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].

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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].

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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

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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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