported
toxicity.[129] We selected H1 and tVal, the two weaker pol III promoters, to drive our
artificial miRNA expression and made constructs for each artificial miRNA to be driven
by each promoter (H1-miRNA2; H1-miGJB2-C; H1-miGJB2-D; H1-miNC1; tValmiRNA2; tVal-miGJB2-C; tVal-miGJB2-D; tval-miNC1).
The artificial miRNAs were next cloned into expression plasmids. We replaced
the more commonly used GFP with DsRed because the GJB2-R75W mice already
express GFP with the transgene. The use of DsRed as a fluorescent label for artificial
73
miRNA expression enables us to detect and compare both transgene expression and
artificial miRNA expression when applied to the transgenic mouse. The DsRed protein
expression is independently driven by a CMV promoter (Figure 21). In total, 8 artificial
miRNA expression plasmids were made: pAAV_H1-miRNA2; pAAV_H1-miGJB2-C;
pAAV_H1-miGJB2-D; pAAV_H1-miNC1; pAAV_tVal-miRNA2; pAAV_tValmiGJB2-C; pAAV_tVal-miGJB2-D; pAAV_tval-miNC1.
Artificial miRNA expressed from a plasmid is markedly different than simple
transfection of naked siRNA and so it was important to assess the effectiveness of the
new constructs before packaging in viral vectors for in vivo use. We used an in vitro cell
culture co-transfection assay similar to the assay used to verify siRNA effectiveness.
CHO cells were again plated in a 24-well plate and co-transfected in triplicate wells with
the target plasmid (pGJB2-R75W_myc-FLAG) and one of the artificial miRNA
plasmids. After 24 hours the triplicate wells were combined, RNA extracted, reverse
transcribed in triplicate and mRNA levels quantified with qPCR in triplicate. GJB2R75W expression results were normalized to levels of CHO-Hprt internal control and
knockdown levels of GJB2-R75W were normalized to negative control artificial miRNA
(pAAV_H1-miNC1) treated cells = 100% expression (0% knockdown). Results for
knockdown of the GJB2-R75W transgene by the artificial miRNA constructs were:
pAAV_H1-miRNA2 (20% ± 5% knockdown), pAAV_H1-miGJB2-C (18% ± 15%),
pAAV_H1-miGJB2-D (20% ± 8%), pAAV_tVal-miRNA2 (1% ± 9%), pAAV_tValmiGJB2-C (33% ± 12%), and pAAV_tVal-miGJB2-D (42% ± 11%) (Figure 22).
The artificial miRNA constructs driven by tVal performed slightly better than
those driven by H1 and so we re-tested the tVal artificial miRNAs to verify knockdown.
The quantitative knockdown data were variable from experiment to experiment, but
pAAV_tVal-miGJB2-D consistently out-performed the other constructs (data not shown).
The greatest degree of knockdown achieved by pAAV_tVal-miGJB2-D in an in vitro
assay was 82% ± 13% (18% ± 13% transgene expression) (Figure 23). As the graphical
74
data make evident, however, the other artificial miRNA-treated samples in this assay
showed increased transgene expression compared with the negative control samples and
so these data are probably not as reliable as the previous assay showing a 42% decrease
in transgene expression for samples treated with pAAV_tVal-miGJB2-D. The variability
in the results will be revisited in the Discussion section. Despite the variability, however,
it was clear that pAAV_t-Val-miGJB2-D achieved the best transgene silencing in vitro
and is our construct of choice for viral vector packaging and in vivo application.
Discussion
This chapter describes our work in designing an RNAi-based therapeutic for in
vivo treatment of dominant-negative GJB2-R75W deafness. It builds on previous work
performed in our lab in which an siRNA (siRNA2) was shown to effectively silence
expression of the exogenously expressed GJB2-R75W transgene in vitro (>80%
knockdown) and in vivo (>70% knockdown).[102] Based on these strong previous results
it may seem redundant to re-test the efficacy of siRNA2 and to re-design and test new
siRNAs targeting GJB2-R75W. However, we felt it worthwhile to design and test new
siRNAs for the following reasons: (i) second-generation computer-based design
algorithms had been developed since we had designed siRNA2 and perhaps these
algorithms could identify even more potent siRNAs; (ii) Dicer substrate siRNAs (DsiRNA) sometimes perform more effectively than traditional siRNAs and therefore could
prove more potent than siRNA2;[130] (iii) different siRNAs will have different potentials
for toxicity through off-target effects and stimulating an immune response; (iv) pooling
multiple siRNAs against a single target has proven useful in some instances;[104, 106]
(v) the more siRNAs we test, the more therapeutic tools we will have at our disposal for
in vivo application and targeting different locations of the same transcript can serve as an
important control for verifying target-specific effects of RNAi-mediated
knockdown.[104, 131]
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siRNA Design
Designing effective siRNAs can be a complex process, with hundreds of
candidate siRNAs per gene and dozens of available algorithms to use in prioritizing the
candidates. Additionally, there are several variations on the standard siRNA structure.
After consultation with collaborators Dr. Mark Behlke and Dr. Scott Rose (Integrated
DNA Technologies, Coralville, IA) we decided to design D-siRNAs for initial silencing
of the GJB2-R75W transgene. D-siRNAs are 25/27-nucleotide duplexes with a two-base
3’ overhang on the anti-sense strand.[105] D-siRNAs are designed to promote predictable
cleavage by Dicer through imitating structural features of pre-miRNAs. Therefore, DsiRNAs are synthesized with a two-base 3’ overhang on the AS strand with the opposite
end of the duplex being blunted to mimic the closed loop of pre-miRNAs. This structure
discourages processing from the blunted end and increases the likelihood that Dicer will
cleave 21-23 nucleotides upstream of the two-base 3’ overhang, releasing the correctly
processed siRNA for incorporation into RISC.[105, 130] Chemically synthesized DsiRNA duplexes have been reported to have up to a 100-fold increase in potency
compared with corresponding 21-mers directed at the same site.[130] This increased
potency is thought to be due to Dicer processing of the D-siRNAs, which leads to more
efficient loading into the RISC.[105]
We used two methods for designing the D-siRNAs: (i) “Eyeball selection” in
which we hand-picked D-siRNAs based on applying Rational Design rules [105, 112] to
regions in the target mRNA where the GJB2-R75W transgene sequence had differed
from the endogenous mouse Gjb2 sequence; (ii) in silico selection based on an computergenerated algorithm developed at Integrated DNA Technologies (IDT). Although five DsiRNAs were selected by each method (see Table 5), none of the candidate D-siRNAs
targeted the actual transition mutation (c.223C>T) underscoring our observation that
c.223C>T is in a region which is less than ideal for siRNA targeting.[102] This dilemma
highlights a potential problem for RNAi therapeutics which require allele-specific gene
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silencing: the choice of siRNA is restricted by the location of the mutation and siRNAs
targeting that location may not be effective.[108] One strategy to circumvent this obstacle
is to introduce an additional permissive mismatch in the siRNA. Thus, the siRNA would
have two mismatches with the non-target allele making it less likely to cause silencing,
but it would only have a single mismatch with the target allele which, if in a permissive
position, should not significantly inhibit silencing.[132] Our lab has successfully used
this strategy to design allele-specific siRNA for the p.G286S (c.856G>A) hearing loss
mutation in KCNQ4. A second approach to overcoming allele-specific targeting problems
is to design siRNA that will non-discriminately silence both alleles, followed by
exogenous replacement of the wild-type allele or protein.[133] To this end, vectors have
been constructed that will simultaneously express both shRNA and a transgene.[134] Our
experimental goal, however, was to prevent translation of the endogenously expressed
human GJB2-R75W transgene and so we exploited species-specific sequence differences
to achieve this goal. This approach greatly increased the size of our target sequence and
improved the likelihood of identifying potent siRNAs.
Design Algorithms
In our experience second generation computer-based algorithms did not perform
better than more basic Rational Design-based selection. Each of our top three most potent
siRNAs (siRNA2, GJB2-C, GJB2-D) was hand-picked based on Rational Design rules
rather than by in silico selection. We caution that our experience with a single computer
algorithm used for targeting a single gene should not be generalized to minimize the
value or effectiveness of in silico tools for siRNA selection. The lesson that we can
emphasize, however, is that computer-based siRNA selection programs are not perfect
predictors of siRNA effectiveness. Actual in vitro verification is absolutely essential
whether hand-picking siRNAs based on Rational Design rules or performing in silico
siRNA selection. Our experience also suggests that the Rational Design rules described
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between 5 and 10 years ago identified many of the most important rules of siRNA design
and remain a viable alternative to in silico design methods. For future siRNA design, we
recommend strategies that incorporate Rational Design methods with in silico scoring
algorithms.[104, 117] Our experience shows that experimentally validating more than
just a few candidate siRNAs may be more important than the selection method used. In
fact, we recommend testing the top 3-5 candidate siRNAs generated from at least two
different selection methods as we did in this study.
Artificial miRNA vs. shRNA
After identifying three potent siRNAs, we redesigned for long-term in vivo
expression. shRNAs and artificial miRNAs, delivered with a viral vector, have both been
used for in vivo RNAi therapy. Data from several studies suggest that artificial miRNA
constructs are advantageous to shRNA constructs. McBride and colleagues observed that
some shRNAs, both gene-targeting and control, caused toxicity when delivered to the
mouse striatum. Delivery of artificial miRNAs, however, was not associated with
neurotoxicity.[123, 124] Additionally, Silva and colleagues found that artificial miRNAs
were more effective that equivalent shRNAs at gene silencing.[135] They observed that
artificial miRNAs were up to 12 times more efficient than shRNAs at producing the small
RNA for loading into RISC.[136] With the desire for increased potency and decreased
risk of toxicity, we selected artificial miRNAs as the design of choice for in vivo use. Our
highly effective siRNAs were re-designed as artificial miRNAs modeled after the
backbone of the primary miR-30 miRNA (Figure 20).[123-125, 135, 136]
Promoter Selection
The promoter that drives expression of shRNA or artificial miRNA is a major
determinant of the intensity and duration of gene silencing.[127] However, it is important
to find the balance between effective silencing and toxicity. High levels of duplex RNA,
even when expressed from a transduced or transfected vector, can activate an innate
78
immune response.[131] Toxicity can also result from high levels of miRNA or shRNA
saturating the endogenous RNAi machinery.[128] This toxicity, assocatied with downregulation of endogenous miRNAs, is due to saturation of Exportin-5, which functions in
nuclear export and stabilization of shRNAs/miRNAs and is readily oversaturated.[128]
The goal for effective in vivo RNAi therapy is to express sufficient siRNA to achieve
silencing without expressing so much that it becomes toxic. Ideally, we would deliver the
lowest levels of siRNA that would effectively silence our target. This therapeutic window
may be relatively small, and tightly controlling levels of siRNA presents a particular
challenge to applications that use shRNA or artificial miRNA expressed from plasmids or
viruses. Promoter choice provides an opportunity to regulate levels of expression.
In a study comparing the effectiveness of different promoters (U6, H1, tVal, and
pol II CMV) in driving shRNA expression and gene silencing, Silva and colleagues found
that the promoter made no significant difference when highly effective shRNAs were
used. However, in cases where less effective shRNAs were used, the pol III U6 promoter
and the pol II CMV promoter provided the best and most consistent silencing.[135]
While both U6 and H1 have been implicated in toxicity from high levels of shRNA
expression in vivo,[128] the tVal promoter has been used drive shRNA expression in the
mouse brain without reported toxicity.[129] tVal appears to be weak enough to prevent
toxic accumulation of exogenous shRNA or artificial miRNA transcripts while still
providing enough expression for gene silencing with effective shRNAs and artificial
miRNAs. We selected the tVal promoter as the safest option for in vivo expression of our
artificial miRNAs. We also made artificial miRNA constructs driven by the H1 promoter
in case stronger expression was required.
In Vitro Testing of Artificial miRNA Constructs
There are a number of differences between the siRNA originally validated in vitro
and the artificial miRNA plasmids designed to be expressed in vivo: (i) artificial miRNA
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hairpin structure rather than siRNA or D-siRNA; (ii) enters the miRNA pathway
upstream of siRNA; (iii) requires transcription in the nucleus; (iv) expressed from a
plasmid using either the H1 or tVal promoter; (v) co-expressed with DsRed fluorescent
marker protein. Any of these factors could impact the overall effectiveness of transgene
silencing. We felt it necessary, therefore, to assess in vitro knockdown potency for each
of these new constructs. Another practical purpose for performing this in vitro
knockdown verification was to avoid the high cost associated with packaging each of
these constructs into a viral vector and testing them in vivo. The most effective artificial
miRNA construct would be selected for packaging into a viral vector and subsequent in
vivo testing.
Assessing gene silencing with the artificial miRNA plasmids was performed
similarly to our initial siRNA transfection assays. We used CHO cells and co-transfected
both the target plasmid and the artificial miRNA plasmid. However, our qPCR results
were not as consistent and gene silencing not as robust as in the siRNA assays. One assay
showed up to 82% transgene knockdown with pAAV_tVal-miGJB2-D (Figure 23), but
the assay with the most reliable results showed a 42% transgene knockdown with the
same construct (Figure 22). It was evident that pAAV_tVal-miGJB2-D achieved
knockdown and that it was our most effective construct, but the actual degree of in vitro
knockdown was difficult to determine. It was not surprising that some of the constructs
would perform better than others and achieve different degrees of knockdown compared
to their siRNA counterparts. We were surprised, however, by the degree of variability
that we observed.
Discussion of Methods and Alternate Strategies
Variability is inherent in all experimental procedures, and a multi-process
experiment such as this has many sources of variability. The major stages in the
experiment are: (i) cell culture, transfection, and RNA extraction; (ii) reverse
80
transcription; (iii) QPCR. In each of the major stages replicates are used to help control
for inevitable variation. Cell culture and transfection is performed in triplicate, as are the
RT reactions and the QPCR reactions. If true triplicate reactions were performed at each
of the three stages, however, the number of data points resulting from a single siRNA or
artificial miRNA test is 27 (3 at transfection, 9 at RT, 27 at QPCR). In order to keep the
number of reactions to a more manageable level, we decided to pool the cells from
triplicate wells at the time of harvesting them and prior to RNA extraction (Figure 24).
This took the number of total data points from a single siRNA or artificial miRNA test to
9 (3 at RT, 9 at QPCR). We felt that pooling samples at this step would not alter the
experimental outcome since the RNA from each of the triplicate wells would be
represented in the downstream reactions. However, we failed to recognize that pooling
the cells at this stage diminishes, and possibly nullifies, any benefits gained from the
initial triplicate transfections. One of the benefits of triplicate reactions is that they allow
for the identification and rejection of outliers. The cell culture and transfection stage of
the experiment is the only stage that deals directly with living cells. This biologic stage of
the experiment is the stage in which there is the most potential for unexplained variability
and, therefore, it is the experimental stage in which replicates would be most beneficial.
The later RT and qPCR reactions are expected to have less intrinsic variability and be
more reproducible than the stage involving living cells. In retrospect, therefore, we may
have achieved more consistent results had we performed true biologic triplicate reactions
by not pooling the cells prior to RNA extraction. For future experiments, in order to keep
the reactions to a manageable level while controlling for variability, we suggest
performing the triplicate transfections but only doing a single RT reaction per RNA
sample and then doing the qPCR in triplicate reactions. This modified experimental plan
would also result in 9 data points (3 at transfection, 3 at RT, 9 at QPCR), but would
maintain the biologic triplicates, perhaps resulting in greater consistency between assays.
81
Although RNAi silences genes through targeted mRNA degradation, proteins are
the functional end product of most genes. Assaying protein levels, therefore, can be
important for determining RNAi potency. For our preliminary siRNA screening assays
we chose to measure expression and knockdown at the mRNA level. CX26 has a
relatively short half-life (1.3-5.0 hours, depending on cell type), which makes it an ideal
target for RNAi-mediated silencing.[43, 137] Because of the short half-life, mRNA
present in the cells after RNAi-treatment should approximate protein levels, an
assumption which would not necessarily hold true for proteins with slow turnover. We
also decided to focus on mRNA levels because of the high quantitative power of qPCR
compared with Western blot.[75] There is another protein-based assay that is now
routinely used for assaying siRNAs: the dual-luciferase assay. This assay (DualLuciferase™ Reporter kit, Promega) uses a target gene sequence cloned downstream of
the Renilla luciferase gene as a reporter system. The same plasmid (psiCHECK™-2,
Promega) also expresses firefly luciferase, which serves as an internal normalizer. As
siRNA silences the Renilla luciferase gene-target gene mRNA, the ratio of Renilla to
firefly luciferase decreases, correlating with degree of silencing. This assay provides a
convenient way to measure RNAi effect without the additional steps of RNA extraction,
DNA removal, and reverse transcription. Although we are confident that qPCR is an
excellent quantitative assay, in the future we recommend verifying qPCR data with the
dual-luciferase assay.
In summary, we used a computer-generated algorithm and Rational Design rules
to select siRNAs for knockdown of the GJB2-R75W transgene. Out of the 11 siRNAs
tested, three achieved ≥80% silencing of the target in vitro (siRNA2, GJB2-C, GJB2-D)
(see Figure 25). We have discussed lessons learned and made recommendations
regarding the process of designing and validating siRNAs. With the goal of minimizing
risk for in vivo toxicity, we re-designed these siRNAs as artificial miRNAs to be
expressed in a plasmid driven by either the H1 or the tVal promoter. Based on in vitro
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knockdown efficiency, we selected pAAV_tVal-miGJB2-D as the most effective
construct for in vivo targeting of GJB2-R75W. This construct will be packaged into a
viral vector for therapeutic delivery to the developing mouse cochlea.
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Figure 17. RNA interference (RNAi) pathways. Endogenous micro RNA (miRNA) and
exogenous small interfering RNA (siRNA) pathways for gene silencing. The
siRNA pathways utilize the cellular machinery of endogenous miRNA
biogenesis and induce degradation of target messenger RNAs (mRNAs).
miRNA biogenesis results predominantly in translational repression of target
genes and in some cases degradation of target mRNAs. Adapted from Renne
2007.
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Table 3. QPCR primer and probe sequences.
85
Table 4. QPCR reaction conditions.
86
Figure 18. Sample QPCR output. Outputs from QRT-PCR amplification of HsGJB2R75W with Applied Biosystems 7900HT Fast Real-Time PCR System and
Sequence Detection Software. Amplification curves of triplicate known
standards (quantity is indicated) are shown on the left. Unknowns were
extrapolated against standards to establish absolute quantitative measurements
using the Applied Biosystems Sequence Detection software and data were
further analyzed. Sequence Detection Software-generated standard curve plot
is shown on the right with unknown samples indicated in red and standards
indicated in blue. Reactions were run at Integrated DNA Technologies
(Coralville, IA).
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Table 5. Sequences of siRNA and D-siRNAs.
Note: siRNA and D-siRNA designs used in knockdown assays targeting HsGJB2-R75W.
siRNA2 is a previously validated 21mer siRNA [102]. The others are newly-designed 27mer DsiRNAs. GJB2-A, B, C, D, E were “hand-picked” using rational RNAi design principles. GJB238, 289, 514, 550, 596 were generated by a computer algorithm at IDT (Coralville, IA, USA).
NC1 is a scrambled-sequence negative control D-siRNA provided by IDT. The sequence of each
duplex is shown with upper case letters representing RNA bases and lower case letters
representing DNA bases. Mismatches between HsGJB2-R75W and MmGjb2 are indicated in
blue. The 19-nucleotide sense sequence of each duplex is underlined. Expected Dicer cleavage
sites are also shown (‘). Bases in red are complementary to the 21-base target sequence.
88
Figure 19. In vitro siRNA knockdown of GJB2-R75W transgene. CHO cells were cotransfected with GJB2-R75W transgene and naked siRNAs to screen for
knockdown. mRNA expression levels were measured using QRT-PCR and are
shown in the graph.GJB2-R75W (or Hprt for positive control) expression
results were normalized to levels of Rpl23 internal control and knockdown
levels of GJB2-R75W (or Hprt) were normalized to negative control siRNA
(NC1) treated cells = 100% expression (0% knockdown). The four most
effective siRNAs for silencing were: siRNA2 (89 % ± 0% knockdown),
GJB2-C (84% ± 6%), GJB2-D (85% ± 4%), and GJB2-596 (79% ± 3%). The
Hprt knockdown with Hprt-targeting siRNA was (79% ± 3%).
89
Figure 20. Predicted artificial miRNA structures. Artificial miRNAs designed
fromsiRNA2, GJB2-C, GJB2-D, and NC1 and modeled after the backbone of
miR-30 are shown in their predicted hairpin folding conformation. Designs
and images courtesy of Scott Rose at Integrated DNA Technologies.
90
Figure 21. Vector maps for pAAV-tVal-CMV-dsRED2 and pAAV-H1-CMV-dsRED2.
Cloning vectors used for artificial miRNAs. The artificial miRNA sequence is
inserted between the NheI and BamHI restriction sites. The weak Pol III
promoters, tVal (orange) or H1 (red), drive artificial miRNA expression. The
dsRED2 fluorescent protein is expressed strongly from the CMV promoter.
This plasmid may be used in the packaging of AAV vectors, including
BAAV.
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Figure 22. In vitro artificial miRNA silencing of GJB2-R75W transgene. GJB2-R75W
expression results were normalized to levels of CHO-Hprt internal control and
knockdown levels of GJB2-R75W were normalized to negative control
artificial miRNA (pAAV_H1-miNC1) treated cells = 100% expression (0%
knockdown). Results for knockdown of the GJB2-R75W transgene by the
artificial miRNA constructs were: pAAV_H1-miRNA2 (20% ± 5%
knockdown), pAAV_H1-miGJB2-C (18% ± 15%), pAAV_H1-miGJB2-D
(20% ± 8%), pAAV_tVal-miRNA2 (1% ± 9%), pAAV_tVal-miGJB2-C (33%
± 12%), and pAAV_tVal-miGJB2-D (42% ± 11%).
92
Figure 23. In vitro artificial miRNA silencing by pAAV_tVal-miRNAs. Results for
knockdown of the GJB2-R75W transgene by the artificial miRNA constructs
were: pAAV_tVal-miRNA2 (124% ± 23%), pAAV_tVal-miGJB2-C (140% ±
7%), and pAAV_tVal-miGJB2-D (18% ± 13%). Expression levels were
normalized to negative control artificial miRNA (pAAV_H1-miNC1) treated
cells = 100% expression (0% knockdown). 82% knockdown was the highest
silencing achieved by any of the artificial miRNA constructs in any of the
repeated assays. The varied data suggests a high degree of variability in the
experimental procedure.
93
Figure 24. Experimental outline for in vitro silencing assays.
94
Figure 25. HsGJB2-MmGjb2 sequence alignment and siRNA target sites. HsGJB2 and
MmGjb2 coding sequences share 87% nucleotide homology. siRNAs were
designed to target regions of variation in order to confer silencing specificity
to the HsGJB2-R75W transgene. Sequences of the top three siRNAs are
indicated: siRNA2 (yellow highlight), GJB2-C (red), GJB2-D (underline).
GJB2-C and GJB2-D target sites overlap.
95
CHAPTER 5: VIRAL VECTOR TROPISM FOR SUPPORTING CELLS
IN THE DEVELOPING MURINE COCHLEA
Abstract
Gene-based therapeutics are being developed as novel treatments for genetic
hearing loss. One roadblock to effective gene therapy is the identification of vectors
which will safely deliver therapeutics to targeted cells. The cellular heterogeneity that
exists within the cochlea makes viral tropism a vital consideration for effective inner ear
gene therapy. There are compelling reasons to identify a viral vector with tropism for
organ of Corti supporting cells. Supporting cells are the primary expression site of
connexin 26 gap junction proteins that are mutated in the most common form of
congenital genetic deafness (DFNB1). Supporting cells are also primary targets for
inducing hair cell regeneration. Since many genetic forms of deafness are congenital it is
necessary to administer gene transfer-based therapeutics prior to the onset of significant
hearing loss. We have used transuterine microinjection of the fetal murine otocyst to
investigate viral tropism in the developing inner ear. For the first time we have
characterized viral tropism for supporting cells following in utero delivery to their
progenitors. We report the inner ear tropism and potential ototoxicity of three previously
untested vectors: early-generation adenovirus (Ad5.CMV.GFP), advanced-generation
adenovirus (Adf.11D) and bovine adeno-associated virus (BAAV.CMV.GFP).
Adenovirus showed robust tropism for organ of Corti supporting cells throughout the
cochlea but induced increased ABR thresholds indicating ototoxicity. BAAV also
showed tropism for organ of Corti supporting cells, with preferential transduction toward
the cochlear apex. Additionally, BAAV readily transduced spiral ganglion neurons.
Importantly, the BAAV-injected ears exhibited normal hearing at 5 weeks of age when
compared to non-injected ears. Our results support the use of BAAV for safe and
efficient targeting of supporting cell progenitors in the developing murine inner ear.
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Introduction
After selecting an animal model and developing a therapeutic agent, a method is
needed to target the therapeutic to the affected tissue. There are a number of surgical
approaches that have been developed to provide access to the inner ear. Likewise,
numerous viral vectors have been studied as vectors for gene delivery to the cochlea. The
various delivery routes and vectors have different safety profiles and allow targeting of
specific cell types. In designing gene therapy, it is important to select a delivery
technique and a delivery vector that are most appropriate for targeting the correct cell
type at the correct stage of development. This chapter will discuss the route of delivery
and the vector for delivery.
Methods of Cochlear Delivery
The inner ear is a particularly suitable organ for delivery of therapeutic molecules
as its bony capsule within the temporal bone provides relative isolation from the rest of
the body but is accessible through the middle ear. A variety of delivery techniques have
been described for the cochlea [138-142]. Delivery techniques can directly or indirectly
access the perilymphatic or endolymphatic systems and different approaches result in
different cochlear effects.
A number of studies have been conducted on rodents in which the perilymphatic
system was approached by direct microinjection to the scala tympani through the round
window membrane (RWM), via a cochleostomy or through the posterior semicircular
canal. Because of the anatomical continuity of membranous perilymphatic space, these
approaches primarily allow transduction in fibrocytes and mesothelial cells lining the
scala tympani, although in some experimental conditions transduction is also seen in the
area surrounding the scala media.
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Surgical approaches to the endolymphatic sac allow drug delivery to the
endolymphatic system, resulting primarily in transduction in the endolymphatic duct
epithelium and transitional epithelium in the utricle and saccule.[141, 143, 144]
Differences in reported sites of expression between experiments may be due to
differences in rodent models, titers and doses of vector (single injection vs. continuous
infusion using an osmotic mini-pump), particle size, presence or absence of viral
receptors, and time points of sacrifice of the drug-treated animals.
The ear is sensitive to insult and there are potential complications associated with
gene delivery into the cochlea regardless of the route of delivery, including iatrogenic cell
damage from hydropressure and ototoxicity of the delivery vector. For example, after
cochleostomy and injection to the rodent scala tympani, damage of inner ear cells and an
inflammatory response are occasionally seen.[145] The damage is usually confined to the
basal turn of the cochlea or the site of injection at the RWM.
A less traumatic method was studied by Jero and colleagues who demonstrated
the feasibility of diffusion of liposome-complexed plasmid and AV vector through the
intact RWM of mice accessed via a ventral surgical approach.[146] These investigators
were able to observe GFP and β-gal expression in the spiral limbus, spiral ligament,
sensory and supporting cells of the organ of Corti, Reissner’s membrane and spiral
ganglion cells 3–7 days after placing gelfoam soaked with a lipocomplexed plasmid
directly on the RWM.[139] Our lab used a similar approach to introduce a
cytomegalovirus (CMV)-driven, dominant-negative GJB2-R75W mutant construct into
the inner ear and also observed expression in the spiral limbus, spiral ligament, epithelial
cells in the basilar membrane, outer hair cells, inner and outer pillar cells, and Claudius
cells in the organ of Corti.[102] Clearly, an indirect approach through the intact or
chemically permeabilized RWM reduces iatrogenic damage, and this technique is one of
the most suitable for accessing the adult rodent cochlea. One drawback, however, is that
transgene expression is relatively low and primarily in the basal turn of the cochlea.
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Each of the above-mentioned delivery techniques were designed to access the
adult rodent cochlea. However, many genetic forms of deafness are congenital due to
developmental abnormalities of the cochlea that ultimately lead to dysfunction and death
of the sensory hair cells. Gene transfer-based therapies for these forms of hearing loss
require intervention prior to cochlear maturation. Several years ago, Bedrosian and
colleagues developed and optimized a method of in utero gene transfer into the
developing mouse inner ear (otocyst) at embryonic day (E) 12-12.5 mouse (Figure
26).[138] They showed that this was a safe and effective method to target progenitor cells
in the developing cochlea.
Mouse models of connexin-related deafness, including the GJB2-R75W
transgenic mice, display significant hearing loss at the time of cochlear maturation
meaning therapeutic strategies must be targeted at the developing cochlea [13, 47, 54,
147, 148]. We selected the embryonic otocyst injection technique as the delivery method
of choice because it allows us to safely target the therapeutic to the cochlea at its earliest
stages of development, prior to onset of pathology. Although the procedure is both
logistically and technically challenging, Dr. Samuel Gubbels (Division of
Otolaryngology, University of Wisconsin) generously shared his expertise and brought
the technique to the University of Iowa (Figure 27).[149] During the course of my
studies, I also became expert at this procedure.
Viral Vectors
Another key component is selection of the appropriate delivery vector. Although
transient transfection can be achieved by plasmid delivery, long-term in vivo silencing of
an endogenous gene by RNAi requires a different strategy, typically using a viral
vector.[99]
Two major criteria must be evaluated in the selection of an optimal viral vector
for cochlear application: tropism and toxicity. A vector’s tropism refers to the cell type
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which it preferentially transduces. Given the variety of cell types in the inner ear, viral
tropism is an important consideration when selecting a vector for gene transfer [13]. As
some viruses have detrimental effects on their targets, the second major consideration is
toxicity. To date, a variety of viral vectors have been used to target genes in the inner ear,
including adenovirus (AV), adeno-associated virus (AAV), herpes simplex virus (HSV)
and lentivirus (LV) [13, 140, 150-152]. Each virus demonstrated a different tropism and
toxicity profile.
Adeno-Associated Virus
Adeno-associated virus (AAV), a member of the parvoviridae family, is a small,
single-stranded DNA virus with maximum packaging capacity of ~4.5 kb. While
relatively small packaging capacity is its major disadvantage, there are many advantages
to its use in gene therapy. Not associated with any human disease, AAV is generally
considered to be safe.[13] It is capable of infecting most tissues and cell types with
potential for long-term expression. AAV vectors do not express any native viral genes
and the capsid displays very low immunogenicity which limits the potential for
cytotoxicity.[153, 154] Additionally, a number of available AAV serotypes display
unique tropism and offer the opportunity to target specific tissues and cell types.[153,
155] Unique tropisms and transduction efficiencies have been reported in the cochlea,
depending on AAV serotype and model system.[154] Serotype AAV2/1 effectively
transduced the sensory hair cells when delivered to the developing otocyst, as Bedrosian
and colleagues initially reported and we have confirmed.[138]
Adenovirus
AV is a non-enveloped, double-stranded DNA virus that has been used
extensively for gene therapy applications. It is a relatively benign human pathogen that is
made replication-defective by deletion of the early gene, E1.[156] AV vectors were the
first to be evaluated for in vivo gene transfer in a wide variety of preclinical models
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because of a number of features which make it well-suited for gene transfer
purposes.[157] One advantage of AV vector is the ability to infect both dividing and nondividing cells with high efficiency. Their broad host range and rapid onset of gene
expression (~2 days) following transduction make them suitable to a variety of
applications. The AV genome also provides a relatively large packaging capacity (up to
10 kb depending on how much of the native genome is retained) and remains as a nonintegrating nuclear episome, minimizing the risk for insertional mutagenesis.[13, 156]
Some characteristics of AV, however, may preclude its usefulness in certain gene therapy
applications. Failure to integrate into the target genome leads to transient expression
which may limit its efficacy as a long term therapeutic vehicle.[157] Most significant,
early generation (vectors carrying deletions in E1 and E3 genes) AV can induce a potent
immune response that may result in cytotoxicity and reduced transgene expression.[156]
However, late-generation (E4-region is also removed) adenovirus serotype 5 (AV5) has
been used effectively to deliver genes to the adult rodent cochlea and to be safer than the
earlier generation AVs.[13, 154] A variety of cochlear cell types have been transduced,
including sensory and non-sensory cells of the organ of Corti, fibrocytes in the spiral
ligament and spiral ganglion neurons (SGNs).[152, 158-162]
Adenovirus attaches to cells via the coxsackie adenovirus receptor (CAR). CAR is
expressed on most cochlear cell types of the neonatal mouse and continues to be
expressed on the supporting and strial cells of the adult mouse cochlea.[163, 164]
Delivery of AV via cochleostomy to the scala media results in high efficiency
transduction of all organ of Corti supporting cell types.[165] AV has never before been
delivered in vivo to the embryonic otocyst.
Bovine Adeno-Associated Virus
Bovine adeno-associated virus (BAAV) is a novel, non-primate adeno-associated
virus which has recently been used for gene transfer to the rodent cochlea.[49, 166, 167]
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Although it displays 79% nucleotide homology with AAV5 and 76% capsid protein
homology with AAV4, BAAV is serologically distinct and shows a unique tropism
profile when compared with AAV2, 4, or 5.[168] Recombinant BAAV was shown to be
11 times more efficient at gene transfer to a murine salivary gland than was AAV2.[168]
BAAV has been recently used for effective in vitro transduction of cultured rat inner ear
epithelia and gene transfer to murine cochlear organotypic cultures.[49, 166] In these
cochlear cultures, BAAV approached 100% transduction of supporting cells, with
preference for cells of the outer sulcus. In vivo delivery of BAAV to the adult guinea pig
cochlea resulted in safe gene transfer to supporting cells, with occasional outer hair cell
transduction also reported.[167]
In order to verify the previously published results and optimize our technique,
initial otocyst injections performed in our lab used AAV2/1. We observed safe and highly
efficient transduction of sensory hair cells, similar to the previous report.[138] While in
utero targeting of sensory hair cells with AAV2/1 provides therapeutic potential for many
types of genetic hearing loss, there are important reasons to identify a virus with tropism
for supporting cells in the developing cochlea. Aside from their role in the most common
form of ARNSHL, cochlear supporting cells serve as the primary targets of intervention
to induce hair cell regeneration [152]. One significant advance in the application of gene
therapy to restore auditory function was the discovery that adenoviral delivery of Atoh1
(Math1) to supporting cells in the guinea pig cochlea resulted in the formation of “hair
cell-like” cells [13, 152]. The promise of hair cell regeneration and the potential
treatment of connexin-related deafness disorders, are compelling reasons to identify safe
vectors with tropism for inner ear supporting cells [149, 169].
AV and bovine adeno-associated virus (BAAV) vectors have been previously
used for successful transduction of cochlear supporting cells in adult rodents [152, 165,
170], but have never been delivered to the developing cochlea in vivo. We hypothesized
that AV and BAAV would efficiently target supporting cells of the organ of Corti. In this
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chapter I discuss our use of the transuterine microinjection approach to examine the
safety and inner ear tropism of three previously untested vectors delivered to the
developing murine cochlea: Ad5.CMV.GFP (early-generation AV), Adf.11D (lategeneration AV) and BAAV.CMV.GFP (BAAV).
Materials and Methods
Virus Production
Early-generation replication-deficient adenovectors (deleted of E1A and a partial
deletion of E1B and E3) were constructed in the Gene Transfer Vector Core Laboratory
at the University of Iowa as described previously [171]. The green fluorescent protein
(GFP) reporter gene expression was driven by the human cytomegalovirus (CMV)
promoter (Ad5.CMV.GFP). Recombinant Ad5.CMV.GFP viruses were grown in human
embryonic kidney (HEK) 293 cells that complement the E1 early viral promoters. Virus
titers of 1.0^11 plaque-forming units per milliliter (pfu/mL) were suspended in
phosphate-buffered saline (PBS) with 3% sucrose and stored at −80°C until use.
An advanced-generation adenovirus vector, Adf.11D, was provided by Dr.
Douglas Brough at GenVec (Gaithersburg, MD, USA). The E1/E3/E4-deleted
adenovector contains an expression cassette of GFP driven by a human CMV promoter.
Construction and production of Adf.11D has been reported previously [172]. Total
particles (pu) are analyzed by a spectrophotometric assay that has been standardized to
reliably quantify the total particles within a lot of adenovector. The lot of Adf.11D used
for these experiments measured 9.6 x 10^11 total particles per milliliter (pu/mL), with
10.7 x 10^10 focal forming units per mL (ffu/mL), a measure of adenovector activity.
Adf.11D was purified and aliquots stored at −80°C until use.
Recombinant BAAV expressing GFP driven by a human CMV promoter was
provided by Drs. John A. Chiorini and Giovanni Di Pasquale at the National Institutes of
Health (Bethesda, MD, USA). The vector was produced as previously described, purified
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using CsCl gradients and particle titers were determined by qPCR [166, 173]. The
recombinant BAAV particle titers were 5.0 x 10^11 DNase-resistant particles per
milliliter (DRP/mL). Purified BAAV.CMV.GFP was dialyzed in PBS and stored at
−80°C until use.
Animals and Virus Administration
All procedures were performed in adherence with institutional and national
guidelines and were approved by the Institutional Animal Care and Use Committee at the
University of Iowa. 6-week to 3-month old BALB/C mice maintained at the University of
Iowa animal care facility underwent timed breeding to generate the fetuses for
transuterine microinjection. For aging of the embryos, noon of the cervical plug date was
considered to be day E0.5.
Transuterine Otocyst Microinjection
Injection of vectors into the left otocyst of E12.5 mouse embryos was performed
via transuterine microinjection as described previously [138, 149]. Briefly, E12.5 dams
were anesthetized with 9 mg/mL Nembutal at a dose of 7μl/g body weight. Using a Leica
M220 F12 surgical microscope (Leica Microsystems Inc., Bannockburn, IL, USA), a 1.5cm midline laparotomy incision was made and the uterine horns were exposed. Embryos
were trans-illuminated to identify anatomical landmarks of the otocyst. Prior to delivery,
8 µL of vector was mixed with 4 µL of 2.5% fast green dye in PBS and loaded into a
specially pulled glass micropipette [149]. Approximately 50 nl of this vector preparation
(Ad5.CMV.GFP, Adf.11D, or BAAV.CMV.GFP) was delivered to the left otocyst of
accessible embryos (average of 3 to 5 embryos per dam) using a glass micropipette, M33
roller bearing micromanipulator (Stoelting Co., Wood Dale, IL, USA) and PLI-100 picoinjector pump system (Harvard Apparatus Inc., Holliston, MA, USA). Contralateral
(right) otocysts were not injected and served as non-injected controls. Uterine horns were
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replaced inside the peritoneal cavity, incisions sutured, and dams allowed to give birth
vaginally and to nurse their young to maturity.
Auditory Brainstem Response (ABR) Testing
At 5 weeks of age, mice were anesthetized by intraperitoneal injection of (10μl/g
body weight) [10mg/mL] ketamine/xylazine and placed in an acoustic test chamber
(Acoustic Systems, Glendale Heights, IL, USA) for testing of auditory-evoked brainstem
response (ABR) thresholds. ABR testing was carried out as previously described [174].
Briefly, after anesthetic has taken effect, active, reference and ground electrode needles
are inserted subcutaneously and the SmartEP software system (Intelligent Hearing
System Inc., Miami, FL, USA) used for the stimulus presentation, ABR acquisition, and
data management. The first stimulus is presented at an intensity of 70 decibels sound
pressure level (dB SPL) and, depending upon the response, is followed by increasing or
decreasing the decibel level, initially in 10 dB- and subsequently in 5 dB-increments, to
determine the auditory threshold (the lowest sound level at which an ABR pattern is
recognizable). Click-stimulus ABR testing presents clicks that cover a range of
frequencies (2-8 kHz) and is used for general screening of auditory function. Pure-tone
ABR testing presents a series of tones to test ability to hear at specific frequencies: 4, 8,
16, and 32 kHz for these experiments.
Sample Preparation and Immunofluorescent Imaging
At P35 anesthetized mice were sacrificed and the inner ears were isolated,
immediately fixed for two hours in 4% PFA, rinsed in PBS, decalcified in EDTA and
prepared for cryosectioning as previously described [69]. 10μm mid-modiolar
cryosections were prepared for immunohistochemistry. For surface preparations of the
sensory epithelium, P5-P7 mouse pups were decapitated, the cochleae dissected, and the
sensory epithelium isolated and placed on a glass slide as previously described [175]. The
sensory epithelium was immediately fixed in 4% PFA for 20 minutes, rinsed in PBS, and
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stored at 4°C in preparation for immunohistochemistry. Immunohistochemistry was
performed according to a previously established methodology [69]. For anti-GFP
immunostaining, sections from each of the left cochleae and several of the right cochleae
were incubated with a 1:500 dilution of rabbit polyclonal anti-GFP primary antibody
(#AB3080 Lot LV1407277; Chemicon, Temecula, CA) followed by incubation with a
1:1000 dilution of a secondary Alexa Fluor 568-conjugated goat anti-rabbit antibody
(#A11036, Invitrogen, Carlsbad, CA, USA). Immunofluorescence imaging using a Leica
DM IRE2 fluorescent microscope (Leica Microsystems Inc., Bannockburn, IL, USA) was
performed to verify GFP expression. Sections from confirmed GFP-positive cochleae
were incubated with a 1:150 dilution of rabbit polyclonal anti-myosin VIIA primary
antibody (#25-6790, Proteus Biosciences Inc., Ramona, CA, USA) which is specific for
inner and outer hair cells. This was followed by incubation with Alexa Fluor 568
antibody as described above. All slides were treated with ProLong Gold anti-fade reagent
with DAPI (#P36931; Invitrogen, Carlsbad, CA, USA), followed by placement of a
coverslip and allowed to sit for 24 hours prior to imaging. Immunofluorescence imaging
using the Leica DM IRE2 fluorescent microscope or a Leica TCS SP5 confocal
microscope (Leica Microsystems Inc., Bannockburn, IL, USA) was used to identify
transduced cell types.
Results
In utero Delivery of Adenovirus to the Developing Murine
Inner Ear
We performed transuterine microinjection of the murine otocyst with earlygeneration (E1/E3-deleted) adenoviral (AV5) vector expressing green fluorescent protein
(GFP) under the control of a human cytomegalovirus (CMV) promoter (Ad5.CMV.GFP).
Otocyst injections were performed on E11.5-12.5 mouse embryos as described by
Bedrosian et al. [138, 149]. Initial screening of injected embryos demonstrated robust
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transduction of sensory epithelium at 1 week after birth on postnatal day 7 (P7) (Figure
28). Next, we assessed expression in injected mice (n=4) at 5 weeks of age
(approximately P35). We observed robust transduction in the organ of Corti with strong
tropism for supporting cells (Figure 29a-f). Transduced cells were also observed in the
stria vascularis, the supralimbal region of the spiral limbus, Reissner’s membrane, and
spiral ganglion neurons (SGNs) (Figure 29a-c,g). We observed no evidence of immune
cell infiltrates in histologic cross-sections. None of the cochleae showed significant or
consistent vestibular expression of GFP, nor did any of the mice display circling or headtilting behavior which would indicate vestibular dysfunction.
Effect of in utero Adenoviral Transduction on Hearing
We assessed the hearing of injected mice at 5 weeks of age using click-stimulus
auditory brainstem response (ABR) testing to determine if transduction of cochlear cells
with AV was associated with ototoxicity. Using the two-sample t-test for equal variances
we determined that hearing thresholds of the ears injected with Ad5.CMV.GFP were
significantly elevated (97.5dB ± 5, n=4) when compared with contralateral non-injected
ears (45.0dB ± 0, n=4; t(6)=21.0, p=7.6x10-7) and with non-injected littermates (45.8dB ±
7.4, n=6; t(8)=12.2, p=1.9x10-6). These results suggest that Ad5.CMV.GFP transduction
is strongly ototoxic to cells of the developing cochlea (Figure 30).
Because early-generation AVs have been reported to be ototoxic to hair cells in
vivo as assessed by distortion-product otoacoustic emission (DPOAE) testing, we
hypothesized that the use of advanced-generation (E1/E3/E4-deleted) AV would result in
diminished ototoxicity. Following the same protocol as with Ad5.CMV.GFP, we injected
9 embryos with Adf.11D, an E1/E3/E4-deleted adenovector expressing GFP from a CMV
promoter. GFP expression at 5 weeks of age was localized to the organ of Corti region.
However, expression was less intense and more variable than with Ad5.CMV.GFP,
making it difficult to definitively identify transduced cell types (data not shown). We
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again performed ABR testing at 5 weeks of age and as with the early-generation AV, the
two-sample t-test for unequal variances showed that hearing thresholds of the ears
injected with Adf.11D were significantly elevated (99.4dB ± 19.8, n=9) when compared
with contralateral non-injected ears (56.1dB ± 15.4, n=9; t(15)=5.19, p=1.0x10-4) and
with non-injected littermates (51.3dB ± 4.3, n=20; t(8)=7.24, p=8.9x10-5) (Figure 30).
These results indicate that both early- and advanced-generation AV vectors induce
ototoxicity when delivered to cells of the developing murine cochlea.
In utero Delivery of Bovine Adeno-Associated Virus to the
Developing Murine Inner Ear
We next tested bovine adeno-associated virus (BAAV) vector expressing GFP
under the control of a CMV promoter (BAAV.CMV.GFP), initially injecting the left ear
in two embryos and assessing cochlear expression at 5 weeks of age. We observed strong
transduction in the organ of Corti with tropism for supporting cells (Figure 32 a-f).
Although transduction occurred in all turns of the cochlea, we noted preferential
transduction toward the apex.
To confirm these findings we injected six additional embryos with
BAAV.CMV.GFP. Two of the injected mice were sacrificed on postnatal day 5 (P5) and
a whole mount preparation of the sensory epithelium was visualized to determine
transduction efficiency along the length of the cochlea (Figure 33). GFP-positive cells
were counted along the entire length of each cochlea. The mean transduction efficiency
was 29.4 ± 16.6 (ranging from 3 to 56) transduced cells/500µm. To assess whether a
statistically significant difference in transduction occurred along the length of the cochlea
we measured the average number of transduced cells in 1 mm segments from the most
apical, middle and most basal regions of the cochleae. The average transduction
efficiency for each of these segments was 39.8 ± 5.3, 29.5 ± 13.1, and 8.3 ± 5.6
transduced cells/500µm in the apical, middle, and basal cochlear turns, respectively
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(Figure 34d). The mean transduction efficiencies at the apex and at the middle of the
cochlea are each significantly higher than the efficiency at the base according to the twosample t-test for equal variances (Apex: t(6) = 8.24, p = 1.7x10-4; Middle: t(6) = 2.99, p =
0.02). The transduction efficiency between the apex and the middle were not significantly
different (t(6) = 1.45, p > 0.05). The trend of preferential transduction toward the apex is
appreciated by graphs showing the number of transduced cells along the linear distance
of individual cochlear preparations (Figure 34c).
Next we analyzed mid-modiolar cross-sections of the injected cochleae in order to
evaluate transduced cell types. GFP expression was predominantly localized to the organ
of Corti region with tropism for supporting cells (pillar and Deiters cells, inner and outer
sulcus cells) (Figure 4c,f). A small number of inner and outer hair cells were also
transduced in each cochlea (Figure 32c). Sporadic transduction was seen in cells of the
spiral prominence, Reissner’s Membrane and stria vascularis. Spiral ganglion neurons
(SGNs) were transduced with high efficiency (51.1% ± 8.0%) (Figure 32g).
Although we observed no apparent structural abnormalities or evidence of
immune cell infiltrates in the transduced cochleae, we screened the hearing of injected
mice at 5 weeks of age. Hearing thresholds determined by click-stimulus ABR showed no
significant difference (p>0.05) between injected ears (43.5dB ± 7.5, n=4), contralateral
non-injected ears (46.3dB ± 4.8, n=4) and non-injected littermate ears (44.0dB ± 5.7,
n=12) (Figure 3a). Click-stimulus ABR assesses hearing across a range of frequencies.
Hair cells in the cochlea are tuned to respond maximally to different sound frequencies
depending on their position along the length of the cochlea. The apex of the cochlea is
attuned to respond to low frequency stimulation and the base to high frequency
stimulation. Because of the linear transduction gradient observed with BAAV we tested
pure-tone ABRs at 4, 8, 16, and 32 kHz (Figure 31). There was no significant difference
(p>0.05) in hearing threshold at any of the tested frequencies between injected ears (4kHz
= 88.3dB ± 2.6, 8kHz = 54.2 ± 2.0, 16kHz = 35.8 ± 2.0, 32kHz = 34.2 ± 5.8; n=6),
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contralateral non-injected ears (4kHz = 88.3dB ± 2.6, 8kHz = 55.0 ± 3.2, 16kHz = 35.8 ±
2.0, 32kHz = 37.5 ± 5.2; n=6), and non-injected littermate ears (4kHz = 89.5 ± 3.2, 8kHz
= 54.8 ± 3.4, 16kHz = 36.3 ± 3.9, 32kHz = 35.8 ± 6.3; n=20). These results indicate that
BAAV transduction of cells of the developing cochlea is safe to at least 5 weeks of age.
Discussion
Gene therapy offers a promising means of modulating gene expression in the
inner ear with the ultimate goal of altering the deafness phenotype. Viral tropism and the
potential for ototoxic effects are important considerations when selecting a vector for
gene transfer. Bedrosian and colleagues have identified AAV2/1 as a safe vector with
tropism for sensory hair cells of the organ of Corti when delivered to the embryonic
otocyst [138]. Here we report cellular tropism and potential ototoxicity profiles of earlyand advanced-generation adenoviral vectors as well as bovine adeno-associated virus.
Our data for AV are consistent with previous in vitro and in vivo studies that show
tropism for supporting cells [141, 152, 165, 176]. The ototoxicity we observed with
Ad5.CMV.GFP delivery is also consistent with reports of ototoxicity associated with
delivery of early-generation AV [154]. However, in contrast to reports in which
E1/E3/E4-deleted adenovectors were safely delivered to the inner ear of adult mice [177],
delivery of Adf.11D to the E12.5 otocyst resulted in significant ototoxicity. Potential
reasons for the ototoxicity include: pressure from the injection, GFP expression,
triggering of immune responses, high concentration of AV delivered, and increased
sensitivity of the rapidly developing otocyst cells to adenovirus.
Pressure toxicity, or toxicity from the otocyst injection technique is not likely
because each of the injections for AV as well as BAAV was performed by the same
person following an identical protocol that allows for safe delivery to the otocyst [149].
In the report by Bedrosian and colleagues lentivirus had ototoxic effects while none of
the pseudotyped AAVs were associated with toxicity [138]. There are reports that GFP
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can be toxic to living cells [178], but there are currently no reports of GFP-induced
ototoxicity. Viruses expressing GFP from a CMV promoter have been safely delivered to
the cochlea of adult rodents [177], Atoh1-GFP plasmid constructs have been injected and
electroporated into the developing murine otocyst without detriment to hearing [179].
AV is known to induce a potent immune response that may produce ototoxicity
[13, 154, 180]. Early-generation adenovectors deleted of the E1/E3 regions have been
shown to be toxic to cochlear hair cells in vivo, likely related to immune responses [154].
To overcome this toxicity, later generation “gutted” adenovectors that contain only
essential packaging sequences and have no immunostimulatory effects were generated
[154, 156]. Advanced-generation adenovectors deleted of the E1/E3/E4 regions have
been reported to be less immunogenic than earlier generations. The E4 region of AV
encodes proteins that modulate function of the host cells and removal of this region leads
to decreased cytotoxicity and immunostimulation [154, 156, 177]. Thus, we were
surprised by the ototoxicity observed following delivery of Adf.11D. In histological
cross-sections of the cochlea we observed no evidence of immune cell infiltrates.
Additionally, the blood-labyrinth barrier maintains the inner ear as a relatively immunoprotected region, minimizing concern for immune-mediated complications in the intact
cochlea [181]. Ototoxicity may also depend on the titer of virus delivered, with lower
titers being safer but achieving less robust transduction [182, 183]. Because of the limited
volume of the otocyst, we delivered high titer AV (10.7 x 10^10 ffu/mL). It may be
possible to achieve sufficient transduction of the organ of Corti and diminish ototoxic
effects by using lower titers of virus. However, the dynamic range for in vivo AV
transduction in the cochlea is reportedly narrow and significantly reducing the titer may
result in negligible transduction [154].
Bovine adeno-associated virus (BAAV) has recently been used in vivo to safely
transduce cells in the adult guinea pig cochlea [167]. We were able to recapitulate these
results when BAAV was delivered to the murine otocyst. BAAV expression after
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embryonic delivery showed consistent tropism for the organ of Corti and for spiral
ganglion neurons (SGNs). In the organ of Corti, supporting cells were preferentially
transduced with occasional transduction of inner and outer hair cells. Other cochlear cell
types were sporadically transduced. While transduction occurred in each turn of the
cochlea, we observed a preference for transduction toward the apex. Both click-stimulus
and pure-tone ABR tests indicated functionally normal hearing in the BAAV-treated ears
at 5 weeks of age. Additionally, none of the injected mice displayed phenotypes
indicative of vestibular dysfunction prior to sacrifice.
The apical transduction preference is unique to the transuterine delivery of BAAV
to the otocyst. No apical preference has been previously observed with BAAV delivery in
vivo or in vitro. Neither has an apical preference been reported for any other vector
delivered via transuterine injection to the otocyst, including AAV2/1, lentivirus, and AV.
This finding suggests that the observed apical preference is a consequence of properties
unique to BAAV transduction of the developing cochlea.
Cell surface receptors are an important determinant of virus entry, infection, and
tropism [184]. AAVs are known to utilize a variety of cell surface carbohydrates as
attachment receptors. For example, AAV2 binds heparan sulfate proteoglycans (HSPGs),
while AAV4 and AAV5 each recognize different forms of sialic acid for cell attachment
and transduction [185, 186]. The platelet-derived growth factor receptor (PDGFR), an
integral membrane glycoprotein with a terminal 2-3-linked sialic acid, is a co-receptor for
AAV5 transduction [187]. BAAV has unique tropism compared with other AAVs and is
able to transduce cells of the inner ear with high efficiency [49, 166, 167]. BAAV
transduction appears to occur in a two-step process: cell-binding followed by cell entry.
Non-terminal sialic acid groups mediate cell binding and terminal sialic acid groups are
essential for cell entry and transduction [188]. Unlike AAV5, which uses a glycoprotein
as its receptor, BAAV requires glycosphingolipids (GSLs) for transduction. Gangliosides,
glycosphingolipids containing sialic acid, are essential for BAAV entry into cells [188].
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Multiple gangliosides are expressed in the mammalian cochlea [189-191]. GM3, the most
abundant ganglioside in the inner ear, is the first product in the biosynthetic pathway of
the ganglio-series gangliosides and serves as a common precursor to many other
gangliosides. GM3 and GT1b, another ganglioside expressed in the inner ear, are found
throughout the cochlea at P3 but after P14 localize to specific regions in the inner ear
including the organ of Corti [191].
The most likely cause of the apical transduction preference observed with BAAV
is differential expression of cellular BAAV receptors during cochlear development. There
is ample evidence for differential expression gradients along the length of the cochlea
during development. Terminal mitosis of cells in the organ of Corti begins apically and
proceeds towards the base between E12.5 and E18.5 [192]. Cytological differentiation of
the organ of Corti occurs in the opposite direction, beginning in the base of the cochlea
and proceeding apically [193]. A cDNA microarray analysis identified 141 genes that
were differentially expressed between the apex and the base of the mouse cochlea [194].
Individual gangliosides have unique spatiotemporal distribution within the cochlea and
their localization is known to dramatically change during early postnatal development
[191]. The transduction gradient observed with BAAV is likely a result of the
spatiotemporal distribution pattern of cellular attachment and internalization receptors
during cochlear development. In order to better understand BAAV’s apical transduction
preference in the developing cochlea it will be important to identify specific cellular
receptors for BAAV transduction and determine the expression patterns of these
receptors during development.
In summary, we have shown our ability to use the embryonic otocyst injection
technique for delivering a variety of viral vectors to the developing murine cochlea. We
have shown that BAAV has minimal impact on inner ear cell activity compared to AV. It
displays strong tropism for inner ear sensory epithelium and has the ability to penetrate
barrier epithelial cells via transcytosis [49]. These properties, together with BAAV’s safe
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and efficient transduction of organ of Corti cells via embryonic otocyst injection, make
BAAV an attractive vector for delivery of long-term gene therapy to the inner ear to treat
connexin-related (i.e. DFNB1) and other forms of deafness associated with pathology of
cochlear supporting cells.
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Figure 26. Microinjection into the developing mouse otocyst. The ex utero approach as
seen through the surgical microscope. (a) The uterus is exposed by a midline
laparotomy and the fetuses appear as a string of beads. The uterine wall
(arrow) is incised to expose the fetus. (b) The microinjection pipette (arrow)
enters the amniotic sac. The triangular injection site is demarked by the
cardinal veins (v) and fourth ventricle (IV). (c) Dorsal view of the embryo at
embryonic day 12 (E12). The left eye (arrow) and fourth ventricle are visible.
Following injection, fast green dye is visible in the otocyst indicating
successful injection. (d) Left sagittal profile of the embryo illustrating the
endolymphatic duct (arrow) protruding away from the otocyst at the 1 o'clock
position. Adapted from Hildebrand et al. 2008.
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Figure 27. Surgical room setup. Equipment for otocyst microinjections (Department of
Otolaryngology, University of Iowa).
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Figure 28. Adenovector transduction of inner ear. Robust transduction of the mouse
cochlear sensory epithelium at age P7 following in utero microinjection of
Ad5.CMV.GFP at E12.5 (a,b).
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Figure 29. Adenoviral transduction of cochlear cells in the adult mouse inner ear.
Microinjection of Ad5.CMV.GFP into the mouse otocyst was performed at
E12.5 and injected mice were sacrificed at 5 weeks. Transduction of cochlear
cells is still evident six weeks after injection of Ad5.CMV.GFP as shown by
GFP fluorescence (a) and immunofluorescence with an anti-GFP antibody (b)
in mid-modiolar cross section of the cochlea. A merged image is shown in (c).
Cells lining the scala media cavity (c) were predominantly transduced by the
AV with sparse transduction of spiral ganglion neurons (data not shown).
Targeted transduction of supporting cells in the organ of Corti is shown by
GFP fluorescence (d). Immunofluorescence with an anti-myosin VIIA
antibody was conducted to specifically mark the inner and outer hair cells (e).
A merged image is shown in (f). Dieters cells that support the outer hair cells
appear to be readily transduced. Adf.11D expression was also localized to the
organ of Corti, but was less intense than Ad5.CMV.GFP expression (data not
shown). IHC = inner hair cell; OHC = outer hair cell; DC = Deiters
(supporting) cells; SV = stria vascularis; SL = spiral limbus; RM = Reissner’s
membrane; IS = inner sulcus
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119
Figure 30. Auditory assessment of mice injected with viral vectors. Five-week old mice
were anesthetized and placed in an acoustic test chamber for measurement of
auditory-evoked brainstem response (ABR) thresholds. Average clickstimulus hearing thresholds of mice injected with virus compared with noninjected contralateral ears and non-injected littermates, respectively:
Ad5.CMV.GFP (97.5dB ± 5, n=4; 45dB ± 0, n=4; 45.8dB ± 7.4, n=6),
Adf.11D (99.4dB ± 19.8, n=9; 56.1dB ± 15.4, n=9; 51.3dB ± 4.3, n=20),
BAAV.CMV.GFP (43.5dB ± 7.5, n=4; 46.3dB ± 4.8, n=4; 44dB ± 5.7, n=12).
Using the two-sample t-test, both early- and advanced-generation AV injected
ears had significant ototoxicity (indicated by asterisks) compared with
contralateral non-injected ears (Ad5.CMV.GFP: t(6)=21.0, p=7.6x10^-7;
Adf.11D: t(15)=5.19, p=1.0x10^-4), while BAAV injections were nonototoxic (p>0.05).
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Figure 31. Pure tone ABR thresholds of mice injected with BAAV. Five-week old mice
were anesthetized and placed in an acoustic test chamber for measurement of
auditory-evoked brainstem response (ABR) thresholds. Average clickstimulus hearing thresholds of mice injected with virus compared with noninjected contralateral ears and non-injected littermates, respectively:
Ad5.CMV.GFP (97.5dB ± 5, n=4; 45dB ± 0, n=4; 45.8dB ± 7.4, n=6),
Adf.11D (99.4dB ± 19.8, n=9; 56.1dB ± 15.4, n=9; 51.3dB ± 4.3, n=20),
BAAV.CMV.GFP (43.5dB ± 7.5, n=4; 46.3dB ± 4.8, n=4; 44dB ± 5.7, n=12).
Using the two-sample t-test, both early- and advanced-generation AV injected
ears had significant ototoxicity (indicated by asterisks) compared with
contralateral non-injected ears (Ad5.CMV.GFP: t(6)=21.0, p=7.6x10^-7;
Adf.11D: t(15)=5.19, p=1.0x10^-4), while BAAV injections were nonototoxic (p>0.05).
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Figure 32. BAAV transduction of cochlear cells in the adult mouse inner ear.
BAAV.CMV.GFP transduction of supporting cells in the organ of Corti (a-f)
and spiral ganglion neurons (g) is shown by GFP fluorescence in 5-week old
mice following otocyst injection at E12.5. Occasional inner (c) and outer hair
cell transduction occurs. Immunostaining with anti-myosin VIIA antibody
marks the inner and outer hair cells (b, e). Merged images are shown in (c, f).
Nuclei stained with DAPI. IHC = inner hair cell; OHC = outer hair cells; SGN
= Spiral ganglion neurons
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Figure 33. BAAV transduction in the cochlear apex and base. BAAV.CMV.GFP
transduction of cochlear sensory epithelium is shown by GFP fluorescence in
P5 explants of cochlear sensory epithelium. Immunostaining with anti-myosin
VIIA antibody marks the sensory hair cells.
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Figure 34. Gradient of BAAV transduction from cochlear apex to base. (a) Graphical
representation of the number of transduced cells in 500 µm segments
extending from the apex to the base of BAAV11-2, a representative explant
from a BAAV-injected cochlea. (b) Mean transduction efficiencies from the
most apical, middle, and most basal 1mm segments of cochleae were 39.8 ±
5.3, 29.5 ± 13.1, and 8.3 ± 5.6 transduced cells/500µm, respectively.
Transduction efficiencies at the apex and at the middle of the cochlea were
each significantly higher than the efficiency at the base according to the twosample t-test for equal variances (Apex: t(6)=8.24, p=1.7x10-4 (**); Middle:
t(6)=2.99, p=0.02 (*)). The transduction efficiencies between the apex and the
middle did not significantly differ (t(6) =1.45, p > 0.05).
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CHAPTER 6: IN VIVO DELIVERY OF BAAV.MIRNA TO GJB2R75W MICE
Introduction
The previous chapters discussed the HsGJB2-R75W transgenic mouse model for
dominant hearing loss, the optimization of RNAi-based therapeutic, the microinjection
technique for delivery to the developing cochlea, and the identification of BAAV as a
safe vector for gene transfer to the supporting cells of the organ of Corti. The selection of
each of these has been directed by our knowledge regarding the molecular mechanism,
pathophysiology, and expression pattern of GJB2-related hearing loss. The next step is to
combine these tools for in vivo application of RNAi-based therapy for deafness.
In a previous study performed by our lab, we verified that siRNA targeting the
HsGJB2-R75W-EGFP construct was sufficient to prevent hearing loss associated with
exogenous delivery of the mutant construct.[102] This study offered proof-of-principle
that RNAi can be effectively used in vivo to protect against hearing loss caused by the
HsGJB2-R75W allele. I seek to build on this work by using RNAi to suppress the
endogenously expressed HsGJB2-R75W transgene in the GJB2-R75W mouse.
GJB2 has been extensively studied since mutations in it were first linked to
deafness in 1997 and much is known about the molecular basis of GJB2-related hearing
loss.[28] Several mouse models of GJB2-related hearing loss have been developed,
providing greater understanding of the role of CX26 in the cochlea and the
pathophysiology of CX26 dysfunction.
GJB2 (CX26) is expressed in the supporting cells of the organ of Corti and
fibrocytes of the lateral wall (see Figure 16). Along with CX30, CX26 establishes two
gap junction networks involved in maintaining homeostasis within the cochlea: epithelial
network (organ of Corti) and the connective tissue network (lateral wall). The epithelial
network is required to maintain the inner and outer hair cells and the connective tissue
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network plays a role in generating the endocochlear potential.[47] The Cx26OtogCre mice
have a targeted ablation of Cx26 in the epithelial gap junction network only. The hearing
loss in these mice indicates that Cx26 expression in the organ of Corti is necessary for
normal hearing.[54] Since a connective tissue network-specific ablation of Cx26 has not
been reported, it is not known whether Cx26 expression in the epithelial network alone is
sufficient for normal hearing. At a minimum, however, connexin function must be
restored to the organ of Corti to prevent hearing loss due to GJB2 mutations.
Differing from targeted ablation of Gjb2 in mice or patients homozygous for lossof-function GJB2 mutations, dominant mutations in GJB2 disrupt the gap junction
networks by exerting a dominant-negative effect on wild-type connexins. Different
dominant GJB2 mutations have slightly different mechanisms for their dominantnegative effect. The R75W mutation leads to a slight reduction of CX26 in the
membrane, impaired connexon-connexon docking, and altered channel permeability.
Knowledge regarding GJB2 and its expression and function in the cochlea have
allowed us to design therapeutics with an increased likelihood of success. Because
haploinsufficiency of GJB2 does not cause hearing loss in humans and mouse mutants
heterozygous for a targeted deletion of Gjb2 (Gjb2 +/−) do not develop hearing loss, it is
clear that silencing the mutant allele with RNAi would be sufficient to prevent hearing
loss associated with dominant-negative GJB2 mutations.[102] In summary, RNAi-based
therapeutics are particularly applicable to dominant hearing loss due to mutations in
GJB2 for the following reasons: the molecular basis and cell type have been identified;
the disease phenotype is caused by a dominant-negative mutation, rather than
haploinsufficiency, which allows for a therapeutic effect through silencing of the mutant
CX26 alone; the inner ear is surgically accessible in human patients and, via the otocyst
microinjection, in mice.
The otocyst microinjection technique allows us to target therapeutic to the
developing murine cochlea at embryonic day 12.5 (E12.5), approximately 3 days prior to
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onset of Cx26 expression and formation of the epithelial gap junction network.[55, 195]
BAAV’s strong tropism for supporting cells in the organ of Corti enables safe and
directed delivery of the therapeutic RNAi to the source of the pathology. We
hypothesized that the pAAV_tVal-miGJB2-D construct packaged in a BAAV vector and
delivered to the embryonic otocyst can suppress expression of the endogenous HsGJB2R75W transgene and prevent hearing loss in the transgenic mice. In this chapter we
discuss packaging the RNAi therapeutic in the BAAV vector, testing for knockdown in
vitro, and delivery to the transgenic mice.
Materials and Methods
BAAV Vector Production
Endotoxin-free pAAV_tVal-miGJB2-D and pAAV_tval-miNC1 plasmids were
restriction digested to ensure integrity of plasmid, and sent to NIH for packaging into
BAAV. The vector was produced as previously described, purified using CsCl gradients
and particle titers were determined by QPCR [166, 173]. After packaging, viral vectors
were tested in cell culture to confirm expression of DsRed and dialyzed in DMEM (for in
vitro assays) or PBS (for in vivo use). The recombinant BAAV particle titers were 1.0 x
1011 Dnase-resistant particles per milliliter (DRP/mL) and 0.5 x 1011 DRP/mL for
BAAV.miGJB2-D and BAAV.miNC1, respectively.
Cell Culture and Transfection
For knockdown assays, CHO cells were seeded at a concentration of 1.0 × 105
cells/well in 6-well plate in 2mL of media per well. pHsGJB2-R75W-myc-FLAG
plasmid transfections (1µg/well) were performed using FuGene6 (Roche) 24 hours after
seeding. Immediately after plasmid transfection, 2 µL of BAAV (1011 DRP/mL) was
delivered to each well (final volume = 1050 µL). After 24 hours media was changed and
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another 5 µL of BAAV was added to a total volume of 1mL per well. 48 hours posttransfection, the cells were lysed and protein extracted for use in Western blot.
Western Blot
Cells were lysed on ice with 400 µL of radioimmunoprecipitation assay (RIPA)
buffer (50 mM Tris pH 7.4, 150 mM NaCl, 1% NP-40, 0.5% deoxycholic acid, 0.1%
SDS) (Thermo Scientific, Rockford, IL) supplemented with Complete Mini protease
inhibitor cocktail (Roche Diagnostics, Indianapolis, IN). Lysates were cleared by
centrifugation and total protein concentrations were determined by DC protein assay
(Bio-Rad, Hercules, CA). Proteins were diluted in PBS to equivalent concentrations and
stored at -80°C.
Whole cell lysates were mixed with standard (2x) protein loading buffer and
heated for 5 minutes at 95°C . Proteins were then separated by SDS-PAGE in a 15-well
4-20% Tris-HCl gradient gel (Bio-Rad, Hercules, CA) at 150 volts for 90 minutes and
then transferred to nitrocellulose membranes for western blotting. Non-specific binding
to membranes was blocked with Odyssey Blocking Buffer (LI-COR Biosciences,
Lincoln, NE). Membranes were incubated with primary antibodies in Blocking Buffer
overnight at 4°C. Primary antibodies were an anti-FLAG antibody raised in mouse and an
anti-β-actin antibody raised in rabbit (Val Sheffield Lab, University of Iowa) at 1:1,000
and 1:10,000 dilutions, respectively. Membranes were incubated with NIR-dye labeled
secondary antibodies in Odyssey blocking buffer for 1 hour at room temperature.
Secondary antibodies were 680-channel anti-rabbit (1:1,000) and 800-channel anti-mouse
(1:10,000) (Michael Henry Lab, University of Iowa). Antibodies were detected using the
Odyssey infrared imaging system (LI-COR Biosciences) with assistance from Michael
Miller (Michael Henry Lab, University of Iowa). Band densities were averaged from
duplicate experiments (two separate transfection wells run on separate lanes) and
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normalized to internal β-actin controls using Odyssey Application Software (LI-COR
Biosciences).
Otocyst Injections and Auditory Brainstem Response
Testing
Injection of vectors into the left otocyst of E12.5 mouse embryos was performed
via transuterine microinjection as described previously.[138, 149] BAAV.miGJB2-D,
BAAV.miNC1 and PBS were injected. BAAV vectors had been dialyzed in PBS and
were at a final concentration of 1.0 x 1011 DRP/mL and 0.5 x 1011 DRP/mL for
BAAV.miGJB2-D and BAAV.miNC1, respectively. Click-stimulus and pure-tone
auditory brainstem response testing was performed at age P21, as described in the
previous chapter.
Results
Packaging of Artificial miRNA into BAAV
After selecting the plasmid expressing artificial miRNA GJB2-D (miGJB2-D)
with the tRNV-valine promoter (tVal) as the therapeutic, we packaged the construct into
a BAAV vector. The packaging was performed at the National Institutes of Health
(Bethesda, MD) by our collaborators, Dr. John Chiorini and Dr. Giovanni DiPasquale.
The therapeutic vector expresses tVal-driven miGJB2 and CMV-driven DsRed
fluorescent protein and is named BAAV.tVal-miGJB2-D.CMV-DsRed (BAAV.miGJB2D) (Figure 35). A scrambled-sequence artificial miRNA was also packaged into BAAV
to serve as a control vector: BAAV.tVal-miNC1.CMV-DsRed (BAAV.miNC1). The
DsRed fluorescent protein will allow for visual verification of transduction. DsRed was
selected because the GJB2-R75W transgenic mouse already expresses GFP
endogenously. This strategy allows visualization of both the transgene-associated GFP
and the BAAV-associated DsRed. After packaging, viral vectors were tested in cell
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culture to confirm expression of DsRed and dialyzed in DMEM (for in vitro assays) or
PBS (for in vivo use). The final virus titers were 1.0 x 1011 DRP/mL and 0.5 x 1011
DRP/mL for BAAV.miGJB2-D and BAAV.miNC1, respectively.
In vitro Assay of Knockdown with BAAV
Prior to delivering the BAAV vectors to the transgenic mice we performed an in
vitro transduction assay. As we had done previously, we performed a silencing assay in
CHO cells. Rather than quantify silencing by measuring mRNA levels with QRT-PCR,
silencing with BAAV.miGJB2-D was quantified at the protein level. RNAi-mediated
silencing occurs at the mRNA level, but protein levels are ultimately responsible for the
deafness phenotype in the transgenic mouse. Therefore, for in vitro cell culture assay
prior to in vivo application, GJB2-R75W protein knockdown was measured with Western
blot.
CHO cells were plated in a 6-well plate, transfected with pGJB2-R75W_mycFLAG and then, in duplicate wells, treated with either BAAV.miGJB2-D,
BAAV.miNC1, or DMEM-alone. 24 hours following transfection and first treatment with
virus cells were visualized to confirm DsRed fluorescence. DsRed was strongly
expressed in a majority of the cells (50-75%) in each of the BAAV-treated wells.
Subsequently, media was changed and a second treatment with virus or control was
administered. After a total of 48 hours from time of initial transfection and treatment,
cells were washed with PBS and lysed in RIPA buffer.
Prior to Western blotting, concentrations of whole cell protein extracts were
determined using the DC protein assay (Bio-Rad) and samples diluted in PBS to
equivalent concentrations (0.7 mg/mL). FLAG-tagged GJB2-R75W protein was probed
for with anti-FLAG antibodies raised in mouse and, as a loading control, β-actin protein
was probed for using anti-β-actin antibodies raised in rabbit. Anti-mouse (800-channel,
1:10000 dilution) and anti-rabbit (680-channel, 1:1000 dilution) secondary antibodies
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labeled with near-infrared (NIR) spectrum fluorescent dyes were used for detection and
membranes were scanned using the Odyssey imaging system. Integrated intensities for
each band were determined using Odyssey application software, normalized to levels of
β-actin control bands and averages from the duplicate lanes determined. BAAV-treated
protein levels were compared to negative control (DMEM-only treatment) protein levels
(set as 100%). Cells treated with BAAV.miGJB2-D showed expression levels of 70.9% ±
1.9% (29.1% knockdown) and cells treated with BAAV.miNC1 showed levels of 67.6%
± 14.7% (22.4% knockdown) compared with controls (Figure 36). There was no
statistically significant difference in protein levels between the BAAV-treated cells and,
based on the two sample t-test for unequal variances, there was no statistically significant
knockdown achieved by either BAAV.miGJB2-D or BAAV.miNC1 compared with the
control (p=0.96 and p=0.42, respectively). We further investigated these negative results
and discovered that there had been a sequence error in the backbone for my artificial
miRNAs. This small error (absence of 2 nucleotides in the loop structure) would, after
processing by Drosha and Dicer, result in an siRNA that is shifted by a single base.
In vivo Delivery
With the understanding that my “therapeutic” BAAV.miGJB-2 would not be
therapeutic after all, I decided that it would still be worthwhile to deliver it to mice. Both
the BAAV.GJB2-D and BAAV.NC1 would serve as controls and, if delivered without
detrimental effects, would indicate that delivering constructs such as these will not cause
harm to the developing mouse cochlea. We moved forward with in vivo delivery of the
construct to GJB2-R75W transgenic mice via injection to the embryonic otocyst. The
GJB2-R75W transgenic mice are maintained in a hemizygous state on a mixed C57BL/6
and CBA background. Due to their breeding efficiency, large litter size, and postnatal
care of pups, BALB/c mice are the best breed for the in utero surgery.[149] I have found
that CBA mice are much less reliable for timed breeding purposes, have smaller litter
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sizes, and are more likely to cannibalize their pups. Based on these observations, our
timed breeding strategy for GJB2-R75W transgenic mice was for set up male transgenic
mice to breed with female BALB/c mice. We set up approximately 60 female BALB/c
mice for timed breeding with GJB2-R75W transgenic male mice (maximum female to
male ratio per cage was 5:1) over the course of 3 nights. Each morning the males and
females were separated, females with vaginal plugs marked as “possibly pregnant” and
non-plugged females used for the next round of timed breeding. At the end of the three
consecutive nights of timed breeding we had approximately 35 “possibly pregnant” mice
in which the day of conception was known.
Three days prior to embryonic day 12.5 (E12.5), we weighed and recorded the
body weight for each “possibly pregnant” mouse. On the morning of E12.5 we reweighed each mouse and labeled those that had gained at least 1 gram as “pregnant.” Of
the approximately 35 “possibly pregnant” mice, we had 22 “pregnant” mice. There were
approximately 5-10 additional mice which did not have vaginal plugs, but appeared to be
pregnant.
Over the course of two days, with the help of Dr. Samuel Gubbels (Division of
Otolaryngology, University of Wisconsin), we performed in utero surgery on a total of 17
pregnant mice. There were a total of 110 viable (beating heart) embryos, and we
attempted otocyst injections on 96 of them (87.3%). 88 out of the 96 injections (91.7%)
appeared to fill the otocyst, producing a total of 80.0% of all viable embryos injected. 60
total embryos were injected with BAAV.miGJB2-D, 14 were injected with
BAAV.miNC1, and 4 were injected with PBS-only control. Only left otocysts were
injected, leaving the right side as a non-injected control. Two of the 17 pregnant mice did
not survive the 24-hour post-operative recovery period. Of the 15 surviving pregnant
mice, a majority gave birth, but remnants of the pups in many of the cages indicated
cannibalization. Unfortunately, only four mice gave birth to a total of 12 pups that
survived the first 48-hour postnatal period. Therefore, only 12 out of 110 viable embryos
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(10.9%) survived to maturity for further analysis. Of the 12 surviving pups, 7 had been
injected with BAAV.miGJB2-D, 1 with BAAV.miNC1, and 4 with PBS.
Hearing Results
The surviving mouse pups remained in the cage with their mothers until age P21.
At 3 weeks of age, the pups underwent auditory brainstem response (ABR) testing to
assess hearing function in each ear. We performed both click stimulus and pure-tone
ABRs. Pure-tone ABRs were measured at 8, 12, 16, and 24 kHz, frequencies which
correspond to the low to middle range of mouse hearing.[56] The mice were not
genotyped prior to ABR testing and so the operator was blinded to the genotype of the
mice as well as to the substance injected. Following ABR testing the pups were weaned
and genotyped.
The high accuracy with which we were able to perform the otocyst injections
resulted in a number of pregnant mice in which 100% of the embryos had been injected.
This injection data verified that 11 out of the 12 surviving pups had received a good
injection in the left otocyst. This information, combined with the genotypes, allowed us
to group the mice for analysis. In total, there were 7 mice (2 transgenic, 5 wild-type)
definitely injected with BAAV.miGJB2-D, 4 mice (3 transgenic, 1 wild-type) definitely
injected with PBS, and 1 wild-type mouse possibly (50% chance) injected with
BAAV.miNC1. This mouse is awaiting immunohistochemistry for confirmation of
injection. The ABR thresholds (dB SPL for 8 kHz, 12 kHz, 16 kHz, 24 kHz, click
stimulus; n=number of mice tested) were measured for each group of mice and reported
in Figure 37. There are no statistically significant differences between any of the
treatment groups.
Discussion
In this chapter we combined all of our preliminary work for in vivo application:
therapeutic RNAi (miGJB2-D) packaged in the delivery vector (BAAV) injected to the
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embryonic otocyst of the GJB2-R75W transgenic mouse model. As expected, given the
sequence error in the artificial miRNA construct, we did not see a therapeutic result. We
first noticed this error after our in vitro assays failed to yield significant knockdown. As
we further reviewed the artificial miRNA construct to try and understand why it did not
silence the transgene, we came across the design error in the artificial miRNA backbone
upon which our constructs were based. We noticed that the loop sequence that we had
used for the miRNA hairpin was missing two nucleotides. After the hairpin is processed
by Drosha and Dicer, this small error would result in a one-base shift of the siRNA that is
loaded into RISC. This slight difference could potentially explain the lack of silencing by
the artificial miRNA. A sequence shift by even a single base can lead to a 10-fold or
greater difference in silencing capacity.[104, 112, 196] This could be the difference
between a highly functional siRNA and a non-functional siRNA.
Dicer processing is not perfectly predictable. D-siRNAs are designed to be
predictably cleaved by Dicer, but even they are subject to alternative processing, resulting
in a majority of 21-base siRNA and a minor fraction of 22-base siRNA.[105, 130] So it is
possible that the single base shift would still lead to production of at least a portion of the
desired siRNA, but certainly not at optimal levels. Similarly, there are regions of mRNA
that are readily targeted by multiple siRNAs across the region and so it is possible that a
shift by a single nucleotide would not significantly alter the results. However, our lack of
in vitro transgene silencing points to a non- or reduced-functioning siRNA which is best
explained by our design error in the artificial miRNA backbone. We are currently redesigning the artificial miRNAs with the correct backbone and will test in vitro prior to
further in vivo application.
I decided to proceed with the in vivo delivery because I realized that there was
still information that could be obtained from in vivo administration of the artificial
miRNA constructs, including: (i) whether DsRed expression driven by CMV in the
developing cochlea would cause toxicity; (ii) whether artificial miRNA expression driven
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by tVal would cause toxicity to the developing cochlea; (iii) confirmation of our previous
studies indicating that BAAV transduction of the developing cochlea is safe.
Our post-injection hearing results indicated no differences in hearing between the
ears injected with the BAAV constructs and non-injected ears or ears injected with PBS.
These results confirm that BAAV is safe for delivery to the embryonic otocyst and
indicate that neither the DsRed protein nor artificial miRNA is toxic to the developing
cochlea. The BAAV.miGJB2-D construct that produces a single-base shift in the
processed siRNA is a particularly good control. Scrambled-sequence siRNA is a
commonly used control, but some sources suggest using a siRNA with high sequence
similarity to the therapeutic siRNA but with changes to a few of the bases. This is
recommended as the optimal control because places a similar molecule in the cells but
has enough difference that it should not induce silencing via the RNAi mechanism.[111]
The lack of detrimental effects in these controls offers encouragement that all aspects of
our delivery system are safe and, given an effective therapeutic, only has the potential to
help rather than to harm.
Otocyst Injections
The transuterine embryonic otocyst injection technique is a powerful method for
delivering genes to the progenitor cells of the cochlea. It provides a means for therapeutic
intervention prior to onset of pathology, even in the case of congenital hearing loss.
While this technique has great power and potential, it comes with several drawbacks that
were especially evident with the injections described in this chapter. The greatest
drawback with the otocyst injection technique is the often low numbers of injected
animals. The procedure is highly technical with extreme variability in efficiency
depending on the surgeon. We have implemented strategies to improve outcomes, but
chance seems to still play a more significant role than we would like. From the first time
timed breeding step all the way through until several days after birth, the number of mice
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can be significantly reduced at each stage (Figure 38). In the past, our rate limiting step
has often been lack of pregnant mice and poor injection efficiency. This time we set up
dozens of breeders to ensure sufficient numbers of pregnant mice and Dr. Gubbels
performed injections with incredible precision, but mice were lost in subsequent steps.
We always expect that a fraction of each litter will resorb and never be born. This
is a natural consequence of manipulating the uterus and the embryos, but the effect is
usually minimized by using extra-fine micropipettes and keeping the duration of surgery
to a minimum. This time was had significantly higher numbers of resorptions than before.
The high number of resorptions was likely due to the high percentage of injections that
were attempted. Usually we only attempt to inject 50-75% of the embryos; this time we
attempted injections on almost 9 out of 10 embryos, injecting 100% of the embryos in
many of the pregnant mice. Although the injections were atraumatic and the surgery time
was kept to a minimum, it is likely that the high percentage of embryos that were injected
led to increased rates of resorption.
Cannibalism is always a possibility for loss pups. To minimize the risk, we house
the pregnant mouse alone, we use BALB/c mice which generally do quite well at
nurturing their pups, and we try not to disturb the cages in the hours during and
immediately following birth. In the most recent injection experiments many of the pups
which were born did not survive the postnatal period; however, it is not clear how many
were cannibalized because the carcasses are not always found. From the number of
carcasses that we did find in the cages it was evident that more of the BALB/c mothers
had cannibalized their pups than we have observed in the past. With other strains of mice,
particularly CBA, we have observed frequent cannibalization of the offspring. It is not
clear why there was increased cannibalization with the BALB/c mice this time. One
potential reason is that, in order to increase the number of mice for breeding, we used
many females that had never been pregnant before. First litters are more likely to be
cannibalized than subsequent litters. In the future, we recommend using mice that have
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already given birth to at least one litter. Fostering is another option which we have not
done because we did not previously observe a need for it. It is an option that we will
consider using in the future to increase survival of the pups.
Hearing Tests
The auditory brainstem response testing performed on the injected mice was
designed to maximize the likelihood of detecting a therapeutic change. The earliest age at
which ABR testing can be performed in mice is approximately P14, just after hearing
onset and opening of the external auditory canal. We predict that the earlier hearing
function is assessed in GJB2-R75W transgenic mice, the higher the likelihood of seeing a
therapeutic effect. However, we have occasionally observed inconsistent results if the
hearing tests are performed too early. We chose, therefore, to test hearing at age P21, the
earliest age at which we are able to consistently measure hearing in mice. In addition to
click stimulus, which is a good general screening test for auditory function over a range
of frequencies, we performed pure-tone ABR testing at 8, 12, 16, and 24 kHz. These
frequencies represent the low to mid frequencies within the hearing range of mice.[56]
Low frequencies are detected by the apex of the cochlea. Based on our previous
observation that BAAV preferentially transduces the apical regions of the mouse cochlea,
we predict that a therapeutic effect is most likely to be measured in the low frequencies.
We did not observe any effect on the hearing of injected mice but, as explained above,
this data serves as a valuable control for the safety of our gene therapy delivery technique
and vehicle.
137
Figure 35. BAAV.miGJB2-D expression construct. tRNA-valine promoter-driven
artificial miRNAs (miGJB2-D and miNC1) were cloned into a rAAV-inverted
terminal repeat (iTR)-containing plasmid upstream of a CMV-driven, dsRED
expression cassette. BAAV.miGJB2-D and BAAV.miNC1 were produced.
138
Figure 36. In vitro silencing of GJB2-R75W transgene with BAAV.miGJB2-D. CHO
cells expressing the GJB2-R75W transgene were treated with
BAAV.miGJB2-D to assess for silencing at the protein level. Protein
expression was assayed by Western blot and quantified using the Odyssey
System. Cells treated with BAAV.miGJB2-D showed expression levels of
70.9% ± 1.9% (29.1% knockdown) and cells treated with BAAV.miNC1
showed levels of 67.6% ± 14.7% (22.4% knockdown) compared with
controls.
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Figure 37. Auditory brainstem response results for mice injected with BAAV constructs.
Pure tone (top) and click stimulus (bottom) hearing thresholds are shown for
wild-type mice (left side) and GJB2-R75W transgenic mice (right side). Mice
were injected with either BAAV.miGJB2-D (“therapeutic), BAAV.miNC1
(“scrambled”), PBS, or non-injected. The ABR thresholds (dB SPL for 8 kHz,
12 kHz, 16 kHz, 24 kHz, click stimulus; n=number of mice tested) were
measured for each group of mice and the hearing results for each group are as
follows: Transgenic + BAAV.miGJB2-D (122.5 ± 3.5, 101.3 ± 1.8, 105 ± 0.0,
81.3 ± 1.8, 77.5 ± 10.6; n=2), Transgenic + PBS (128 ± 0.0, 107.5 ± 2.5, 110.0
± 0.0, 80.8 ± 1.4, 82.5 ± 2.5; n=3), Transgenic + No Injection (126.9 ± 2.5,
104.5 ± 3.3, 112.0 ± 4.1, 81.5 ± 3.4, 79.0 ± 6.3; n=5), Wild-type +
BAAV.miGJB2-D (97.0 ± 2.1, 80.5 ± 2.7, 79.0 ± 1.4, 74.5 ± 5.7, 46.5 ± 3.8;
n=5), Wild-type + PBS (95, 80, 77.5, 77.5, 42.5; n=1), Wild-type +
BAAV.miNC1 (95, 82.5, 80, 75, 52.5; n=1), Wild-type + No injection (97.5 ±
4.1, 77.9 ± 4.4, 78.2 ± 1.9, 61.4 ± 16.9, 47.5 ± 4.1; n=7). There are no
statistically significant differences between any of the treatment groups.
140
Figure 38. Progressively diminishing returns in the otocyst injection procedure. Mice are
inevitably lost at each step of the experimental procedure. This necessitates
beginning experiments with large numbers of mice.
141
CHAPTER 7: CONCLUSIONS AND LESSONS LEARNED
Summary
It is important to identify new approaches to prevent or cure disease. In recent
years, the field of genetics has continued to expand rapidly. Today, geneticists have tools
at their disposal which would have been unimaginable just decades ago. These include
transgenic, knockout, and knock-in animal models;[197, 198] embryonic, adult and
induced pluripotent stem cells;[199-201] the human genome sequence, and completed or
working genome projects for hundreds of other organisms;[1, 2] RNA interference;[97]
genotyping and expression microarrays, and massively parallel sequencing
technologies.[202] Despite these advancements, the technology and data available to the
researcher will continue to expand exponentially. One of the great challenges for the
future of genetics and medicine will be to determine the best use for these tools and data
and how to translate them into improving longevity and quality of life.
In this Thesis, I have applied genetic tools to the treatment of hereditary hearing
loss. Hearing loss, the most common sensory deficit, is commonly treated using hearing
aids or cochlear implantation. While major improvements have been made to these
modalities over the years, they cannot substitute for a functional cochlea. It is estimated
that mutations in several hundred genes may potentially lead to hearing impairment.[13]
Through the study of human pedigrees and mutant mice over sixty deafness genes have
already been identified.[10] Because murine inner ear anatomy and physiology are
similar to human, and in many cases the human deafness phenotype is faithfully
recapitulated in the mouse, mouse models have provided tremendous insight into the
molecular physiology of hearing and pathophysiology of deafness.[13]
My Thesis work has focused on the GJB2-R75W transgenic mouse model of
dominant-negative deafness. GJB2 is the most commonly mutated gene leading to
hereditary hearing loss.[9] The ability to prevent or reduce the severity of GJB2-related
142
deafness would make an enormous impact. Effective gene therapy requires a therapeutic
substrate, a delivery route, and a delivery vector. With the goal of applying gene therapy
to the GJB2-R75W transgenic mouse, I completed the following Specific Aims:
Specific Aim 1: To further characterize the phenotypes of the GJB2-R75W
transgenic mouse model.
Specific Aim 2: To design and validate siRNAs for potent and specific
knockdown of the GJB2-R75W transgene.
Specific Aim 3: To identify and test in vitro a viral vector suitable for safe and
efficient delivery of therapeutics to the supporting cells of the developing murine
cochlea.
Specific Aim 4: To deliver and assess efficacy of in vivo gene therapy for
dominant-negative GJB2-R75W deafness.
First, I further characterized the mouse model. Importantly, I used QRT-PCR to
quantitate cochlear expression levels of the mutant GJB2 compared with levels of the
endogenous wild-type Gjb2. Although the mutant was on average expressed at a slightly
higher level (~1.5:1 transgene to wild-type ratio), I was encouraged that the cochlea was
not so dominated by transgene as to make therapeutic efforts futile. Second, I selected
RNAi as the therapeutic of choice for this dominant model of deafness. The discovery of
RNAi has provided a new mechanism for intervention in diseases caused by a dominantnegative effect.[92] Previous proof-of-principle studies in our laboratory showed that
siRNA could suppress a GJB2-R75W construct exogenously delivered to the adult mouse
cochlea and prevent hearing loss.[102] I built on this work by identifying multiple potent
siRNAs and designing artificial miRNA hairpins for in vivo delivery to target the
endogenous GJB2-R75W human transgene expressed in mice. Third, I studied the inner
ear tropism and safety profile of multiple viral vectors (AAV, early-generation AV, lategeneration AV, and BAAV) delivered to the developing mouse cochlea via transuterine
microinjection to the embryonic otocyst. BAAV was identified as a vector for safe
143
delivery of therapeutic to the supporting cells of the organ of Corti, the major site of
pathology in GJB2-related hearing loss. Finally, we delivered artificial miRNAexpressing BAAV to the embryonic otocysts of both wild-type and GJB2-R75W mice.
During the course of this study I became proficient at this delivery approach. However,
due to a single base error in the artificial miRNA construction we did not observe a
therapeutic effect. Importantly, we did demonstrate that our system of RNAi-delivery to
the mouse cochlea is safe.
Lessons Learned
Throughout the course of this study I have learned numerous lessons that will be
helpful to future efforts directed at gene therapy for deafness and other genetic diseases.
RNA Interference
There are dozens of algorithms available for designing functional siRNAs. I tested
both Rational Design rules and a computer-generated algorithm. The siRNAs designed
with the more traditional Rational Design rules proved the most potent. However, it is
important to concede that no currently available siRNA design strategy is always superior
to the rest. I recommend using at least two different design strategies and testing the top
candidates from each. For future siRNA design, there are two approaches in the literature
which have caught my interest because they incorporate basic Rational Design rules and
new bioinformatics tools.[104, 117]
The first approach, described by Birmingham and colleagues,[104] has the user
identify a target mRNA sequence and use Microsoft Excel to manually generate all
sliding-window 19-base sequences within the target. This list of all candidate siRNAs is
then manually filtered by removing candidates that fail to qualify under established
Rational Design rules such as GC content and positional base preferences (Microsoft
Excel worksheets and online calculators may be used to speed up this filtering process).
Each remaining candidate is prioritized according to its functionality score derived from
144
online scoring algorithms. The highest scoring candidates are analyzed in silico for their
potential to target unintended transcripts. Finally, at least 4 of the top scoring candidates
with the least potential for off-target effects are selected for in vitro testing.
The second approach is the i-Score Designer.[117] The inhibitory-score (i-Score)
is an siRNA prediction algorithm that is based exclusively on base preferences at each
position in an siRNA. The i-Score Designer is a Microsoft Excel VBA program which
calculates 11 different siRNA functionality scores for all possible siRNA sequences
within a user-defined sequence. It also determines the ΔG values, GC content and length
of GC stretches for each siRNA. The program ranks the highest scoring candidates in
each of the scoring algorithms and the authors suggest that the best candidate siRNAs for
in vitro testing are those which receive high scores in most of the algorithms. Either of
these approaches should produce good candidates for in vitro verification.
My experience shows that experimentally validating more than just a few
candidate siRNAs may be more important than the selection method itself. There are
many examples of non-functional siRNAs that were predicted in silico to be highly
functional, and vice versa. Therefore, in vitro verification of siRNAs is essential. Many
design strategies recommend selecting three to five siRNAs for verification; I would
recommend testing as many as can reasonably be tested. In most cases eight to twelve
siRNAs can easily be tested in duplicate or triplicate on a 24-well plate without
significant work or cost.
I selected QRT-PCR as the optimal method for measuring knockdown based on
its high sensitivity and specificity as well as the rapid turnover rate of CX26 protein (~3
hours). For more stable proteins silencing should be confirmed at the protein level. Even
for short-lived proteins like CX26, measurement of silencing at the protein level is
important for therapeutic approaches. Assays with greater sensitivity and fewer
processing steps than immunoblotting are appealing for future knockdown assays, such as
dual luciferase assays and in-cell Westerns.
145
After selection of a potent siRNA, the sequence needs to be re-designed to
accommodate a hairpin backbone that can be expressed in vivo by a viral vector. There
are numerous options for hairpin design including traditional shRNAs and artificial
miRNAs. My decision to use an artificial miRNA based on the miR-30 backbone for in
vivo RNAi delivery was an attempt to maximize gene silencing while minimizing toxic
side effects. Although I feel confident that this is a sound strategy, it is not the only
strategy.[136, 203-206] Silva and colleagues also used the miR-30-based backbone but
found that including additional sequence flanking the miRNA improved the processing
efficiency and spacer sequence between the promoter and the miRNA increased
knockdown efficiency.[135, 136] The artificial miRNA expression plasmids designed by
Silva and colleagues allow for new artificial miRNAs to be inserted easily with several
different promoter options.[135] These plasmids are now available from Open
Biosystems.[136]
Li and colleagues found that shRNAs are as efficacious as artificial miRNAs
provided that they are designed correctly. According to their findings a 19-nt stem and 9nt loop hairpin outperformed a standard 4-nt loop shRNA or miR-30-based artificial
miRNA.[203] This provides a further option for hairpin design. A third design involves a
cassette that allows expression of multiple shRNAs by a single vector. Shan and
colleagues showed that a triple miR-155-based artificial miRNA achieved excellent gene
silencing, and Wang and colleagues designed cassettes which can express up to six
shRNAs.[205, 206] Any or all of these strategies has the potential to improve in vivo
hairpin potency and should be a consideration for future experiments.
Otocyst Injections
I have spent three years of my PhD studies learning and optimizing the
transuterine otocyst microinjection technique.[149] Successful outcomes for our otocyst
microinjections were depended on two variables: the number of otocysts accurately
146
injected and the number of embryos that ended up surviving post-partum (see Figure 38).
Aside from proficiency in performing the injection, my colleagues and I identified
several other ways to improve the outcomes of this technique.
The otocyst injection technique is extremely time-intensive. From beginning to
end surgery for each pregnant mouse requires approximately one hour. This does not
include timed breeding setup, preparation of equipment, cleanup and post-operative
monitoring of the mice. Minimizing the surgery time increases the survival of both the
dam and injected embryos.
One frustration was obtaining sufficient pregnant mice. Even when breeding
significant numbers of mice the ascertainment of pregnant dams is generally low. It is
much better to have too many pregnant mice than too few. I recommend, therefore,
setting up approximately four times as many mice for breeding as you actually plan to
inject. Although not always possible, decreasing the female to male ratio in the timed
breeding cages often increases the likelihood of success. Breeding mice in single pairs
rather single males with multiple females is optimal Another way to increase breeding
success is to setup breeding cages over the course of two or three days and plan to
perform the surgeries over the same period. Females that were not plugged after the first
night will often be plugged after the second. Finally, the strain of mice is very important
for success. Consistent with the studies of Brigande and colleagues,[179] I recommend
the use of BALB/c mice because they breed well and the surgical landmarks are much
more easily visualized in non-pigmented animals. CBA/J mice are poor breeders and the
landmarks can be difficult to see in pigmented C57BL/6 mice. This can become a present
a problem because mice used in hearing research are often placed on the CBA
background due to its decreased susceptibility to age-related hearing loss. Fortunately, I
found that using BALB/c females and CBA background males for timed breeding
provided acceptable results, both in terms of breeding and injection success. BALB/c
mice are the optimal strain that we have used and should be used if at all possible.
147
BALB/c mice are susceptible to age-related hearing loss and so would not be suitable for
studies with timepoints greater than four months. I am currently breeding the GJB2R75W mice onto the BALB/c background for use in future otocyst microinjection
surgeries.
Another step that I’ve implemented to minimize the risk of operating on nonpregnant mice is to weigh the mice several days prior to the day of surgery (E12.5) and
then again on the day of surgery. Only mice that gained more than 1 gram in the 2-3 days
prior to surgery have a high likelihood of being pregnant and are injected. This technique,
in combination with the presence of a post-breeding vaginal plug and visual inspection of
the abdomen size, has made us highly accurate at identifying truly pregnant mice.
The most frustrating challenge in the entire surgical process is loss of injected
embryos due to death of the dam, trauma-induced resorption, or postnatal cannibalization.
It is also the most difficult to prevent. The use of a heating pad under the surgical
platform significantly increased maternal survival. It is evident that efficient surgeries,
both in terms of time and degree of embryonic manipulation, result in increased survival
for both dam and embryos. The importance of not over-manipulating the embryos was
underscored in our most recent surgeries when we attempted injections in approximately
90% of embryos. Although the time of surgery was kept low (between 30 and 40
minutes) and the injections were performed with high accuracy, many of the embryos did
not survive to term. I suspect that the high percentage of embryos that were injected
contributed to the low survival rates (~10%). It was known that embryos adjacent to a
resorbing embryo are also likely to resorb themselves. Striking the balance between
increased number of injections and increased risk of resorption is difficult, but in the
future I recommend attempting injections on only 50-75% of the embryos in any given
litter.
In the most recent series of injections a higher than usual degree of postnatal
cannibalism was observed. This is a particularly challenging problem. There is likely an
148
environmental component to this behavior and anything that stresses the dams during
pregnancy or postnatal period may contribute. In the past, BALB/c mice have performed
very well at nurturing the pups. In contrast to the frequently observed cannibalism with
CBA mice, I have only observed several cases of cannibalism with BALB/c mice in the
past. Other than to check for date of birth I do not disturb the cages during birth or the
early postnatal period. Given the amount of time and effort that goes into each injection, I
think that cross-fostering is an option that should be considered in the future. Crossfostering will require slightly more preparation, but will be worth the extra effort if it
preserves the survival of treated mice.
Scarcity is a universal problem that is especially evident in cochlear research
using the otocyst injection. The mouse cochlea is smaller than a grain of rice with a
maximum of two cochleae per mouse. A maximum of one cochlea per mouse can be
injected. Furthermore, there are only three ways to identify which mice were injected: (i)
inject all mice; (ii) keep track of the injected embryos’ positions within the uterus and
remove them prior to parturition; (iii) the injected substance expresses a fluorescent
protein and its presence is verified visually after sacrifice and removal of the cochlea.
Injecting all mice is not always possible and may lead to decreased survival. We always
record the uterine position of each injected embryo and have used this technique for
verifying transduction prior to birth. However, this method requires sacrifice of the
embryos prior to birth prevents both evaluation of hearing function and of transduction at
later timepoints. The third option, verifying presence of a transgene or fluorescent protein
using immunohistochemistry, is our standard protocol. It allows hearing tests up until the
time of sacrifice and it allows visual verification of transduction by viewing crosssections of the cochlea. However, it is not compatible with a surface preparation of the
sensory epithelium which allows determination of transduction efficiency along the
length of the cochlea. It is also not compatible with extracting DNA, RNA, or protein for
quantification purposes. Thus, after thoughtful consideration is given to the information
149
that is most sought, difficult decisions need to be made regarding the time of animal
sacrifice and the methods of cochlear processing and transduction verification.
Future Studies
The work reported in this Thesis provides a foundation for continued efforts
toward therapy for connexin-related hearing loss. I have identified RNAi that potently
silences the mutant transgene and shown that our delivery system safely and effectively
targets the supporting cells in the murine cochlea. Looking to the future, our top priority
is to treat the GJB2-R75W transgenic mice with an efficacious artificial miRNA
construct. The correct hairpin construct is currently being synthesized and will be
validated in vitro prior to delivery to the transgenic mice. I am optimistic that this
strategy will work, provided we are able to achieve sufficient transduction and expression
of the miRNA in supporting cells in vivo. If strong in vivo knockdown is not attained then
others strategies, such as stronger promoters or multiple-shRNA expression systems, will
need to be considered.
Our system for gene therapy delivery can also be adapted to other models of
connexin-related hearing loss. For example, gene replacement therapy could be applied to
recessive models of connexin-related deafness. BAAV vectors expressing wild-type Gjb2
or Gjb6 could be delivered to the developing cochleae of conditional Cx26-null or Cx30null mice, respectively. Cochlear pathology in Cx30-null mice is reported to have slightly
later onset and to be slightly less widespread than in conditional Cx26-null mice. A
mouse with the dominant T5M mutation in Cx30 was recently reported to have mild
hearing loss.[55] These results indicate a less severe pathology compared with Cx26
mutants and thus therapy directed at Cx30 mutants would be expected to have a higher
likelihood of success.[56]
Another option for future therapeutic trials would be to create a new mouse model
for dominant-negative Cx26 deafness using a mutation that is functionally less severe
150
than the R75W mutation. The D179N mutation in GJB2 causes ADNSHL. The
phenotype of patients with the D179N mutation is less severe than with other dominant
mutations in GJB2. The hearing loss is later onset and is not associated with other
pathologies.[207] Corresponding to the less severe phenotypic manifestations, in vitro
functional studies indicate a less severe dominant-negative effect with the D179N mutant
than with other dominant CX26 mutants, including R75W.[208] Rather than completely
preventing channel function, the effect of D179N was to reduce function to a level
intermediate to that of wild-type and the other dominant CX26 mutations tested. I predict
that the weaker dominant-negative effect of this mutation could be more easily overcome
by therapeutic intervention. I propose that the D179N mouse model be created as a
targeted knock-in resulting in one wild-type and one mutant copy of the gene, expressed
by the endogenous Gjb2 promoter. This would recapitulate the genetic context of the
human GJB2 mutation and increase the likelihood of attaining therapeutic levels of
silencing.
Another potential genetic therapy for dominant-negative CX26 deafness is based
on the observation that the root problem is one of relative connexin quantity. Since
connexin oligomerization is thought to be a stochastic process, the predicted composition
of connexons within a cell is predictable and will reflect the relative abundance of
available of connexins. An individual heterozygous for a mutation in GJB2 will have
approximately 50% wild-type and 50% mutant CX26 proteins available for connexon
assembly and the relative abundance of different connexons is modeled by a bell-shaped
distribution (Figure 39). Small changes in the ratio of available connexins can result in
substantial changes in composition of expressed channels.[209] Our approach has been to
use RNAi to reduce the levels of mutant relative to levels of wild-type, thereby shifting
the curve toward the formation of connexons composed completely or mostly of wildtype connexins. I hypothesize that over-expression of wild-type connexins will result in
151
the same shift of the curve (Figure 39). A sufficiently large shift could result in levels of
functional channels at or above that required for normal hearing.
This hypothesis can be tested using transgenic mice that have already been
created: BAC-Cx26 and BAC-Cx30 mice. Dr. Erick Lin (Emory University, Atlanta,
GA) created these transgenic mice to over-express Cx26 (BAC-Cx26) or Cx30 (BACCx30).[147] The mice express approximately twice the endogenous level of the each
connexin and are phenotypically indistinguishable for non-transgenic mice. These mice
have already been used to therapeutic advantage. Through breeding, the BAC-Cx26
transgene was transferred to Cx30-null mice. The presence of the BAC-Cx26 transgene
resulted in a complete rescue of the hearing of the Cx30-null mice.[147] I hypothesize
that genetic over-expression of Cx26 and/or Cx30 using the BAC-Cx26 and BAC-Cx30
transgenic mice has the potential to overcome the dominant-negative mutation and
restore hearing to the GJB2-R75W transgenic mice. This hypothesis is supported by a
variety of in vitro studies indicating that increased ratios of wild-type connexin, coexpressed with a dominant-negative mutant, can improve channel function.[210, 211] I
have obtained the BAC-Cx26 and BAC-Cx30 transgenic mice and have started breeding
them with the GJB2-R75W transgenic mice. Breeding and assessing the hearing function
is an ongoing project in our laboratory. If the over-expression is not sufficient to prevent
hearing loss, futures experiments could combine over-expression of wild-type with
silencing of mutant to achieve even more favorable ratios. If the dominant-negative effect
is strong, increasing the ratio of wild-type protein may not be sufficient. This was true in
experiments performed in our laboratory on a dominant-negative DFNA2 mouse model
carrying a KCNQ4 mutation. Supplementation of wild-type KCNQ4 in the cochlea of
these mice was not sufficient to overcome the dominant-negative effect of the mutant
KCNQ4 and prevent hearing loss (unpublished data).
Much progress has been made during the past decade in the field of hearing and
deafness research. However, there is much still to be done. Improving technologies will
152
enable more detailed study of the cochlea; genetic tools will lead to the identification of
new genes involved in hearing and deafness and provide more accurate diagnostic and
prognostic information for patients and their families; therapeutic safety and vector
delivery will be optimized for application to human patients. The ability to quickly
identify and deliver potent RNAi molecules to silence pathogenic dominant genes in the
ear would represent a big step towards personalized therapies.[85] As we continue to
press forward with zeal founded upon knowledge, genetic therapies for hearing loss will
emerge as viable treatment options.
153
Figure 39. Predicted connexon channel composition based on stoichiometric ratios. The
bell-shaped curve (dark blue) indicates the predicted % of connexons
(hemichannels) in a cell with the six different potential combination of mutant
and wild-type connexins (shown on the x-axis) given equal ratios of wild-type
to mutant connexins in the cell (e.g. heterozygote). The curve is skewed to the
right (light blue) when the ratio of wild-type connexins increases relative to
mutant connexins. Increasing wild-type to mutant ratios, either through
silencing the mutant or by over-expressing the wild-type can result in
substantial changes in the composition of connexon channels within a cell.
Calculations were performed using the probability mass function of the
binomial distribution.
154
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