Lymphatic Drainage from the Mouse Eye and the Effect of Latanoprost
by
Alex Lai Chi Tam
A thesis submitted in conformity with the requirements for the degree of Master of Science
Department of Laboratory Medicine and Pathobiology
University of Toronto
© Copyright by Alex Lai Chi Tam 2013
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Lymphatic Drainage from the Mouse Eye and the Effect of Latanoprost Alex Lai Chi Tam
Master of Science
Department of Laboratory Medicine & Pathobiology University of Toronto
2013
Abstract
Glaucoma is a leading cause of world blindness, often associated with elevated eye
pressure. Current glaucoma treatments aim to lower eye pressure by improving aqueous humor
outflow from the eye. Ocular lymphatics have been demonstrated to contribute to aqueous humor
outflow in human and sheep. It is not known whether any glaucoma drugs target this lymphatic
drainage. The mouse is a valuable model with similar aqueous humor dynamics and
pharmacology as human. Using in vivo hyperspectral fluorescence imaging combined with
intracameral quantum dot injection, we identified an ocular lymphatic drainage in mouse.
Immunofluorescence and confocal microscopy revealed lymphatic channels in the ciliary body,
sclera, and orbit that may be responsible for this lymphatic drainage. We showed that latanoprost,
a prostaglandin F2α analog widely used to treat glaucoma, increases this ocular lymphatic
drainage. Our findings provide the framework for future development of novel glaucoma drugs
that stimulate the ocular lymphatic drainage.
iii
Acknowledgements
I owe a debt of gratitude to both, Dr. Yeni Yücel and Dr. Neeru Gupta, for their guidance,
vision and support throughout the past years. Thanks for taking a chance on an inexperienced
student and patiently provided him with superior scientific direction and life advice. The
accomplishments I have achieved over these years have only been possible with their unending
patience, understanding, and rigorous training.
Thanks must go to Dr. Bhagu Bhavnani for his time and advice. His expertise has
undoubtedly helped in the completion of this thesis.
Thank you to the lab members of the Eye Research Lab for the moral support, stimulating
discussion and their assistance during various stages of the work.
Special thanks are due to my friends - Scott, for introducing the world of research to me;
Jessica, for believing in me; Adrian, for his motivational words; and Yvonne, for her loving
support of my studies.
My biggest debt of gratitude, however, must be to my parents and Andy, whose constant
support and encouragement has never wavered. Thanks, thanks, and thanks again.
A final word of remembrance is for my grandfather, M.Y. Woo, and grandmother, P.C.
Woo, who inspired me to pursue this degree and who are both much missed.
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TABLE OF CONTENTS
Precontent
Abstract ii
Acknowledgements iii
List of Publications vi
List of Abbreviations vii
List of Figures and Tables ix
Contributions xi
1. Introduction 1
1.1 Overview 1
1.2 Glaucoma and Its Relation to Aqueous Humor Dynamics 2
1.3 Aqueous Humor - Composition, Formation and Dynamics 2
1.3.1 Trabecular Meshwork Outflow (Fig. 1-1) 4
1.3.2 Uveoscleral Outflow (Fig. 1-1) 5
1.3.3 Ocular Lymphatic Drainage (Fig. 1-1) 6
1.3.3.1 Non-Ocular Lymphatics 6
1.3.3.2 Ocular Lymphatics in Relation to Ocular Lymphatic Drainage 7
1.4 Glaucoma Treatment 8
1.5 Prostaglandins and Non-Ocular Lymphatics 10
1.6 Background on Using a Mouse Model 11
1.7 Justification of In Vivo Imaging for Study of Lymphatics 12
1.7.1 Conventional Lymphatic Drainage Imaging 12
1.7.1.1 Cannulation of Lymphatics to Detect Injected Radioactive Tracers 13
1.7.2 Non-Invasive In Vivo Imaging of Lymphatic System with Organic
Fluorophores
13
1.7.3 Non-Invasive In Vivo Imaging of Small Animals with QDs 14
1.8 Rationale and Hypothesis 15
1.8.1 Rationale 15
1.8.2 Hypotheses 16
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2. Material and Methods 19
2.1 Subjects 19
2.2 Quantum Dots 20
2.3 Topical Application of Substances Prior to QD Tracer Injection 20
2.4 Intracameral Injection of QD Tracer 20
2.5 IOP Measurement 21
2.6 In Vivo Hyperspectral Fluorescence Imaging 22
2.7 Control Experiments for QD drainage 24
2.8 Tissue Collection and Processing 24
2.8.1 Head and Neck Specimen 24
2.8.2 Orbit Specimen 25
2.9 Frozen Sections 25
2.9.1 Head and Neck Specimen 25
2.9.2 Orbit Specimen 26
2.10 Quantification of Tracer Drained into Lymph Node 26
2.11 Immunofluorescence and Nuclear Staining 26
2.11.1 Submandibular Lymph Node 26
2.11.2 Right Eye and Orbit Specimen 27
2.12 Bright Field Microscopy 28
2.13 Combination of Fluorescence and Bright Field Microscopy 28
2.14 Confocal Microscopy 29
2.15 Statistical Analysis 29
3. Results 31
4. Discussion 51
4.1 Limitation of the Studies 59
4.2 Future Directions 61
4.3 Conclusion 64
5. References 65
vi
List of Publications
1. A.L.C. Tam, N. Gupta, Z. Zhang, Y.H. Yücel. Quantum Dots Trace Lymphatic Drainage from the Mouse Eye. Nanotechnology 2011 Sep 21; 22(42):425101.
2. A.L.C. Tam, N. Gupta, Z. Zhang, Y.H. Yücel. Latanoprost Stimulates Ocular Lymphatic
Drainage: An In Vivo Nanotracer Study. Translational Vision Science & Technology 2013 Aug 7; 2(5):3, http://tvstjournal.org/doi/full/10.1167/tvst.2.5.3.
3. A.L.C. Tam, N. Gupta, Y.H. Yücel. Identification of Lymphatics in Ciliary Body, Sclera, and Orbit of the Mouse Eye. (Manuscript in preparation).
vii
List of Abbreviations
AqH Aqueous humor
BSA Bovine serum albumin
CAI Carbonic anhydrase inhibitors
DAPI 4',6-diamidino-2-phenylindole
H&E Hematoxylin and eosin
IOP Intraocular pressure
LEC Lymphatic endothelial cell
LYVE Lymphatic vessel hyaluronan receptor
NIR Near infrared
NPE Non-pigmented ciliary epithelium
PBS Phosphate buffered saline
PE Pigmented ciliary epithelium
PG Prostaglandin
PGA Prostaglandin analog
PGF2α Prostaglandin F2α
Prox Prospero-related homeodomain transcription factor
PVA-DABCO Polyvinyl alcohol-1,4-diazabicyclo[2.2.2]octane
QD Quantum dot
RGC Retinal ganglion cell
SC Schlemm's canal
viii
TM Trabecular meshwork
TRITC Tetramethylrhodamine-5-(and-6)-isothiocyanate
TSA Tyramide signal amplification
UVS Uveoscleral
VEGFR Vascular endothelial growth factor receptor
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List of Figures and Tables
Chapter 1
Figure 1-1
Aqueous Humor Outflow Pathways 17
Figure 1-2
Chemical structures of PGF2α and commercially available PGAs 18
Chapter 2
Figure 2-1
Timeline for IOP measurements 22
Figure 2-2
Timeline for in vivo hyperspectral imaging 24
Table 2-1
Exposure times (ms) at various magnifications for frozen sections of orbit specimen
29
Chapter 3
Figure 3-1
In vivo hyperspectral fluorescence imaging of head and neck in dorsal (A-C) and ventral (E-G) views
33
Figure 3-2
Post-mortem analysis of neck tissue 34
Figure 3-3
Immunofluorescence of left submandibular lymph node 35
Figure 3-4
QD distribution in left eye 6h following intracameral injection in control mice
37
Figure 3-5
Podoplanin-positive lymphatic channels in conjunctiva as positive control
39
Figure 3-6
Podoplanin-positive lymphatic channels in ciliary body
41
Figure 3-7
Podoplanin-positive lymphatic channels in sclera
42
Figure 3-8
Podoplanin-positive lymphatic channels in posterior orbit
43
Figure 3-9
Effect of latanoprost on IOP 44
Figure 3-10
In vivo hyperspectral imaging of mouse neck region 46
Figure 3-11
In vivo QD signal detection over post-injection imaging times 47
x
Figure 3-12
In vivo QD signal detection rate 48
Figure 3-13
Localization of QD in lymph node of latanoprost-treated and control mice
49
Figure 3-14
QD intensity in the submandibular node in latanoprost-treated and control groups
50
xi
Contributions
Zhexue (Josh) Zhang assisted in intracameral injection and initial in vivo imaging experiments.
Zazeba Chowdhury BSc. assisted in sectioning frozen tissues.
1
1
Introduction
1.1 Overview
Glaucoma is a leading cause of world blindness. The major risk factor is elevated
intraocular pressure (IOP) [1]. Current medical and surgical therapies attempt to lower IOP by
improving drainage through aqueous outflow pathways [2-4]. Lymphatics are channels known to
drain extracellular fluid, solutes and proteins in most organs. Lymphatics in the human [5] and
sheep eye [6] have been shown to contribute to aqueous humor (AqH) drainage. However, the
existence of ocular lymphatic drainage remains elusive in mouse. Since mouse is a valuable
model with similar AqH dynamics [7] and pharmacology [8, 9] as human, developing methods to
visualize ocular lymphatic flow in vivo in additional to investigating the effect of the most
commonly prescribed glaucoma drug, latanoprost, on the ocular lymphatic drainage would be
highly relevant to glaucoma studies.
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1.2 Glaucoma and Its Relation to Aqueous Humor Dynamics
Glaucoma is estimated to affect 60 million people in the year of 2010 to 80 million
people in 2020 [10]. A major risk factor for the development and progression of glaucoma is
elevated IOP [11]. High IOP leads to death of retinal ganglion cells (RGCs), damage of optic
nerve axons, and eventually blindness [11]. Lowering IOP with medicated eye drops helps to
preserve vision or slow progression of vision loss in glaucoma patients [12-14]. This is achieved
mainly through increasing AqH outflow from the eye [14].
In a healthy state, AqH outflow against resistance generates an average IOP around 16
mmHg in human eyes [15]. IOP is required to maintain the curvature of the cornea and the
refractive properties of the eye. IOP is determined by three factors: the rate of AqH production
by the ciliary body, the resistance to AqH outflow across the trabecular meshwork-Schlemm’s
canal system, and the level of episcleral vessel pressure [14]. Healthy IOP reflects a balance
between inflow and outflow of AqH.
1.3 Aqueous Humor - Composition, Formation and Dynamics
AqH is a transparent fluid contained in the anterior and posterior chambers of the eye. It
is responsible for the supply of nutrients to and removal of metabolic wastes from the avascular
tissue of the eye such as lactate, pyruvate, and carbon dioxide [16]. It is crucial for the
maintenance of the shape, IOP, and optical properties of the eye [17]. Moreover, it transports
ascorbic acid into the anterior segment of the eye to scavenge free radicals, and plays a role in
immune responses during inflammation and infection [18, 19].
The major components of AqH are organic and inorganic ions, carbohydrates, glutathione,
urea, amino acids and proteins, oxygen, carbon dioxide, and water. In a number of mammalian
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species [20-22], except from the eyes of rhesus monkeys [23], AqH was demonstrated to be
slightly hypertonic to plasma. The greatest differences in AqH relative to plasma, moreover, are
the concentrations of protein (20 times less) and ascorbate (20 to 50 times higher) [24]. The
protein content of AqH has both quantitative and qualitative differences compared to plasma.
Most AqH proteins are intrinsic glycoproteins of the vitreous, which are secreted by non-
pigmented ciliary epithelium (NPE) [25]. In addition, molecules that play a role in maintaining
the extracellular matrix, such as collagenase, have been identified in AqH and may influence
trabecular meshwork (TM) outflow resistance and, consequently, the IOP [26].
AqH is formed and secreted by the ciliary processes which are composed of a double
layer of epithelium over a core of stroma and a rich supply of fenestrated capillaries [27]. The
outer layer of pigmented ciliary epithelium (PE) faces the stroma while the inner NPE faces the
posterior chamber of the eye. The NPE cells contain numerous mitochondria, microvilli, and
Na+,K+-ATPase and are the main production site of AqH [28].
Three mechanisms are involved in AqH formation: diffusion, ultrafiltration and active
secretion [29]. Diffusion and ultrafiltration are responsible for the accumulation of plasma
ultrafiltrate in the stroma, behind the tight junctions of the NPE layer, from which the posterior
chamber AqH is derived [30, 31]. These two processes are passive without involvement of active
cellular participation. During the diffusion step, lipid soluble substances are transported through
the lipid portions of the membrane of the tissues between the capillaries and the posterior
chamber [32]. Next, with ultrafiltration, water and water soluble substances flow across
fenestrated ciliary capillary endothelium into the ciliary stroma [32].
4
Active secretion, which accounts for 80% to 90% of total AqH formation [33-37], is
comprised of three sequential steps. First, NaCl ions are transported into the PE cells from the
stroma via secondary active transport (paired Na+/H+ and Cl-/HCO3- antiports) [38, 39]. Next,
gap junctions between NPE and PE cells create a functional syncytium allowing for passage of
NaCl ions from PE to NPE [38, 39]. Finally, Na+ and Cl- ions are released into the posterior
chamber through Na+, K+-activated ATPase and Cl- channels [38, 39], which are influenced by
bicarbonate formation with the enzyme carbonic anhydrase through the conversion of CO2 and
H2O to HCO3- and H+ [40]. AqH then flows into the anterior chamber through the pupil [41],
around the anterior chamber, and into the three drainage pathways via the anterior chamber angle:
conventional TM outflow (pressure-dependent) [42, 43], unconventional uveoscleral (UVS)
outflow (pressure-independent) [32], and the ocular lymphatic drainage [5, 6].
1.3.1 Trabecular Meshwork Outflow (Fig. 1-1)
Most of the AqH in humans exits the eye through the TM outflow pathway. This pathway
is comprised of the TM, juxtacanalicular connective tissue, the endothelial lining of Schlemm’s
canal (SC), the collecting channels and the aqueous veins [44-46]. The TM is made up by the
uveal and corneoscleral meshworks, which form connective tissue lamellae or beams that are
covered by flat TM cells which rest on a basal lamina [44-46]. These beams attach to one another
in several layers, forming a porous filter-like structure to allow the passage of AqH. The TM
structures also provide the main resistance for AqH outflow to build up IOP [47]. When the IOP
is high enough, AqH will then flow across the TM into SC. Once in the lumen of SC, it was
demonstrated in the cynomolgus (Macaca fascicularis) monkey that AqH then flows into the
endothelial tubules that penetrate mostly through the sclera and join the episcleral venous plexus,
where AqH is finally drained into the venous system [48].
5
1.3.2 Uveoscleral Outflow (Fig. 1-1)
While a fraction of AqH leaves via the TM outflow pathway, part of the AqH leaves the
eye through the UVS pathway. The UVS outflow pathway is a drainage route for AqH from the
anterior chamber that successively includes extracellular spaces within the iris root, the ciliary
muscle, the anterior choroid and suprachoroidal space, and the adjacent sclera [49-52]. Fluid then
leaks into the surrounding periocular tissues directly through the sclera or through the loose
connective tissue surrounding penetrating blood vessels and nerves [53]. The amount of AqH
that leaves the eye via the UVS outflow routes varies in different species - mouse was reported to
be 82% [7], primates such as cynomolgus monkey 55% [54], sheep 22% [6], and cat 3% [55]. In
human eyes, various results were reported [56, 57].
The UVS pathway is relatively pressure-independent as it does not depend on the IOP to
the same extent as the TM outflow [58]. TM outflow increases linearly as IOP increases while
the UVS flow remains relatively constant. Nevertheless, at extreme conditions, such as removing
almost all of the resistance offered by the ciliary muscle through cyclodialysis, clear signs of a
pressure-dependence can be observed as the UVS flow was increased from 3 to 54% in rabbits
[59] and a four-fold increase was seen in monkeys [60].
The UVS flow can also be affected by the degree of contraction of the ciliary muscle [61].
It was observed that increasing the ciliary muscle tone with pilocarpine almost completely
blocked the UVS flow [62]. In fact, when combining the treatment with pilocarpine and PGF2α,
pilocarpine abolishes all the effects of PGF2α suggesting PGF2α increases flow through the ciliary
muscle [63]. Later experiments in cynomolgus monkey demonstrated that PGF2α and PGF2α
analog, latanoprost, increases UVS outflow [64, 65].
6
1.3.3 Ocular Lymphatic Drainage (Fig. 1-1)
1.3.3.1 Non-Ocular Lymphatics
While the majority of AqH leaves via the TM and UVS outflow pathway, a portion of the
AqH leaves the eye through the lymphatic pathway in human and sheep [5, 6]. The lymphatic
system is responsible for returning the interstitial protein-rich fluid, extravasated plasma proteins
and cells back to bloodstream. It also plays a role in immune surveillance of the body [66] and
absorbs lipids from the intestinal tract [67]. The lymphatic system is composed of a network of
collecting vessels known as initial lymphatics. They are blind-ended structures that lack
fenestrations, a continuous basal lamina, and pericytes [68]. Initial lymphatics are composed of a
single-cell layer of overlapping lymphatic endothelial cells (LECs) [69] which can be identified
by detecting molecules that are specifically expressed by LECs such as prospero-related
homeodomain transcription factor (Prox1) [70], vascular endothelial growth factor receptor-3
(VEGFR-3) [71], lymphatic vessel hyaluronan receptor (LYVE)-1 [72], and the membrane
glycoprotein podoplanin [73]. LECs exhibit large interendothelial pores [74, 75], which make
them highly permeable to large macromolecules, and migrating cells in the interstitium. Once the
interstitial fluid and proteins get into the initial lymphatics, they are collectively known as lymph
[76]. Lymph will then travel along the initial lymphatics, which coalesce into larger collecting
ducts. The collecting ducts are described as pre-nodal ducts if they lead to the lymph nodes and
as post-nodal ducts if they are drained away from lymph nodes. Collecting ducts contain a
continuous smooth muscular layer, an adventitial layer, a basement membrane, and valves to aid
in unidirectional flow [77]. The smooth muscle layer enables each functional unit of the
collecting ducts, lymphangion, to pump lymph [78]. Lymph from the left side of the head, the
left arm, left part of the chest region, and lower portion of the body enters the thoracic duct,
7
which in turn empties into the blood venous system at the juncture of the left internal jugular
vein and left subclavian vein [68]. Lymph from the right side of the neck and head, the right arm,
and parts of the right thorax enters the right lymphatic duct, which empties into the blood venous
system at the juncture of the right subclavian vein and internal jugular vein [68].
1.3.3.2 Ocular Lymphatics in Relation to Ocular Lymphatic Drainage
With the specific lymphatic endothelial cell markers, such as such as LYVE-1 [72], and
podoplanin [73] detected with D2-40 antibody [79], immunofluorescence staining and cryo-
immunogold electron microscopy were performed to reveal numerous fine lymphatic channels
through the ciliary body stroma in human and sheep eyes [5].
Intracamerally injected fluorescent tracers were localized in the lumen of lymphatic
channels, confirmed by LYVE-1 staining, in the ciliary body of the sheep [5]. When radioactive
tracers were injected into the anterior chamber of the sheep, they were drained to the head and
neck lymph nodes [5]. Subsequently, a sheep model was developed to quantitatively assess
lymphatic drainage along with TM and UVS outflows. Following intracameral injection of 125I-
bovine serum albumin (BSA), lymph and blood samples were continuously collected. Lymphatic
and TM drainage were quantitatively assessed by measuring 125I-BSA recovery. This quantitative
sheep model enabled assessment of relative contributions of lymphatic drainage (1.64% ±
0.89%), TM (68.86% ± 9.27%) and UVS outflows (19.87% ± 5.59%) [6]. Altogether, these
studies demonstrated the presence of lymphatic channels in the eye and the role of these
lymphatic channels played in fluid drainage from the eye.
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1.4 Glaucoma Treatment
The goal in glaucoma treatment is to prevent further loss of vision [80, 81]. The
preservation of vision is mainly achieved by lowering IOP to a level that is safe for the RGCs.
Topically applied ocular medications are usually the first step in controlling the IOP [14]. If eye
drops do not lower IOP sufficiently, then surgical procedures such as argon laser trabeculoplasty,
trabeculectomy, and shunts implantations will have to be performed with possible sight-
threatening post-surgical complications [82].
There are five classes of glaucoma drugs currently used to manage glaucoma. They are
either decreasing AqH production or increasing the aqueous outflow. They are categorized into
cholinergic agonists, α2-adrenergic agonists, inhibitors of carbonic anhydrase, β-adrenergic
antagonists, and prostaglandin analogs (PGAs).
Cholinergic agonist, also known as miotics, is the oldest effective pharmacological
treatment of glaucoma. The most commonly prescribed cholinergic compound is pilocarpine,
which lower IOP by stimulating the M3-type muscarinic receptors [83] of the ciliary muscle that
widens the anterior chamber angle, resulting in increased conventional TM outflow [84].
Pilocarpine typically reduces IOP by about 20 - 30% in human [85]. While pilocarpine is an
effective IOP-lowering agent, it is not commonly prescribed due to its side effects such as
decreased visual acuity due to pupillary constriction [86] and a 3 or 4 times daily dosage
requirement [84, 87, 88].
Adrenergic agonists in general decrease AqH production and increase UVS outflow [89,
90]. One of the α2-adrenergic agonists, apraclonidine hydrochloride (IOPIDINE® Ophthalmic
Solution), decreases AqH production as well as episcleral venous pressure and improves UVS
9
outflow [84, 89, 90]. Apraclonidine affects vascular tone and might cause vasoconstriction [84].
Apraclonidine, however, is rarely used for long-term therapy because of its high rate of allergic
blepharoconjunctivitis [91].
Carbonic anhydrase inhibitors (CAIs) decrease AqH production via enzymatic inhibition
[92]. In the CE, carbonic anhydrase isoenzyme II catalyses the conversion of CO2 and H2O to
HCO3- and H+, a process important for the production of AqH described previously. By
inhibiting this conversion, AqH formation is decreased. Dorzolamide was approved as the first
topical CAI for glaucoma therapy. At a dosage of three times daily, 2.0% ophthalmic solution
lowers IOP by 18-22% [93]. Nevertheless, local side effects from topical CAIs include stinging,
burning and itching still exist [93].
Beta-receptor antagonists reduce IOP by decreasing fluid formation through inhibition of
cyclic adenosine monophosphate production in CE in the ciliary body [94]. Most topical
ophthalmic β-adrenoceptor antagonists used today include timolol, levobunolol, metipranolol
and carteolol. While β-adrenoceptor antagonists are very effective IOP-reducing agents, their
popularity has declined due to their systemic side effects, such as exercise induced tachycardia, a
reduction of cardiac output, congestive heart failure, hypotension, and heart block [95, 96]. In
addition, depression, amnesia, and fatigue are the most common central nervous system adverse
effects [97].
Prostaglandin analogs are a relatively new class of ocular hypotensive agents and were
introduced into market in 1996. PGAs are currently the first-line treatment of glaucoma [98].
PGAs are all multicarbon-chain, lipophylic molecules, derived from arachidonic acid and sharing
similar structural features to PGF2α (C24H45NO8). The four different PGAs approved for clinical
10
use - latanoprost, unoprostone isopropyl, travoprost, and bimatoprost, are typically the most
potent class of IOP-reducing drugs available (Figure 1-2). They offer excellent control of diurnal
IOP fluctuation [99, 100]. Except for minor ocular effects such as iris pigmentation changes,
eyelash hypertrichosis, and ocular inflammation, they have few systemic side effects [101-103].
Amongst the PGAs, latanoprost is the only PGA to have received a formal first-line
usage approval from the FDA. Latanoprost (17-phenyl-13,14-dihydrotrinor PGF2α) is an analog
of the pro-drug PGF2α isopropyl ester. The pro-drug penetrates the cornea and becomes
biologically active after being hydrolyzed by corneal esterase [104]. Just like other PGAs,
latanoprost increases fluid drainage from the eye in glaucoma patients [9] and this is ascribed to
its action on the UVS pathway [105, 106] by loosening the intercellular spaces and induces
remodeling of extracellular matrix adjacent within the ciliary body [104]. Latanoprost has been
compared with timolol [101-103] and was found to be significantly more effective at lowering
IOP with a lower dosage. Mean IOP reduction was 6.7± 3.4 mm Hg for latanoprost compared
with 4.9 ± 2.9mm Hg for timolol after 6 months of treatment [101]. Moreover, latanoprost has
been shown to be significantly more effective at reducing IOP than other classes of glaucoma
medication such as dorzolamide [107] and brimonidine [108]. Most importantly, latanoprost has
no deleterious effect on blood pressure, pulse, and pulmonary function in patients [109].
1.5 Prostaglandins and Non-Ocular Lymphatics
Prostaglandins (PGs) (PGA2, PGB2, PGF2α) [110], along with various compounds, such
as serotonin [111], and thromboxane A2 [112], are known to stimulate the lymphatic smooth
muscle cells to increase the contractility; thereby, increasing the circulation of lymph. Of the PGs
that were shown to excite lymphatic smooth muscle cells, PGF2α has the greatest effects on
11
bovine mesenteric lymphatic smooth muscle cells [110], suggesting a significant effect of PGF2α
in facilitating lymph flow. As described previously, PGF2α analogs such as latanoprost are the
most potent, and most commonly prescribed worldwide to treat glaucoma. It is not clear whether
PGs act on the lymphatic outflow pathway of the eye. Developing a model to understand the
effect of latanoprost on the lymphatic drainage is critical in gaining insights into, and developing
novel treatment strategies for glaucoma.
1.6 Background on Using a Mouse Model
The mouse model is the focus of the experiments laid out in this thesis. Although
nonhuman primates most closely approximate human anatomy and physiology, experimentation
and handling of nonhuman primates can be challenging and is expensive [113]. Common
alternative models such as cats, dogs and rabbits are easier to handle and data for AqH dynamics
are relatively easy to gather [114-116]. However, in those models, only a small proportion of
total outflow exists via the UVS routes, with less than 10% of total outflow in cats [55] and
rabbits [59] and around 15% in dogs [117]. Since ocular lymphatic channels and drainage have
been demonstrated in the ciliary body in human and sheep [5, 6], it is possible that the ocular
lymphatic drainage is closely related to the UVS outflow. With only a small proportion of total
outflow via the UVS, cats, rabbits, and dogs will be expected to have even less proportion of
total outflow via the ocular lymphatic drainage. Hence, these species may not be as helpful in
understanding the ocular lymphatic drainage as mice, whose total outflow exits via the UVS was
reported to be 82% [7].
The mouse model also offers several advantages for investigating ocular diseases because
of the availability of transgenic mice, their similar genetic background and relatively quick
12
breeding [118]. The low cost and ease of handling of this species allows for large-scale
experiments to be performed. With the development of techniques that permit reproducible
measurement of aqueous flow, episcleral venous pressure, and total outflow facility in the mouse
[7, 119] as well as reliable measurement of IOP [120-125], this species becomes an ideal species
for the study of AqH dynamics. The mouse has a normal IOP range averaging approximately 10
to 20 mmHg, which is similar to that of healthy human eyes at 16 ± 3 mmHg [126]. Compared to
the common models such as cows and rabbits, mice have better developed TM and SC in the eye
[127]. In fact, mouse and human eyes have similar anatomic structure, outflow pathways, and
response to IOP-lowering drugs [8, 128]. It has been demonstrated that topical application of
PGF2α, which lowers IOP and increases UVS outflow in monkey and human eyes [9, 64], also
lowers IOP in mouse eyes [8]. Finally, advances in in vivo imaging modules and nanotechnology
have enabled visualization of the lymphatic drainage in small animals, including mice.
1.7 Justification of In Vivo Imaging for Study of Lymphatics
1.7.1 Conventional Lymphatic Drainage Imaging
Conventional lymphatic imaging involves the use of radionuclide scintigraphy and dyes.
These methods can effectively image lymphatic vessels and delineate lymphatic drainage.
Lymphoscintigraphy is a special type of radionuclide imaging which requires injection of
gamma-emitting radionuclides labeled macromolecules into a local region of tissue to evaluate
lymph drainage function [129-131]. For imaging with dyes, such as isosulfan blue, patent blue V,
Evans blue or fluorescent dyes, dye molecules are typically injected intradermally or
subcutaneously into the interstitial tissue of animals or human beings. A visual signal of the
draining lymphatic vessels and lymph nodes can then be obtained from indirect micro-
13
lymphangiographies of superficial lymphatic vessels [132]. Cutaneous lymphatic vessels and
lymphatic drainage from the skin can also be macroscopically visualized with this method [132-
134]. However, there are various drawbacks for these methods. Lymphoscintigraphy has
radiation exposure while both techniques have relatively low resolution. Moreover, the intrinsic
fluorescence from dyes has very little penetration through tissue. Hence, invasive procedures
such as incisions to expose organs or tissues of interests must be performed to visualize
lymphatic drainage.
1.7.1.1 Cannulation of Lymphatics to Detect Injected Radioactive Tracers
In human, radioactive lymphography through direct cannulation of lymphatics to
visualize lymphatic drainage is now abandoned due to the difficult and potentially life-
threatening procedures [135]. In animals such as sheep [6], rabbits and cats [136], however,
lymphatic drainage was studied by cannulation of lymphatic ducts. Following radioactive tracers
injection into site of interest, continual collection of lymph through the cannula can provide
quantitative data for understanding the role of lymphatic system in aqueous or cerebrospinal
drainage. However, cannulation of lymphatics is difficult to perform in small animals such as
mouse due to the small size of lymphatic vessels and the invasive nature of the procedure.
Therefore, further non-invasive techniques are required to study lymphatic drainage in small
animals.
1.7.2 Non-invasive In Vivo Imaging of Lymphatic System with Organic Fluorophores
With the advances of optical imaging, lymphatic system can now be studied non-
invasively using organic fluorophores. In vivo optical imaging allows for imaging at molecular
level, mapping of lymphatic drainage, and visualization of multiple nodes at a relatively high
14
resolution without radiation exposure [135]. Fluorophore such as indocyanine green has been
approved in humans for assessing cardiac and hepatic function, and ophthalmic angiography
[137-141]. Studies have also shown the potential of this dye to image lymphatics in animal
models and in humans [142]. The principal downsides of optical imaging agents remain low
depth penetration [143] and having emission spectra overlapping that of autofluorescence
generated by tissue and melanin [144]. Probes that emit in the near infrared (NIR),
approximately 600–800 nm, help to maximize the target to background ratio and have improved
depth penetration [145] as cells, tissues, and biological fluids autofluoresce minimally when
stimulated in the NIR [146]. NIR organic fluorophores such as Alexa Fluor 705, IRDye780, Cy7,
and Cy5.5 thus can be used to detect lymphatic drainage. These organic fluorophores can also be
conjugated with antibodies, proteins, and peptides to effectively enter and remain in the
lymphatic system [143, 147]. However, optical resolution with organic fluorophores is hampered
by narrow excitation spectra, broad emission spectra, and susceptibility to photobleaching [148].
Hence, the design of high-emission fluorophores is necessary for both animal research and
clinical imaging applications [147]. The recent advance of nanotechnology has led to the
development of quantum dots (QDs) that circumvent these drawbacks.
1.7.3 Non-invasive In Vivo Imaging of Small Animals with QDs
The unique physical characteristics of QDs such as broad excitation spectra, narrow size-
tunable emission spectra and a high photobleaching threshold make them suitable for non-
invasive in vivo imaging. Optical properties of QDs such as CdSe-ZnS core-shell nanoparticles
are advantageous for long-term single or simultaneous multiple color imaging of live cells [149]
and animals [150, 151]. QDs can be detected by hyperspectral imaging due to an intrinsic
brightness greater than regular organic fluorescent dye [150, 152]. QD655 with a hydrodynamic
15
size of 19 nm provides optimal uptake and retention inside lymphatic vessels [151] and lymph
nodes [153]. The emission spectrum of 655 nm penetrates a depth of up to 2 centimeters [151]
and can be distinguished from autofluorescence mainly produced by melanin, which has an
emission range of 360–560 nm [154], in pigmented animals [144].
1.8 Rationale and Hypothesis
1.8.1 Rationale
Lymphatic channels have previously been reported in sheep and human eyes [5]. The
contribution of lymphatic pathway to AqH drainage from the sheep eyes has been quantified [6].
However, the existence of ocular lymphatic drainage and lymphatic channels are relatively
unexplored in the mouse models. Since the ocular anatomy and aqueous dynamics between
human and mouse are similar [7], in addition to the advantages of mouse models in glaucoma
research [8, 155], experiments were undertaken in the first part of this thesis to examine the
existence of a lymphatic drainage and lymphatic channels in the mouse eye.
Among the anti-glaucoma drugs, PGF2α analogs are the most potent, and the most
commonly prescribed worldwide. Latanoprost, an PGF2α analog, increases fluid drainage from
the eye in glaucoma patients [9]. Increased fluid drainage from the eye is ascribed to the action
of latanoprost on the UVS pathway [106]. Although PGAs are well established to stimulate the
lymphatic drainage in non-ocular tissues [156-158], it is not clear whether PGs act on the
lymphatic outflow pathway [5]. Based on the demonstration of ocular lymphatic drainage from
the mouse model [159], we aimed to determine the effect of latanoprost on the lymphatic
drainage.
16
1.8.2 Hypotheses
Hypothesis 1
The first hypothesis is that lymphatic drainage from the mouse eye exists. The hypothesis
will be tested through intracameral injection of QDs followed by in vivo hyperspectral imaging
over a period of imaging time points to visualize the trajectory of the tracers.
Hypothesis 2
The second hypothesis is that lymphatic channels exist in the mouse eye and surrounding
orbital tissue. This will be determined by performing immunofluorescence staining frozen eye
and orbit sections for lymphatic channels and blood vessels with anti-podoplanin and anti-
collagen IV antibodies, respectively. Stained sections will then be examined through confocal
and bright field microscopy.
Hypothesis 3
The third hypothesis is that a PGF2α analog, latanoprost, stimulates lymphatic drainage
from the mouse eye. The in vivo lymphatic drainage rate will be compared through topical
application of latanoprost or artificial tear as control, followed by intracameral injection of QDs
and in vivo hyperspectral imaging. QD signal intensity will also be quantified in tissue sections
using post-mortem hyperspectral imaging.
17
Figure 1-1. Aqueous Humor Outflow Pathways. AqH is produced by the ciliary body and it flows from the posterior chamber through the pupil into the anterior chamber. From there it flows out via three pathways at the anterior chamber angle: conventional outflow (red arrow) via the TM and SC; the unconventional UVS route (blue arrow) via the ciliary muscle and sclera; and the ocular lymphatic drainage pathway (green arrow) via the lymphatic channels in the ciliary body. (Adapted from reference [160])
18
Figure 1-2. Chemical structures of PGF2α and commercially available PGAs.
19
2
Material and Methods
2.1 Subjects
The experiments reported in this thesis were designed and performed in accordance with the
guidelines of the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research
with the approval by the institutional Animal Care Committee. The experiments utilized
randomly bred male pigmented mice (129SVE from Taconic, Hudson, NY) with weight ranging
from 19g to 28g. Housed under constant 12 light/dark cycles, they were provided with standard
food and water ad libitum including the day before experiments. In the study of identification of
ocular lymphatic drainage and channels in mouse, 27 mice, inclusive of the mice utilized in
control experiments, were used. In the study of the effects of latanoprost on the ocular lymphatic
drainage, 10 mice were used to determine the effects of latanoprost on IOP, 14 mice were used in
the latanoprost-treated group and 14 were used as control group for in vivo imaging of ocular
lymphatic drainage.
20
2.2 Quantum Dots
The nanotracers used in the experiments reported in this thesis are QDs (QD655, Invitrogen,
Eugene, OR, USA) with core/shell CdSe/ZnS, are ellipsoid (6 nm x 12 nm), with maximal
emission at 655nm. QDs were coated with carboxylic acids and negatively charged (QD-COOH,
pH 9.0). Three μL of 8 μM solution in a 50 mM borate buffer was used for intracameral injection.
2.3 Topical Application of Substances Prior to QD Tracer Injection
In the study of mouse ocular lymphatic drainage, three drops of artificial tears (Refresh tears,
Allergan Inc., ON, Canada) were topically administrated 17 hours before and 1 hour to 24 mice
before the experiment took place to ensure the eyes stay moisturized. In the study of the effect of
latanoprost, 28 mice were split into two groups. In one group, 3 drops of latanoprost 0.005%
(Xalatan, Pfizer, QC, Canada) (n=14) were instilled to the eyes 17 hours and 1 hour prior to the
tracer injection. The 14 mice instilled with artificial tears described in the mouse ocular
lymphatic drainage experiments were used as the control group.
2.4 Intracameral Injection of QD Tracer
Intracameral tracer injections were performed under general anaesthesia using either inhalation
of 3% or 1.5% isoflurane (Abbott Labs Inc., Saint-Laurent, QC, Canada), in 70% N2O and
balance of O2. The hair on the head, neck, abdomen regions and forelimbs were gently shaved
with a hair trimmer (Chromini Type 1591, Wahl, Sterling, IL, USA) [144]. Under an operating
microscope (Leica M655, Wetzlar, Germany), 3 μL of QD655 was injected into the anterior
chamber of the left eye using a 33 gauge needle (Hamilton Company, Reno, NV, USA) and
either a 100 μL or 25 μL Hamilton syringe (Hamilton Company, Reno, NV, USA). QD toxicity
suggested by in vitro studies [161, 162] was not specifically studied, though acute signs of ocular
toxicity were not observed.
21
2.5 IOP Measurement
Under general anaesthesia as described above, IOP was measured non-invasively using a
handheld rebound tonometer (TonoLab, Type TV02, Icare Finland Oy, Helsinki, Finland) [120-
125] before the tracer injection and at the end of experiments to monitor any IOP changes for all
in vivo imaging experiments in this thesis. Generally, the tonometer probe was applied
perpendicularly to the centre of the cornea under operating microscope. Multiple readings were
taken until six steady consecutive measurements could be obtained, and the mean value of these
six measurements was calculated. No statistically significant difference in IOP was observed
between measurements performed before tracer injection and 6 hours after tracer injection for
latanoprost-treated and control mice (latanoprost-treated group: before tracer injection: 17.9 ±
6.6 mmHg and 6 hours after tracer injection: 15.1 ± 5.2 mmHg, t -test, p>0.2; control group:
before: 17.6 ± 4.2 mmHg, and 6 hours after tracer injection 15.5 ± 4.5 mmHg, t-test, p>0.2).
For experiments performed to confirm the IOP reduction following topical application of
latanoprost in 10 randomly bred male pigmented mice (Figure 2-1). One drop of 0.005%
latanoprost (Cayman Chemical Co., Ann Arbor, MI, USA) or 0.02% benzalknonium chloride in
1x phosphate buffered saline (PBS) (Sigma-Aldrich, St. Louis, MO, USA) was topically applied
to the left eye at 5 pm and 16 hours later. Right and left eye IOPs were measured prior to each
instillation, and 1, 1.5, 2, 3 and 5 hours following last instillation. All IOPs were measured one
minute after induction of general anesthesia to minimize the effects of anesthesia on IOP [8]. The
effect of isoflurane on IOP during in vivo imaging was not studied in the present experiments.
The IOP decrease in the latanoprost-treated group (n=5) was compared to that in the control
group (n=5). Mean IOP in latanoprost-treated mice was compared to controls at each time point
using two-sample t-tests assuming equal variances.
22
Figure 2-1. Timeline for IOP measurements. Timeline for inhalation anesthesia (IN), IOP
measurement (IOP), and topical application (TA).
2.6 In Vivo Hyperspectral Fluorescence Imaging [163, 164] (Figure 2-2)
Prior to in vivo imaging, a hair trimmer (Chromini Type 1591, Wahl, Sterling, IL, USA) was
used to shave the head, neck, abdominal and forelimb areas [144]. Under general anaesthesia in
vivo hyperspectral fluorescence imaging was performed before injection, and various time points
after injection of QD into the anterior chamber of the left eye. Mice were anesthetized through
induction of isoflurane for 60 sec in a chamber and maintenance through an air tube for 3 - 4
minutes during hyperspectral system imaging (Maestro™, Cambridge Research &
Instrumentation Inc., Woburn, MA, USA). Anesthesia was discontinued after the imaging and
mice were placed back in their cage between imaging sessions. The anesthesia was repeated only
during the imaging sessions at various time points. The in vivo imaging time points in the study
of identification of lymphatic drainage were: before injection, and 6 and 24 hours after injection
(n=3) or before injection, 5, 20, 40, and 70 minutes; 2 and 6 hours after injection (n=14) of QD
into the anterior chamber of the left eye. In determining the effect of latanoprost, the time points
were before tracer injection, and at 5, 20, 40, 70, 120, and 360 min after injection (n=28). Two
subjects were excluded from analysis due to Maestro – computer interface problems (Animal
23
ID#16, 17), with 4 excluded due to QD leakage from the eye following injection (#24, 25, 28,
29). Time to QD detection was defined as the earliest in vivo detection of QD signal to the neck
region after eye injection. QD signal detection rate (60/time to in vivo detection) (hours-1) was
calculated to assess lymphatic drainage.
All mice were monitored throughout the experiment. No abnormal behavior was observed due to
general anesthesia. The excitation filter and the emission filter were 503–555 nm, 580 nm long
pass, respectively. The tunable filter was automatically stepped in 10-nm increments from 500 to
800 nm. The exposure time was 900 ms. Animals under anaesthesia were gently placed in a
light-tight chamber. Captured images were analyzed using the Maestro 2.4 Imaging Software
(Cambridge Research & Instrumentation Inc., Woburn, MA, USA), with unmixing algorithms to
separate autofluorescence from QD signal [165, 166]. Green signal was set to represent spectra
of autofluorescence mainly from melanin [144] while red signal was set to represent the QDs
spectrum.
24
Figure 2-2. Timeline for in vivo hyperspectral imaging. Timeline for inhalation anesthesia
(IN), topical application (TA), IOP measurement (IOP), intracameral injection (IC), in vivo
hyperspectral imaging (HI), and euthanasia (E).
2.7 Control Experiments for QD drainage
To ensure QDs drain similarly in the right eye, QDs were injected into the anterior chamber of
the right eye in 5 mice. In vivo imaging was performed before injection, 2 and 6 hours after
injection (n=5). Two mice were excluded from analysis due to injection problems (#2, 5)
Because some leakage from anterior chamber into the conjunctiva can occur during intracameral
injections, control experiments were conducted to verify that QDs draining to the lymph node
was not due to conjunctival lymphatics. To test this, QDs were topically administrated onto the
conjunctiva (n=5). In vivo imaging was performed before injection, 2 and 6 hours after injection.
2.8 Tissue Collection and Processing
2.8.1 Head and Neck Specimen
Mice were sacrificed with CO2 inhalation at either 29 (n=3 from study in identification of
lymphatic drainage) or 6 hours (n=14 for latanoprost-treated, n=14 for tear-treated, n=10 in
control experiments) after injection of QDs. The upper body including head and neck, thorax and
front limbs was dissected. Specimens were immersion-fixed in 4% paraformaldehyde (Electron
25
Microscopy Sciences, Hatfield, PA, USA) for 48 h at 4°C. The specimens were then
cryoprotected by immersion in 10% glycerol (Bioshop, Burlington, ON, Canada) and 2%
dimethyl sulfoxide (Fisher Scientific Company, Ottawa, ON, Canada) in 0.1 M phosphate buffer
for 4 days and 20% glycerol and 2% dimethyl sulfoxide in 0.1 M phosphate buffer for 6 days.
The upper body of mouse was scanned using Maestro™ for the location of QDs after skin
removal to get rid of the autofluorescence signal generated by the melanin pigment in hair and
skin [144]. Neck tissues containing QD signal were harvested. The tissue block and the
remainder of the upper body were scanned again to confirm the presence of QD signal and its
absence, respectively. Mouse heads were harvested followed scanning. To orient the neck tissue
block, its apex and right side were marked by red and blue surgical pathology ink, respectively
(Triangle Biomedical Sciences, Inc., Durham, NC, USA).
2.8.2 Orbit Specimen
Orbit specimens were collected and cryoprotected in an identical manner as the head and neck
tissue. To orient the orbit tissue block, its superior and nasal side were marked by yellow and
blue surgical pathology ink, respectively.
2.9 Frozen Sections
2.9.1 Head and Neck Specimen
Tissue blocks from the neck region [167] containing QD signal were frozen in isopentane cooled
by dry ice [5] and embedded into cryomatrix resin (Shandon Cryomatrix, Thermo Scientific, MI,
USA). All blocks were serially sectioned (140 μm thick) using a sliding microtome (Leica
DM2400, Leica, Germany). Sections were mounted onto double-frosted glass slides (Fisher
Scientific Company, Ottawa, ON, Canada) with a PVA-DABCO (Polyvinyl alcohol-1,4-
26
diazabicylo[2.2.2]octane) anti-fade mounting medium (Sigma-Aldrich, St. Louis, MO, USA). All
sections were imaged under the Maestro™ and sections with QD signal were selected for
histological analysis.
2.9.2 Orbit Specimen
Mouse orbit specimens were frozen in isopentane cooled by dry ice and embedded into
cryomatrix resin. Blocks were serially sectioned (140μm thick) through the coronal plane using a
sliding microtome. Sections were mounted onto double-frosted glass slides with a PVA-DABCO
anti-fade mounting medium.
2.10 Quantification of Tracer Drained into Lymph Node
Serial neck region sections scanned by hyperspectral imaging (450 ms exposure time) were
analyzed in a masked manner for QD signal intensity on 696 x 520 pixel size images (Image J
1.43 u National Institutes of Health, Bethesda, USA). The area and mean gray value of the region
with QD signal was measured using thresholds of 60 and 255, respectively. QD signal intensity
for each mouse was calculated by adding the product of the area and mean gray values in serial
sections with QD signal.
2.11 Immunofluorescence and Nuclear Staining
2.11.1 Submandibular Lymph Node
To verify the presence of QDs in lymph node, selected sections containing the submandibular
lymph node were washed three times in 0.1 mol 1x PBS (pH 7.4) for 6 min each, and incubated
for 10 min in 0.2% TritonX-100 (Fisher Biotech, NJ, USA) in PBS. After three more washes in
PBS for 6 min each, sections were incubated in 3% hydrogen peroxide (EMD, Darmstadt,
27
Germany) in PBS for 15 min. Sections were then washed 3 times for 6-min in PBS, and
incubated for 1 h with 1% blocking reagent (Tyramide signal amplification (TSA) kit, Invitrogen,
Oakville, ON, Canada) in PBS. Sections were incubated in collagen IV antibody (1:100, rabbit
polyclonal, Abcam, Inc., Cambridge, MA, USA) with 1% blocking reagent overnight at 4°C.
After three 5-min washes in PBS, sections were treated in the dark with a 1:100 dilution of the
Tyramide Alexa Fluor 555 (TSA kit) for 10 min in amplification buffer. Sections were washed
three times in PBS for 5 min and incubated for 20 min with a 1:1000 dilution of Sytox Green (a
nuclear staining, Invitrogen, Eugene, OR, USA). Sections were washed three times in PBS for 6
min each before cover-slipping with a PVA-DABCO anti-fade mounting medium as described
above. All sections were treated at room temperature and with mild agitation at all steps unless
otherwise noted. Negative controls were obtained by omitting primary antibody.
2.11.2 Right Eye and Orbit Specimen
Frozen right eye and orbit sections (n=5) were selected for immunofluorescence and nuclear
staining based on the following criteria: 1) conjunctiva, ciliary body, sclera, and orbital tissue
must be present under examination by bright field microscopy; and 2) ocular structures must be
intact and not fragmented. Immunofluorescence and nuclear staining were performed as
described above except primary antibody used to label lymphatic channels and blood vessels
were podoplanin (1:100, hamster polyclonal, eBioscience, San Diego, CA, USA) and collagen
IV antibody (1:100, rabbit polyclonal, Abcam, Inc., Cambridge, MA, USA), respectively.
Secondary antibody targeting podoplanin and collagen IV were goat anti-hamster Alexa Fluor
488 and anti-rabbit Alexa Fluor 647 (1:100 dilution, Invitrogen, Oakville, ON, Canada) ,
respectively. Sections were incubated for 20 min with a 1:500 dilution of propidium iodide.
28
2.12 Bright Field Microscopy
Hematoxylin and eosin (H&E) stained sections were viewed with upright brightfield microscope
(BX51,Olympus, Tokyo, Japan). Images were taken with MicroFire® True Color Firewire
Microscope Digital CCD Camera 1600 x 1200P (Optronics, California, USA). Exposure times
were set to automatic.
2.13 Combination of Fluorescence and Bright Field Microscopy
To visualize and confirm the QDs outflow pathway in the left injected eye and orbit, frozen
coronal sections (140µm thick) of left orbit specimens from control group (n=14) were examined
using an upright fluorescence microscope (BX51, Olympus, Tokyo, Japan). Two subjects were
excluded due to QD leakage from the eye following injection (#24, 28). A DAPI (4',6-diamidino-
2-phenylindole) filter was used to capture blue emission signal from autofluorescence of tissues
while a TRITC (Tetramethylrhodamine-5-(and-6)-Isothiocyanate) filter was used to capture red
emission from QD signal in left orbit specimens only. Images were captured with a DP72 digital
camera using cellSens software (Olympus, PA, USA) and a 4x, 10x, 20x, or 40x objective lens.
Bright field images were superimposed on fluorescence images to visualize tissue structure.
Exposure times for QDs and tissue in left orbit specimens were manually set to avoid over
saturation of signals (Table 2-1).
In right orbit sections without QD signal, exposure time for tissue was manually set to 15ms with
40x objective lens.
29
Table 2-1. Exposure times (ms) at various magnifications for frozen sections of orbit specimen.
Objective lens Tissue QDs
4x 50 50 - 75
10x 20 - 25 10 - 30
20x 5 - 20 1 - 25
40x 5 - 10 0.5 - 10
2.14 Confocal Microscopy
Confocal microscopy was utilized to verify the presence of QDs in lymph node, and to identify
lymphatic channels in the eye and orbit specimen from immunofluorescence-labeled sections.
Stained sections were viewed with a confocal laser scanning microscope (TCS SL, Leica,
Wetzlar, Germany) and images of 1024 x 1024 pixel resolution were captured. Excitation
wavelengths used were 488 nm for Alexa Fluor 488 and Sytox green nuclear stain, 543 nm for
Alexa Fluor 555, and 633nm for Alex Fluor 647. QD655 was excited with the shortest available
wavelength, 488 nm, and the emission window was specifically set at 620 – 680 nm to capture
its maximum emission spectrum.
2.15 Statistical Analysis
Mean QD signal detection rate in latanoprost-treated mice was compared to controls using two-
sample t-tests assuming unequal variances. Mean total QD intensity (log scale) in the latanoprost
30
group was compared to the control group using two-sample t-tests assuming equal variances.
P<0.05 was considered statistically significant. Statistical analysis was carried out with
Microsoft Excel (Version: 14.0.6112.5000; Microsoft Office Professional Plus 2010, Microsoft
Corporation, Seattle, WA, USA). Results were graphed with GraphPad Prism (GraphPad
Software, Inc., La Jolla, CA, USA).
31
3
Results
Whole body in vivo imaging confirmed the presence of strong QD signal in the left eye at
5, 20, 40, and 70 minutes, 2 hours after tracer injection. Signals were consistent with the QD
spectral profile (Figure 3-1D,H). At these time intervals, no QD signals were detected in other
parts of the body. At six hours after injection, while a strong QD signal was still detected in the
injected left eye (Figure 3-1B), a focus of QD signal was noted only in the left neck region in 14
out of 17 mice (Figure 3-1F). The spectral analysis of the signal in the left eye and left neck
confirmed the QD spectral profile (Figure 3-1D,H). Twenty-four hours following injection, the
strong QD signal was still observed in the left neck region (Figure 3-1G), while QD signal
intensity observed in the left eye was reduced (Figure 3-1C).
A consistent background pattern of green signal from facial and limb hair, and faint
orange signal from mouth, forelimb and ear was observed (Figure 3-1A-C, E-G).
32
Control Experiments
QD signal was noted only in the right neck region 6 hours following injection in all mice
(n=3) successfully injected with QDs in the right anterior chamber. Post-mortem imaging with
skin removal confirmed, in all mice (n=3), the presence of QDs in the neck tissue where the right
submandibular lymph node was located.
No QD signal was observed during the in vivo imaging sessions in the head and neck
region of the control mice (n=5) with QDs were topically applied onto the conjunctiva. Moreover,
QD signal was not observed in anywhere else in the body except remaining at the site of topical
application. Post-mortem imaging with skin removal revealed no QD signal in the head and neck
tissue or in any organ or tissue for all mice (n=5).
33
Figure 3-1. In vivo hyperspectral fluorescence imaging of head and neck in dorsal (A-C) and ventral (E-G) views. Imaging performed prior to QD injection (A) shows no signal in the left eye (arrow). At 6h (B) and 24h (C) following injection, QD signal in red is noted in the left eye (arrow). Strong QD signal is noted in the left neck region (arrow) at 6h (F) and 24h (G), and is not seen prior to injection (E). Scale = 10 mm. Emission spectral profiles of the signals in the eye (arrow in B) and in the left neck region (arrow in F) at 6 hours are shown in dotted lines in (D) and (H), respectively. Spectral profiles of both signals confirm the QD spectral profile (red), compared to the background signal profile (green) (D, H).
34
Having observed QD signals draining from the anterior chamber to the neck tissue, we
next aimed to determine the anatomical structures where the intracamerally injected QDs drained.
Based on post-mortem imaging using hyperspectral imaging, intense QD signals were observed
within a block of neck tissue on the left side (Figure 3-2A) after skin removal. This soft tissue
area adjacent to the submandibular salivary gland was isolated (Figure 3-2B,C). Hematoxylin
and eosin (H&E) stained sections revealed classic lymph node architecture (Figure 3-2D).
Figure 3-2. Post-mortem analysis of neck tissue. Neck tissue containing QD signals (red) (A) was harvested (B), and isolated (C). Hematoxylin and eosin stained section shows a lymph node (D). Scale = 10 mm (A-C) and 250 μm (D).
35
To further confirm our histological findings, immunofluorescence and confocal
microscopy were performed on frozen sections of the neck tissue containing the submandibular
lymph node. QDs in red were detected inside the submandibular lymph node (Figure 3-3).
Immunofluorescence labeling with antibody against collagen IV outlined the morphology of a
lymph node (Figure 3-3) while counterstaining with Sytox green demonstrated architecture of the
node (Figure 3-3). QDs were consistently detected in the cortex region at the subcapsular sinus
close to one pole of the submandibular lymph node. No QD signal was noted in the paracortex or
medulla regions.
Figure 3-3. Immunofluorescence of left submandibular lymph node. QDs in red are contained within the lymph node surrounded by capsule in blue (anti-collagen IV) against a green background of cell nuclei of the node parenchyma (Sytox green). Scale = 250 μm.
36
Altogether, the data suggests a lymphatic drainage from the mouse eye draining QDs
from the left anterior chamber to the left submandibular lymph node. Nevertheless, we cannot
exclude the possibility that the Maestro was unable to detect minute amount of QDs that is below
detection threshold in the contralateral lymph node or elsewhere in the body. To further
understand this lymphatic drainage, we first set out to determine the locations within in the eye
and orbit of where QDs were drained from the anterior chamber. Through examination by
fluorescence and bright field microscopy, of frozen coronal sections of left orbit specimen from
the control group (n=12), we observed QD drainage from the anterior chamber to various ocular
compartments 6 hours following QD injection. QDs were observed in ciliary body (Figure 3-
4A,B) in all 12 mice, in choroid in 11 of 12 mice (Fig. 3-4C,D), and in the sclera of all mice
(Figure 3-4C,D). QDs were observed in the posterior orbit in 10 of 12 mice (Figure 3-4E,F). The
signal appeared to be a continuation from choroid, to sclera, and to orbital tissue and onward.
Although QD signal was observed in the sclera, we were not able to distinguish whether the QDs
were located in the anterior or posterior portion of sclera based on coronal sections. Transverse
sections will be needed to pinpoint the portion of sclera into which QDs were drained. QDs were
not observed in the lens, retina, or vitreous. QD signal was not observed in control non-injected
right eye and orbit specimens. In the frozen sections, lymphatic or blood vessels could not be
observed under the magnification of the bright field microscope. As such, further studies with
higher magnification are needed to determine whether or not QDs are located in ocular lymphatic
channels and that intraocular lymphatics play a role in draining the QDs from the anterior
chamber to the submandibular lymph node. However, the existence of lymphatic channels in the
mouse eye has not been explored. This opened up the opportunity for us to first identify whether
lymphatic channels exist in the mouse eye.
37
Figure 3-4. QD distribution in left eye 6h following intracameral injection in control mice treated with artificial tears. Merged images from fluorescence and light microscopy showed QDs (red), uveal tissue (black), all other ocular tissues (blue) and optically empty space between tissues (bright field). QDs observed in boxes in panels A, C, and E are shown at higher magnification in B, D, and F, respectively. Scale bar = 500 μm (A,C,E), 100 μm (B,F), and 50 μm (D). Ciliary body (CB), lens (L), choroid (C), sclera (S), retina (R), and orbit (O). (Published in supplementary material in reference [168])
38
With the observation of QD signals in the ciliary body, sclera, and orbit (Figure 3-4), we
focused on identifying lymphatic channels in these structures. The morphologic criteria used to
establish a structure as a lymphatic vessel were as follows: 1) endothelial cell-lined
immunostained vessel; 2) a very thin-walled vessel with central lumen, usually with a wavy,
irregular contour and often collapsed; and 3) a poorly developed or absent basal lamina indicated
by absence of collagen IV staining.
With the criteria above, we first made sure the primary antibodies, podoplanin and
collagen IV, were specifically targeting lymphatic channels and blood vessels, respectively. We
selected sections with conjunctiva as a positive control. The morphology of conjunctiva was
confirmed by collagen IV-positive basement membrane and multiple layers of nucleus
counterstained by propidium iodide (Figure 3-5A). Conjunctiva sections showed podoplanin-
immunoreactive lymphatic channels that were distinct from collagen IV-positive conjunctival
blood vessels in 5 out of 5 subjects (Figure 3-5B). Negative controls omitting primary antibody
showed no non-specific staining (Figure 3-5C).
39
Figure 3-5. Podoplanin-positive lymphatic channels in conjunctiva as positive control (A) Conjunctiva immunostained with collagen IV antibody (blue), counterstained with PI (red) (arrow indicates the substantia propria where lymphatics are seen). (B) A podoplanin-positive lymphatic channel in green (arrow) is distinct from a collagen IV-positive blood vessel in blue (asterisk). Both signals are absent in the negative control without primary antibodies (C). Scale bar = 30 μm (A), and 10 μm (B,C).
40
Having determined the specificity of the primary antibodies, we examined the existence
of lymphatic channels in the ocular structures in the following order: ciliary body, sclera, and
orbital tissue. Within the ciliary body stroma, podoplanin-immunoreactive lymphatic channels
were noted (Figure 3-6B) and they were distinct from collagen IV-positive blood vessels (Figure
3-6C). These lymphatic channels were thin-walled vessels without collagen IV staining.
Lymphatic channels were observed in the ciliary body of all mice (n=5). Non-specific staining
was absent in negative controls omitting primary antibody (Figure 3-6D).
Podoplanin-positive lymphatic channels were observed in the episclera region (Figure 3-
7A,B), and were distinct from collagen IV-positive blood vessels (Figure 3-7C). These lymphatic
channels were endothelial cell-lined vessels that had a central lumen and wavy and thin-walled
contour. They lacked basal lamina indicated by absence of collagen IV staining. Lymphatic
channels were noted in 3 out of 5 mice. Negative controls omitting primary antibody showed no
non-specific staining (Figure 3-7D).
Finally, podoplanin-positive lymphatic channels were found in the orbital tissue. The
Harderian gland, which was surrounded by a fine connective tissue capsule stained by collagen
IV, was used as a landmark to locate the posterior orbit (Figure 3-8A). Podoplanin-
immunoreactive lymphatic channels were observed in the posterior orbit, adjacent to the
Harderian gland, in 5 out of 5 subjects (Figure 3-8B). These orbital lymphatic channels were
distinct from collagen IV-positive blood vessels (Figure 3-8B). Lymphatic channels in the
posterior orbit were thin-walled, were collapsed in contrast to the blood vessels with rounder
contour, and were not collapsed. Non-specific staining was absent in negative controls (Figure 3-
8C). Other regions of the posterior orbit were not examined in this study; thereby, we cannot
exclude the possibility of the existence of lymphatic channels elsewhere in the orbit.
41
Figure 3-6. Podoplanin-positive lymphatic channels in ciliary body. (A) Ciliary body immunostained with collagen IV (blue) and counterstained with PI (red) (arrow indicates where lymphatics are seen in ciliary body bounded by ciliary processes (CP) and sclera (S)). (B) Lymphatic channel in green (arrow) is identified with anti-podoplanin immunofluorescence. (C) Lymphatic channel in green (arrow) is distinct from collagen IV-positive blood vessel in blue (asterisk). Both are absent in the negative control without primary antibodies (D). Scale bar = 100 μm (A), and 5 μm (B,C,D).
42
Figure 3-7. Podoplanin-positive lymphatic channels in sclera. (A) Sclera (S) adjacent to the ciliary processes (CP) was identified with merging fluorescent and bright field microscopic images. (B) The structure was confirmed in the identical section stained with collagen IV (blue) counterstained with PI (red) (arrow indicates where lymphatics are seen in the sclera). (C) Lymphatic channel in green (arrow) is identified with anti-podoplanin immunofluorescence, distinct from collagen IV-positive blood vessel in blue (asterisk). Both signals are absent in the negative control without primary antibodies (D). Scale bar = 50 μm (A, B), and 10 μm (C, D).
43
Figure 3-8. Podoplanin-positive lymphatic channels in posterior orbit. (A) Posterior orbit section containing the retina (R), choroid (Ch), and Harderian Gland (H) stained with collagen IV (blue) and counterstained with PI (red) (arrow indicates where lymphatics are seen in the posterior orbit). Sclera was not visible in the staining. (B) Podoplanin-positive lymphatic channel in green (arrow) is distinct from collagen IV-positive blood vessel in blue (asterisk). Both signals are absent in the negative control without primary antibodies (C). Scale bar = 200 μm (A), and 10 μm (B,C).
44
The existence of lymphatic channels in the eye and orbit provided a framework for future
studies in determining the role of ocular and orbital lymphatics in the ocular lymphatic drainage
observed from in vivo hyperspectral imaging [159]. Most importantly, our data so far provided
evidence of a lymphatic drainage from the mouse eye. Next, we asked whether latanoprost, a
PGF2α analog, stimulates ocular lymphatic drainage. We first confirmed the IOP lowering effect
of latanoprost to ensure the drug has its action in our mouse strain. Latanoprost-treated left eyes
(n=5) showed a significant difference in mean IOP compared to control left eyes (n=5) at 2 hours
following last instillation (9.1±1.1 mmHg vs. 14.9±2.4 mmHg; (Mean±SD), P < 0.001, two-
sample t-test) (Figure 3-9). No significance difference was observed in other time points for left
eyes and in all time points for right eyes.
Figure 3-9. Effect of latanoprost on IOP. Latanoprost-treated left eyes (n=5, black squares) showed a significant difference in mean IOP compared to control left eyes treated with 0.02% benzalknonium chloride in 1xPBS (n=5, open circles) at 2 hours following last instillation. *P<0.001.
45
Next, we examined the effect of latanoprost on the ocular lymphatic drainage in mouse.
In the latanoprost-treated group, QD signal in the left neck region was detected in 2 mice within
20 minutes of QD eye injection (Figure 3-10A), and in 7 additional mice by 70 minutes (Figure
3-10C and Figure 3-11). Control mice treated with artificial tears showed no neck signal at 20
minutes (Figure 3-10B), 40 minutes, or 70 minutes (Figure 3-10D) after QD injection (Figure 3-
11). By 6 hours, all latanoprost-treated (Figure 3-10E) and control mice (Figure 3-10F) except
#36 showed neck signal (Figure 3-11). No QD signal was detected except at the injection site
using in vivo hyperspectral imaging prior to observation of QD signal in the left submandibular
lymph node. QD signal detection rate was increased in the latanoprost-treated group compared to
controls (1.23±1.06 hours-1 vs. 0.30±0.17 hours-1, mean±SD, P< 0.02) (Figure 3-12).
46
Figure 3-10. In vivo hyperspectral imaging of mouse neck region. Following QD injection into the left eye, latanoprost-treated mice showed QD signal (red) in the left neck region (arrow on ventral view) at 20 minutes (A, #20), 70 minutes (C, #43), and 6 hours (E, #39) by in vivo hyperspectral fluorescent images. In controls treated with artificial tears, QD signal was not detected at 20 min (B, #38) or 70 minutes (D, #42) post-injection imaging times, but was present at 6 hours (F, #34). Green signal corresponds to background and red to QD signal. Scale = 10 mm.
47
Figure 3-11. In vivo QD signal detection over post-injection imaging times. In most latanoprost-treated mice (black squares), QD signal was detected in the neck at earlier post-injection times compared to controls treated with artificial tears (open circles).
48
Figure 3-12. In vivo QD signal detection rate. Histogram shows mean and standard deviation of QD drainage rate (hours -1) for latanoprost-treated group (black, n=11) and control group treated with artificial tears (white, n=11) groups. *P<0.05.
49
Frozen sections were examined to ensure QDs were drained from the left anterior
chamber to the left submandibular lymph node in both groups. Indeed, QDs were detected inside,
and beneath the capsule of the left submandibular lymph node in both latanoprost-treated (Figure
3-13A) and control (Figure 3-13B) mice. Collagen IV immunofluorescence staining outlined the
capsule with characteristic bean-shaped morphology of a lymph node (Figure 3-13A,B) while
counterstaining with Sytox green demonstrated lymph node cellular architecture (Figure 3-
13A,B).
Figure 3-13. Localization of QD in lymph node of latanoprost-treated and control mice. QDs in red are located in the left submandibular lymph node in both latanoprost-treated mouse (A, #37) and control mouse treated with artificial tears (B, #23). Both lymph nodes are surrounded by capsule in blue (anti-collagen IV) against a green background of cell nuclei (Sytox green). Scale = 250 μm.
50
In addition to demonstrating an increased lymphatic drainage, we aimed to provide
evidence of an increased amount of QDs being drained to the submandibular lymph node. When
examining the QD intensity in the left submandibular node, mean total QD intensity was
significantly greater in latanoprost-treated mice compared with controls (10.55±1.12 vs.
9.48±1.24, mean±SD, P<0.05) (Figure 3-14). QD signal was not detected in the right
submandibular node in both groups. Overall, our findings from in vivo and post-mortem analysis
show that latanoprost stimulates the ocular lymphatic drainage from the mouse eye [168].
Figure 3-14. QD intensity in the submandibular node in latanoprost-treated group and control group treated with artificial tears. Box-plot shows the median, 25th, and 75th percentiles (solid line box), and the minimum and maximum intensity (whiskers) for the natural log-transformed total QD intensity gray value measured. *P<0.05.
51
4
Discussion
The present thesis provides, for the first time, evidence of lymphatic drainage from the
mouse eye [159]. A non-invasive approach combining intracameral injection of nanotracers and
hyperspectral fluorescence imaging was developed to visualize in vivo lymphatic flow from the
eye. We were first to identify lymphatic channels in the mouse ciliary body, sclera, and orbit that
may be responsible for this ocular lymphatic drainage. The establishment of lymphatic outflow
from the eye provided opportunities to investigate whether any drugs target this drainage.
Through the application of the novel in vivo lymphatic imaging technique, we demonstrated for
the first time, latanoprost, the most potent and widely prescribed glaucoma drug, stimulates
ocular lymphatic drainage [168]. Collectively, our findings support the use of this mouse model
in combination with the in vivo imaging technique for future screening studies of drugs that may
target the ocular lymphatic circulation.
The existence of lymphatic drainage from the mouse eye is in keeping with lymphatic
drainage reported in sheep [5, 6]. The submandibular lymph node [167] appears to be the initial
52
draining lymph node from the eye as strong QD signal was observed following intracameral
injection. The presence of QDs in other tissues at concentrations below the threshold of the
imaging system used cannot be excluded. Our study shows that QDs used to detect lymphatic
drainage from peritoneal and pleural spaces, and skin [169-171], are useful to study ocular
lymphatic drainage, a newly described route for fluid out of the eye. However, our results differ
from the literature regarding the location of the lymphatic drainage. We only observed the
intracameral injected QDs draining toward the submandibular lymph node, but not cervical or
facial lymph nodes [5, 6, 172]. It is possible that the retention of fluorescent dextrans or
radioactive albumins in lymph nodes is different from QDs. QDs are known to accumulate in the
first lymph node they were drained to and have excellent retention [173]. This may explain why
QDs were only observed in one lymph node. Tracers with various sizes, charges, and optical
properties [6, 52, 151, 174] can be used to examine whether or not locations of drainage via the
ocular lymphatics in mouse are affected by selection of tracers.
The distribution of QDs in the subcapsular sinus close to one pole of the submandibular
lymph node matches closely the afferent flow into the lymph node [175]. Our results are in
keeping with the distribution of the QDs across the subcapsular sinus of the sentinel lymph node
following subdermal injection [150]. It is possible that the size and charge of QDs may affect
their distribution pattern and this requires further study. It is known that mouse skin and fur can
reduce the QD detection sensitivity [176, 177]. This is supported by the stronger QD signal we
observed following skin removal in post-mortem imaging. For this reason, we cannot rule out the
possibility that some QDs were drained earlier than the initial detection time following tracer
injection. This reduction in QD655 sensitivity may not be optimal for quantitative analysis of in
vivo hyperspectral imaging [178].
53
The data from frozen left eye and orbit specimens revealed that following intracameral
injections, QDs were observed in the orbit and in the tissue compartments that typically comprise
the UVS outflow pathway - the ciliary body, choroid, and sclera. The tissue compartments from
the UVS pathway in which the fluorescent tracer appeared in our findings are the same as
observed in the NIH Swiss mouse eyes [128], supporting the use of QDs, like fluorescent dextran,
as tracers for time course study of AqH drainage. Our results, however, differ in the intensity of
fluorescence from QDs in tissue compartments. In the present thesis, an elimination of
fluorescence within the elements of UVS outflow pathway [128] was not observed at 6 hours
following intracameral injection. Instead, QDs fluorescence remained at moderate to high
intensity amongst the tissue compartments. A possible reason for this may be the injection
volume. Instead of 1.5 µL of tracer used, it is possible that 3 µL of QDs is sufficient for a
continuous transport of tracers through the UVS pathway; thereby, maintaining the observed
intensity in the frozen sections. Another possibility is that the size of the tracer may be important.
The ocular distribution of dextrans following intracameral injection varies with various sizes of
the tracers [52], 40-kDa dextran was shown to be optimally recovered in the anterior uvea,
anterior sclera, and posterior sclera for normal eyes; while 150-kDa dextran was recovered most
efficiently in inflamed eyes [52]. Therefore, additional experiments using tracers with various
sizes may further clarify the properties of macromolecular transport of AqH outflow to the
various ocular tissue compartments.
The identification of lymphatic channels in the mouse ciliary body using podoplanin, a
specific lymphatic endothelial cell marker supports the previous findings of lymphatics identified
in the human and sheep ciliary body using D2-40 to specifically detect podoplanin and LYVE-1
antibody for the lymphatic endothelial hyaluronan receptor [5]. The characteristics of the
54
lymphatic channels in mouse ciliary body are in keeping with lymphatics in human and sheep
ciliary body as they are not covered by collagen IV-positive basement membrane, distinguishing
them from blood vessels [179, 180]. Larger lymphatic collecting ducts covered by collagen IV-
positive basement membrane were not observed in our study. Further experiments using electron
microscopy may be helpful in locating larger collecting ducts in the orbit.
The presence of lymphatics in the sheep ciliary body provides a unique pathway for the
drainage of AqH [5, 6]. In sheep, tracers injected in the anterior chamber are drained to the head
and neck lymph nodes via the lymphatic in the ciliary body. Other studies have confirmed that
intraocular antigens can traffic to local draining lymph nodes [181, 182]. It is possible that the
injected tracer in the mouse eye draining to the submandibular lymph node [159] is partially due
to the lymphatics in the ciliary body. When taking into account of both identification of
lymphatics in ciliary body and the observation of QDs in the ocular compartments composed of
the UVS pathway, for instance, ciliary body, choroid, sclera, it is possible that QDs were drained
through the lymphatic channels in the mouse ciliary body. This network of lymphatics may
branch off to join with lymphatic channels in the orbital tissues in the posterior orbit. Altogether,
these lymphatic channels can drain QDs from the anterior chamber to the submandibular lymph
node. Further studies are needed to determine whether the lymphatics in the ciliary body and
orbit contribute to AqH drainage.
Previous studies using corrosion casting and serial section microscopy have indicated that
fluid can be drained from the suprachoroidal space through preformed scleral channels at the
inner aspect of the anterior sclera into the intrascleral venous plexus [183, 184]. Later studies
using 5'-nucleotidase, an early marker for LECs [185], failed to identify these scleral channels as
lymphatics [186]. In the present study, lymphatic channels were identified at the episclera with
55
podoplanin. These lymphatics are not covered by collagen IV-positive basement membrane,
distinguishing them from blood vessels [179, 180]. The presence of lymphatic channels in the
episclera may provide a unique pathway for the drainage of fluid out of the suprachoroidal space
into the sclera and finally into the orbital tissues [187]. Further studies are needed to determine
the exact location of the distributions of lymphatic channels in the sclera to better understand the
drainage of fluid out of the eye.
Mouse conjunctival lymphatics were identified recently with LYVE-1 and podoplanin
[188]. We confirmed podoplanin-positive conjunctival lymphatic channels in this study in which
they were also used as positive controls. Though LYVE-1 can also detect macrophages [188], it
was demonstrated that podoplanin staining was negative on the LYVE-1+ non-endothelial cells.
Since our study used podoplanin as the lymphatic endothelial cell marker, the possibility of the
luminal structures identified in this study being macrophages is unlikely.
In the orbit, lymphatics have previously been identified in the lacrimal gland and the dura
mater of the optic nerve in human [185, 189], and in the Harderian gland of rats [190].
Lymphatics in the extraocular muscles and in the connective tissues of the extraocular muscle
cones in mice [188] were not investigated in the present study due to problems with processing
and sectioning of the extraocular muscles and their connective tissues. Nevertheless, using
podoplanin as a lymphatic endothelial cell marker, we identified the existence of lymphatic
channels near the Harderian gland in mice. The presence of the lymphatic channels in the orbit
may provide a drainage pathway of the ocular/orbital tissue fluid. Since AqH drained via the
UVS pathway exits the portal in the sclera [62], it is also possible for these orbital lymphatic
vessels to drain fluid originating from the exit sites of the scleral portals. Further studies are
needed to determine the origin and destination of lymph drained via the orbital lymphatics.
56
The fact that the eye is frequently affected by inflammations [191], and intracamerally
injected antigens are drained to neck lymph nodes [159, 192, 193], suggest that ocular
lymphatics [5] identified in this thesis may play a role in immune responses. Indeed,
experimentally induced inflammation of ciliary body in cynomolgus monkeys increases the UVS
outflow [194]. Hence, our findings may be important for studying ocular inflammation. In terms
of intraocular tumors, metastasis can occur via spreading to regional lymph nodes [195, 196].
The risk of metastasis and mortality are higher when tumors invade the ciliary body [197, 198].
Podoplanin staining ciliary body of posterior uveal melanoma with and without extraocular
extension showed lymphangiogenesis in the ciliary body [199]. Our findings of an ocular
lymphatic drainage [159] and lymphatic channels in the normal ciliary body and sclera support
the notion of intraocular peritumoral lymphatics originating from preexisting lymphatics in the
ciliary body.
The existence of an ocular lymphatic drainage in mouse [159] led to opportunities to
study pharmacology of this drainage pathway. Here we provided the first evidence that a topical
PG, latanoprost, a widely prescribed glaucoma drug, stimulates lymphatic drainage from the eye
[168]. Quantification of the total QD intensity from all serial lymph node sections demonstrated
significant increase of QD signal in left submandibular lymph node in latanoprost-treated group
compared to controls. In addition, the fact that QD signal was not detected elsewhere in both
latanoprost-treated and control mice suggests that topical application of latanoprost increases the
drainage rate without altering the location of QD drainage. The increased lymphatic drainage rate
from the eye to the lymph node in the latanoprost-treated group provides evidence that
latanoprost stimulates lymphatic drainage from the eye. Cellular changes induced by latanoprost
may be implicated in this process. Ciliary muscle relaxation [200], by PG action on
57
Prostaglandin F (FP) receptors [200-203] may contribute indirectly to increased ocular lymphatic
drainage. PGs act on lymphatic endothelial and contractile cells surrounding lymphatic channels
responsible for the peristaltic movement of lymph [204]. Latanoprost remodels extracellular
matrix, increasing matrix metalloproteases (MMP), proMMP-1 and proMMP-3 [205], while
decreasing collagens, fibronectin, laminin, and hyaluronan [206, 207]. It is unknown whether
these cellular changes observed following long-term latanoprost treatment, are relevant to
increased lymphatic drainage observed in this study.
The increased detection rate of intracamerally injected QDs in the left submandibular
lymph node in the latanoprost-treated group compared to controls, suggests that latanoprost may
act directly on the lymphatic vessels within the eye [5]. In addition to this uveolymphatic
“pathway”, our findings from fluorescent and bright field microscopy on the QDs outflow
pathways in the left eye and orbit specimens confirm that a portion of tracers were drained by the
UVS pathway to sclera [128] and then to the orbit. It is possible that some of these tracers in the
orbit are drained via the orbital lymphatics [185, 188, 190] to the regional lymph node. Further
studies are needed to determine the role of orbital lymphatics in AqH drainage.
The hydrodynamic size and intrinsic brightness of QDs provide optimal retention in
lymphatic vessels and lymph nodes [151] in addition to emit signal that can penetrate up to 2 cm
of tissue [150] making this in vivo imaging method very sensitive to changes in lymphatic
drainage. The optical resolution of this hyperspectral imaging system, however, restricted in vivo
visualization of QDs in TM, UVS pathway, and iris. While localization of QDs in the UVS
pathway was demonstrated in coronal orbital sections, QDs in anterior chamber angle structures
such TM and SC that are involved in the conventional pathway could not be assessed, and the
use of transverse frozen sections may help to locate QDs in these structures. Without visualizing
58
QDs in the TM, we cannot exclude the possibility that some QDs were drained through the
conventional pathway into the venous circulation and be taken up by the liver and spleen [208].
However, no QD signal was observed in the liver or spleen from any mice in the latanoprost-
treated (n=11) or control group (n=11) during in vivo and post-mortem imaging performed 6
hours following intracameral injection. It is possible that concentrations of QDs in the spleen and
liver were below the detectable threshold by the imaging system used. Further experiments with
more sensitive techniques [208, 209] may be needed to investigate the QDs localization in the
liver and spleen following intracameral injection.
AqH dynamics have been studied in several species [210, 211] including the mouse
[155]. Our observation that an increase in lymphatic drainage rate following latanoprost
application matches temporally with a decrease in IOP suggests that enhanced ocular lymphatics
may be implicated in the IOP-lowering effect of latanoprost in normal mice [8, 155]. As mouse
aqueous dynamics are similar to humans [7], the presence of lymphatic drainage from the mouse
eye provides the framework for developing glaucoma treatments through pharmaceutical
manipulation of the lymphatics to decrease IOP. In addition, since lymphatic drainage from the
eye may play an important role in controlling pressure in the eye, further investigations in
glaucoma mouse models [212] will help to determine whether stimulating lymphatic drainage
from the eye using latanoprost, is implicated in its IOP lowering effect in glaucoma.
59
4.1 Limitation of the Studies
In demonstrating, in vivo, a lymphatic drainage from the mouse eye, our experiments
relied on both the hyperspectral imaging system and the nanotracer, QDs. We acknowledge that
both tools inherently include some limitations. The detection of QDs by the hyperspectral
imaging system are dictated by two factors. First, the intrinsic brightness of QDs can penetrate
the tissue of a depth up to 2cm [150]. Hence, our detection of QDs are restrained by the
proximity to the surface of the tissue structure in which QDs locate. Second, the intensity of QDs
penetrated through the tissue and skin has to reach a certain detection threshold. Taken together,
it is possible that QDs were drained into deep tissues from where QD signals cannot penetrate
toward the surface. Similarly, traces of QDs that are below the detection threshold could be
drained to other tissues aside from the submandibular lymph node. Therefore, we cannot exclude
the possibilities that some QDs may have been drained into structures other than the
submandibular lymph node within the 6 hour in vivo imaging time period. Further studies
involved sacrificing animals in earlier time points, sectioning whole body, and increasing the
exposure times can be helpful in determining whether QDs can be detected elsewhere other than
the submandibular lymph node.
The optical resolution of the hyperspectral imaging system is another limitation we faced
when we aimed to confirm the aqueous outflow of QDs within the eye during in vivo imaging.
The system is sensitive to the changes in QD intensity in gross anatomical structure such as
lymph nodes. However, the optical resolution of the system is unable to provide sufficient
magnification to visualize finer ocular structures. Hence, we were unable to confirm in vivo AqH
outflow through the TM and UVS pathway. Post-mortem studies with examination of frozen eye
60
and orbit sections will help understand and characterize the trajectory of QD within in the eye
and orbit following intracameral injection.
In the identification study of lymphatic channels, it is important to note that the value of
immunofluorescence relies heavily on the specificity of the primary antibody toward the
lymphatic endothelium. In our study, anti-podoplanin stained specifically the lymphatic channel
but not the blood vessel. We did not select tissues without lymphatics to eliminate the
possibilities of false positive results from anti-podoplanin staining for non-specific non-
endothelial cells. Nevertheless, we relied on the morphological criteria, such as having a full
central lumen and partially-collapsed structure, to determine whether podoplanin-positive
structures are indeed lymphatic channels. Further improvement of this study would be to use a
different primary antibody targeting lymphatic endothelium, for instance, anti-LYVE-1, and
compare results to further confirm the existence of lymphatic channels in the eyes and orbits.
Finally, in vivo quantification was restrained in studying the effect of latanoprost on the
lymphatic drainage. The intrinsic brightness of QDs led to oversaturation of signals in the eye
during in vivo imaging. Hence, QDs intensity changes in the eye could not be quantified and
compared between the latanoprost-treated and control mice. Further studies using tracers with
less intrinsic brightness and adjustment of imaging parameters will be helpful in determining the
changes of QDs intensity in the eye.
61
4.2 Future Directions
Role of Ocular Lymphatics in AqH Drainage
Ocular lymphatics identified in this thesis may play an important role in draining AqH;
thereby, controlling pressure in the eye. Hence, determining their role in AqH drainage is highly
relevant for glaucoma studies. It has been shown in sheep that intracamerally injected tracer are
located in the lumen of lymphatic channels in the ciliary body [5], which demonstrated
lymphatic channels playing a role in AqH drainage. It is highly likely that intracamerally injected
QDs in the mouse left eye will also be located in the lymphatic channels in the conjunctiva,
ciliary body, and sclera. Immunofluorescence targeting lymphatic channels in the frozen eye
sections with QDs followed by examination of these sections with confocal microscopy will be
needed to localize QDs in the lumen of lymphatic channels.
The localization of QDs in the posterior orbit raises the possibility of orbital lymphatics
involving in AqH drainage. The location to where tracers are drained by the orbital lymphatics is
currently unknown. QDs can be directly injected into the orbital tissue to determine the AqH
outflow pathways through there.
Mapping AqH Outflow Pathway to Submandibular Lymph Node
Understanding the general pathways in which QDs drained from the anterior chamber to
the submandibular lymph node is crucial. This can be achieved through intracamerally injecting
tracers conjugated with specific antibody against LECs such that the tracers can bind to the
lymphatics as they are being drained. It is possible that the lymphatic drainage begins in the
extracellular spaces of the ciliary muscle and enters into the suprachoroidal space and anterior
choroid. Then the network exits the periocular tissues directly through the sclera or through the
62
loose connective tissue surrounding penetrating vessels and nerves. QDs will then be drained
toward the tissues toward the submandibular lymph node.
Dissection of Ocular Lymphatic Drainage from UVS Outflow
Since PGF2α analogs affect both lymphatic vessels [156-158] and UVS drainage [105,
106], it may be difficult to separate the component of increased UVS pathway from the direct
effect on lymphatics. Future studies using other drugs such atropine [62], that have been shown
to increase the UVS pathway via a different mechanism from PGF2α analogs, or the use of
selective drugs that do not affect UVS outflow but have direct effect on lymphatics may be
helpful to pharmacologically dissect out ocular lymphatic drainage from UVS outflow. Better
characterization of lymphatics in the eye and in the orbit in mouse and the localization of PGF2α
receptors and other receptors in the lymphatics in the eye and orbit may help to identify new
therapeutic targets to modulate lymphatic drainage from the eye.
Studying Ocular Lymphatic Drainage in Glaucoma Mouse Models
Existing glaucoma mouse models provide opportunities to study whether ocular
lymphatics can be targeted to control IOP; thereby, delaying onset or progression of glaucoma.
In human, it is known that glucocorticosteroid administration may lead to the development of
ocular hypertension and open-angle glaucoma [213-216]. It has been demonstrated that
application of dexamethasone, a glucocorticosteroid, to the mouse eye will lead to ocular
hypertension [217]. The mechanism behind the increased IOP was thought to be morphologic
and biochemical changes in the TM, which led to reduction in AqH outflow [218]. Similarly,
transgenic mice, such as a mouse strain expressing the Tyr423His myocilin point mutation can
also be used [219, 220]. Myocilin is a secreted glycoprotein [221, 222] that is expressed in high
63
amounts in the TM [223-225]. In human, the secreted myocilin is present in the aqueous humor
[226-228]. Mutated myocilin was demonstrated to accumulate in the TM cell cytoplasm and was
prevented from secreting into the extracellular space in mouse [219], which leads to detachment
of the endothelial cells of TM and contributes to development of glaucoma [219]. These
glaucoma models are especially useful for studying the effect of ocular lymphatics in controlling
IOP as only the TM was predominately affected while the UVS outflow remains unaffected.
These studies would allow the screening various compounds that stimulate the ocular lymphatic
drainage and lower IOP in the glaucomatous mice .
Immune Response, Tumor Metastasis, and the Eye
The findings of an ocular lymphatic drainage provided opportunities to study ocular
immune responses and metastasis. It has been demonstrated that experimentally induced
inflammation of ciliary body in cynomolgus monkeys increased UVS outflow [194]. Studies
involving inducing ocular inflammation in mouse and assessing lymphatic drainage with in vivo
hyperspectral imaging can be helpful in understanding the relationship between ocular
inflammation and the lymphatic drainage. Intraocular tumors can metastasize via spreading to
regional lymph nodes [195, 196]. Lymphatics in the ciliary body may play a role in metastasis
of uveal melanoma [199, 229]. Studies involving stimulating lymphangiogenesis or ocular
lymphatic drainage followed by observing the extraocular extension and metastasis would be
helpful in understanding the role of ocular lymphatics play in spread of the tumour [230]. The
results may enable development of anti-angiogenic drugs or drugs that slow ocular lymphatic
drainage to prevent metastasis of intraocular tumors.
64
4.3 Conclusion
This work provides direct evidence of lymphatic drainage from the mouse eye.
Lymphatic drainage from the eye has for the first time been visualized in vivo with QDs and
hyperspectral fluorescence imaging techniques. Lymphatic channels identified in the ciliary body,
sclera, and orbit in mice may play a role in AqH drainage. Using this model in conjunction with
in vivo imaging, a stimulatory effect of latanoprost on ocular lymphatic drainage was shown.
The findings in this thesis support the use of this mouse model as a potential platform for future
screening studies of drugs that target the ocular lymphatic circulation. Due to limited sensitivity
to detect QD signal in deep tissue by in vivo hyperspectral imaging system, it may be difficult to
localize QD signal in deep anatomical structures. Future development of biocompatible tracers
and more sensitive hyperspectral imaging methods may nevertheless circumvent these
drawbacks and will enable translation of pre-clinical findings to clinical trials devoted to the
treatment of blinding eye disease.
65
5
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