Extracellular fluid systems in the brain and the ... · Extracellular fluid systems in the brain...

Extracellular fluid systems in the brain and the pathogenesis of hydrocephalus

by

Gurjit Nagra

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Graduate Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Gurjit Nagra 2010

ii

Extracellular fluid systems in the brain and the pathogenesis of hydrocephalus

A thesis submitted in conformity with the requirements for the degree of

Doctor of Philosophy Graduate Department of Laboratory Medicine and Pathobiology

University of Toronto, 2010 Gurjit Nagra

ABSTRACT

Fundamental questions related to the locations of Cerebrospinal Spinal Fluid (CSF)

absorption deficit and causes of the pressure gradients that expand the ventricles with

hydrocephalus remain largely unanswered.

Work in the Johnston lab over a 15 year period has demonstrated that CSF moves through the

cribriform plate foramina in association with the olfactory nerves and is absorbed by a

network of lymphatic vessels located within the olfactory turbinates. A kaolin-based rat

model of communicating hydrocephalus was developed as a collaborative effort with Drs.

McAllister, Wagshul and Li. After developing a method to quantify lymphatic CSF uptake

in rats, we examined and observed that the movement of a radioactive tracer into the nasal

turbinates was significantly reduced in the kaolin-injected animals compared to saline

injected controls. However, it was possible that while lymphatic CSF uptake was

compromised, other CSF absorption pathways may have compensated. To answer this, we

measured the CSF outflow resistance (Rout) and observed it to be significantly greater in the

kaolin group compared with animals receiving saline and there was a significant positive

correlation between CSF Rout and ventricular volume. Nonetheless, it is not clear how

impaired CSF clearance could lead to a dilation of the ventricles since the ventricular and

iii

subarachnoid compartments are in communication with one another and pressure would

likely increase equally in both.

At this point, we came across a theoretical paper that postulated that a drop in

periventricular interstitial fluid pressure might provide an intraparenchymal pressure gradient

favouring ventricular expansion. In addition, studies in non-CNS tissues indicated that a

disruption of beta-1 (β1) integrin-matrix interactions could lower tissue pressure. Based on

these suppositions and data, we examined if these concepts had relevance to the brain. For

this, we measured pressure in the brain and observed a decline in periventricular pressures to

values significantly below those monitored in the ventricular system following the injection

of the anti integrin antibodies. Many of the animals developed hydrocephalus over 2 weeks

post antibody injection. These data provide a novel mechanism for the generation of

intraparenchymal pressure gradients that is likely contributing to ventricular expansion.

iv

DEDICATION

From: Sukmani Sahib, Siri Guru Granth Shib Ji

I Thank my Dearest Grandfather, Gurdev Singh for teaching me so many things from a very young

age. However, the word “Thank” seems very small to capture the gratitude I feel towards a person

who was nothing but great and will always be in my heart....

I Thank my parents, Surinder & Harbhajan

My Mom: Pure Unconditional Love, Self-Sacrificing, Always Giving, My Very First

Loving Teacher, Always Believing in Me...

My Dad: Best Teacher, Example of Trust, Truth, Simplicity, and Belief...

My Siblings: My Dearest Brother Dara, Dearest Sisters Sarbjit, Rajwinder, Sukhwinder, Baljinder

and my “new” sister Lisa Wells .

My brother Dara, is the world’s greatest brother.... I could not possibly ask for more... He has

always been there for me...supported me... I thank him dearly...

I thank Sarbjit, Rajwinder, Sukhwinder, Baljinder, and Lisa for always loving me unconditionally....

I thank my sister in law Avnit, dearly along with my brother in laws, Jinder, Kashmir, Kulwant. I thank my nephews & nieces for giving my heart joy: Tejbir, Parmbir, Kiranjit, Satbir, Parbjot,

Amarjot, Amarvir, Rasjovan, Gurmeet, Hermeet, Manpreet.

v

ACKNOWLEDGEMENTS

My graduate studies experience in Johnston Lab has been very much enjoyable. I feel that

the statement “Time flies when you are having fun” is true especially because I am so

fortunate to have a masterful supervisor, Dr. Miles Johnston. It is my uttermost pleasure to

work under his supervision. Under his mentorship, I learned many valuable lessons some of

which include, the importance of preparedness, value of respecting time, greatest ideas

formulating from informal discussions, and the art of responding to critique. I am very

thankful to be a student of someone who is ‘Triple M’ - marvellously Mature, magnificently

Masterful, and genuinely Miles.

My past and present colleagues Ms. Lena Koh, Mrs. Dianna Armstrong, Ms. Amy

Baker, Ms. Sara Moore, Dr. Harold Kim, and Ms. Laura Kim of the lab along with my ‘R-

wing’ family Ms. Carrie Purcell, Mr. Alberto Jassir, and Mrs. Lisa Wells who have made my

graduate experience more enjoyable with their support and friendship.

I thank my advisory committee comprised of Dr. Isabelle Aubert, and Dr. James

Drake for their valuable feedback on the progression of my work. Additionally, I want to

express my gratitude to Dr. Aubert for ‘adopting’ me in her lab. I especially thank Ms. Kelly

Markham Coultes, Ms. Lillian Weng for helping me learn new molecular techniques. Finally,

I would like to thank Ms. Jessica Jordao for her support.

vi

TABLE OF CONTENTS

Abstract ii Dedication iv Acknowledgments v Table of Contents vi List of Tables xvii List of Figures xviii List of Abbreviations xx Statement xxii

vii

CHAPTER 1: GENERAL INTRODUCTION 1 1.1 Overview 2

1.2 Hydrocephalus 2

1.2.1 Prevalence of Hydrocephalus 2

1.2.2 Clinical Treatment Problems 3

1.2.3 Defining and Classifying the term “Hydrocephalus” 4 1.3 Proposed Theories of Hydrocephalus Induction 6

1.3.1 A lymphatic CSF absorption deficit as a cause of hydrocephalus 7

1.3.2 A novel mechanism to explain the development of pressure gradients in

hydrocephalus 8

viii

PART A CHAPTER 2: INTRODUCTION TO CSF-LYMPHATIC ABSORPTION 9 2.1 Cerbrospinal Fluid (CSF) 10

2.1.1 CSF Flow 12

2.1.2 CSF Function 12 2.2 Barriers of the Brain: CSF-Blood and Blood Brain Barrier 12 2.3 A CSF Absorption Defect: Classical Way of Thinking about Hydrocephalus 13

2.3.1 CSF Absorption into Lymphatic Vessels Vs Arachnoid Projections 14

2.4 Objectives of the Experimental Studies Outlined in Part A of this Thesis 15

ix

CHAPTER 3: DEVELOPMENT OF MODEL TO ASSESS LYMPHATIC CEREBROSPINAL FLUID ABSORPTION DEFICIT 17

3.1 Abstract 18

3.2 Introduction 18

3.3 Material and Methods 21

3.3.1 Animals 21

3.3.2 Tracer Infusion and Tissue Sampling 21

3.3.3 Sectioning Olfactory Turbinate Sample and Radioactivity Assessment 22

3.3.4 Visualization of Transport Across the Cribriform Plate 23

3.3.5 Analysis of Data 24

3.4 Results 25

3.4.1 Visualization of CSF Transport Across the Cribriform Plate 25

3.4.2 Comparison of Tracer Concentrations in Various Tissues Over Time 28 3.5 Conclusion 32

x

CHAPTER 4: DOES A CSF LYMPHATIC ABSORPTION DEFICIT EXIST IN A KAOLIN MODEL OF HYDROCEPHALUS AND DOES THE LYMPHATIC DEFECT CORRELATE WITH HYDROCEPHALUS? 33

4.1 Abstract 34

4.2 Introduction 35


4.3.1 Animals 37

4.3.2 Induction of Communicating Hydrocephalus 37

4.3.3 Assessment of Ventricular Size 38

4.3.4 Assessment of Lymphatic CSF Absorption 39 4.3.5 Visualization of Transport Across the Cribriform Plate in Kaolin and Saline

Injected Animals 39


4.4 Results 41

4.4.1 Development of Hydrocephalus 41

4.4.2 Assessment of CSF Transport Through Cribriform Plate 45

4.4.3 Transport of CSF Tracer into Olfactory Turbinates: Comparison Between

Kaolin and Saline injected Animals 47

4.5 Conclusion 53

xi

CHAPTER 5: DOES A LYMPHATIC DEFICIT CORRELATE WITH A GLOBAL CSF ABSORPTION DEFICIT? 54

5.1 Abstract 55

5.2 Introduction 57


5.3.1 Animals 58 5.3.2 Details of Collaboration 58

5.3.3 Induction of Communicating Hydrocephalus 58

5.3.4 Assessment of Ventricular Size 59

5.3.5 Assessment of Lymphatic CSF Absorption 59

5.3.6 Outflow Resistance 59


5.4 Results 61

5.4.1 Development of Hydrocephalus 61

5.4.2 Transport of CSF Tracer into Olfactory Turbinates: Comparison

Between Kaolin and Saline injected Animals 61

5.4.3 CSF Outflow Resistance 66

5.5 Conclusion 69

xii

CHAPTER 6: LYMPHATIC CSF ABSORPTION AND HYDROCEPHALUS DISCUSSION 70

6.1 Discussion of Results 71

6.1.1 Quantifying Lymphatic CSF Absorption 71

6.1.1.1 Limitations of Quantitative Method 71

6.1.1.2 Physiological Characteristics of Tracer Movement Across the Cribriform

Plate 72

6.1.2 Lymphatic Absorption in Kaolin Model of Hydrocephalus 74

6.1.3 Outflow Resistance in Kaolin Model of Hydrocephalus 79

xiii

PART B

CHAPTER 7: ROLE OF CELL β1-INTEGRIN – MATRIX INTERACTIONS IN

GENERATING A TRANSPARENCHYMAL PRESSURE

GRADIENTS FAVOURING THE DEVELOPMENT OF

HYDROCEPHALUS 81

7.1 Abstract 82 7.2 Introduction 83

7.3 An alternative explanation for ventricular expansion 84

7.4 Objective of Part B in this Thesis 85


7.5.1 Animals 86

7.5.2 Surgical Procedures for Pressure Measurements 86

7.5.3 Measurement of Pressure using the Servonull System 87

7.5.4 Antibody Injection for Pressure Measurement 89

7.5.5 Chronic Experiments 89

7.5.6 Assessment of Function Blocking Status of the Anti-β1 Integrin IgG Antibody 92

7.5.7 Penetration of Immunoglobins into Parenchymal Tissues 92

7.5.8 Data Analysis 93

7.6 Results 94

7.6.1 Acute Experiments 94

7.6.1.1 Pressure Analysis 99

7.6.1.2 Parenchyamal Pressure Following Injection of Anti- β1 Integrin antibody or

their isotype controls (Figure 7-4 A) 99

7.6.1.3 Comparison of Ventricular and Parenchymal Pressure after the Injection of

xiv

antibodies to β1 Integrin (Figure 7-4 B) 99

7.6.1.4 Comparison of Ventricular Pressure after the Injection of Antibodies

to β1 Integrin or their Isotype Controls (Figure 7-4 C) 100

7.6.1.5 Comparison of Ventricular and Parenchymal Pressure after the Injection of

IgG/IgM Isotype Controls (Figure 7-4 D) 100

7.6.1.6 Pressure Gradients 103

7.6.2 Chronic Experiments 103

7.6.3 Assessing the Characteristics of IgG anti- β1 Integrin 107

7.6.4 Penetration of Immunoglobins into the Peri-Ventricular Parenchyma Following

Intraventricular administration 109

7.7 Conclusion 111

xv

CHPATER 8: THE BRAIN INTERSTITIUM AND HYDROCEPHALUS

DISCUSSION 112

8.1 Summary of Conclusions Contained in Part B of this thesis 113

8.2 Disscussion of Results 114

8.2.1 Integrin and Matrix Elements in the Brain 114

8.2.2 Antibody Issues 116

8.2.3 Dissociation of Ventricular and Parenchymal Interstitial Pressure 117

8.3 Pressure Gradients and Hydrocephalus 118

8.4 Perspective and Significance of cell-matrix concept 119

xvi

CHAPTER 9: GENERAL DISCUSSION 120

9.1 Summary of Conclusions Contained in this Thesis 121

9.2 The development of hydrocephalus: Can one reconcile the 'classical' CSF absorption

deficit concept with the notion that the brain interstitium may be the 'epi-centre' of dysfunction in ventricular expansion 122

9.3 Issues for Future Investigation 127 REFERENCE LIST 129

APPENDIX 146

xvii

LIST OF TABLES

Table 2-1 CSF secretion, volume, and turnover rate in rat and human

Table 7-1 Antibodies and Isotype controls used in the Integrin study

Table 7-2 Impact of antibody injections on ventricular size

xviii

LIST OF FIGURES

Figure 3-1 CSF transport into olfactory turbinates in the rat.

Figure 3-2 Average CSF tracer recoveries (percent injected dose/gm tissue) in

various tissues (including the turbinates) as a function of time.

Figure 3-3 Average CSF tracer recoveries (percent injected dose/gm tissue) in

various tissues as a function of time.

Figure 4-1 Impact of saline or kaolin injection on the ventricular volumes.

Figure 4-2 Average CSF tracer enrichment (percent injected dose/gm tissue) in the

olfactory turbinates and blood as a function of time.

Figure 4-3 Comparison of lymphatic CSF absorption in the saline - and kaolin

injected animals.

Figure 4-4 Comparison of lymphatic CSF absorption in the saline (n=9) and kaolin-

injected animals (n=10).

Figure 5-1 Example of hydrocephalus induced with administration of kaolin into the

basal cisterns. (A) saline injected, (B) kaolin injected

Figure 5-2 Ventricle size and lymphatic CSF uptake.

Figure 5-3 Average CSF outflow resistance in intact, saline or kaolin-injected

animals.

Figure 5-4 Relationship between CSF outflow resistance and ventricular volumes

with both parameters measured in the same animals (n=16).

Figure 7-1 Sections of a rat head illustrating the position of the micropipette tip used

to measure periventricular interstitial fluid pressures (asterix in circle).

Figure 7-2 Examples of periventricular interstitial fluid pressures. Times of injection

are illustrated with arrows.

xix

Figure 7-3 Examples of periventricular and intraventricular pressures. Times of

injection are illustrated with arrows.

Figure 7-4 Comparison of the averaged pressures following injection of antibodies or

isotype controls. The normalization of the data and the method for

averaging have been described in the Materials and Methods. Injections

occurred at time 0.

Figure 7-5 Coronal sections of rat brains for assessment of hydrocephalus 2 weeks

after antibody/isotype control injection into a lateral ventricle. A ruler in

1 mm increments is illustrated in each image.

Figure 7-6 Adhesion of U937 cells to VCAM-1.

Figure 7-7 Penetration of the antibodies into the periventricular parenchyma after

administration into a lateral ventricle.

Figure 9-1 Two-Hit hypothesis for CH

Figure 9-2 An example of ventricular and parenchymal pressure measured

simultaneously in a 2 weeks post kaolin injection animal with evens ratio

of 0.51.

Figure 9-3 Example of (A, B) parenchymal and (C) ventricular pressure measured

after (A, C) 100µl heparinised blood and (B) 100µl heparinised saline

injection.

xx

LIST OF ABBREVIATIONS

α2 Alpha 2

α4 Alpha 4

α2 β1 Alpha 2 beta 1

β1 Beta 1

BBB Blood brain barrier

BCB Blood cerebrospinal fluid barrier

CH Communicating hydrocephalus

CNS Central nervous system

CSF Cerebrospinal fluid

CP Choroid plexus

CT Computed tomography

ECF Extracellular fluid

EGF Epidermal Growth Factor

FM Foramina of Monro

125I-HSA Iodinated human serum albumin

ICP Intracranial pressure

LN Lymph node

LV Lymphatic vessels

MRI Magnetic resonance imaging

Non-CH Non communicating hydrocephalus

NPH Normal pressure hydrocephalus

OB Olfactory blub

PDGF Platelet derived growth factor

xxi

Rout Outflow Resistance

SAS Subarachnoid space

TGF-β Tumor growth factor-beta

VCAM-1 Vascular cell adhesion molecule-1

xxii

STATEMENT

Some of the physiological experiments were complex in nature and required a team effort.

The data analysis was performed with the help of Marko Katic and Alexander Kiss,

Department of Research Design and Biostatistics, Sunnybrook Health Sciences Centre. The

studies outlined in chapter 4 and 5 were possible due to the availability of a kaolin

communicating model of hydrocephalus developed with our collaborators Dr. Pat McAllister,

Dr. Mark Wagshul and Dr. Jie Li. The contribution of key individuals to the work in this

thesis is outlined below.

Mrs Dianna Armstrong, the “den mother” of the lab helped me with the experiments

and taught me various surgical procedures. Her exceptional organizational skills

made it possible to carry out difficult and challenging experiments.

Andrei Zakarov, previous member of our lab, a skilful neurosurgeon who taught me

the stereotactic technique used to quantitative CSF absorption in lymphatic vessels

outlined in chapter 3.

Ms. Lena Koh, previous master’s student in Johnston Lab set up the servonull -

micropressure measurement system that was used for the studies outlined in chapter

7. She has been instrumental in troubleshooting any technical issues with the system.

Ms. Sara Moore, our senior technician helped with the measurement of outflow

resistance experiments outlined in chapter 5. She is a key member of the Johnston

lab. She helps set up and organizes the experiments.

Ms. Kelly Markham Coultes and Ms. Lillian Weng helped me perform western blot

analysis, which is illustrated in Figure 7-7.

Dr. Myron Cybulsky and Mr. Jacob Rullo helped me conduct the cell adhesion assay,

illustrated in Figure 7-6.

xxiii

Ms. Iris Lui (summer student) helped me perform the pressure measurement

experiments with the injection of heparinised blood outlined in chapter 9, Figure 9-3.

CHAPTER 1:

GENERAL INTRODUCTION

1

1.1 Overview

Hydrocephalus is a complex disorder. However, it is generally understood that it involves an

abnormal amount of cerebrospinal fluid (CSF) in the brain accompanied by ventricular

expansion. Many hydrocephalus models have been produced but the biomechanical cause(s)

remain elusive, as many investigators have failed to measure pressure gradients suitable for

causing ventricular expansion. My project was to first, re-consider the classical view of this

disorder, a cerebrospinal fluid absorption (CSF) deficit from a “lymphatic” perspective and

secondly, investigate a new parenchymal-based view of ventromegaly. Collectively, these two

objectives might provide a new approach for studying and hopefully treating hydrocephalus in

the future.

1.2 Hydrocephalus

1.2.1 Prevalence of Hydrocephalus

Hydrocephalus afflicts people of all ages, and is especially common in children. The incidence

of infantile hydrocephalus is 0.5 per 1000 live births (Gupta et al., 2007). It has been reported

that there are about 38,200-39,900 hospital admissions each year and total hospital charges of

$1.4-2.0 billon (in 2003) for pediatric hydrocephalus in the U.S (Simon et al., 2008). In the

U.S., estimates of shunt placement range from 5500 (Bondurant & Jimenez, 1995) to 18000

(Patwardhan & Nanda, 2005) in children and adults annually. There is significant lifelong

morbidity associated with problematic shunt placement for the treatment of hydrocephalus along

with adverse social functioning issues (Gupta et al., 2007).

While it is commonly assumed that hydrocephalus is associated with children and the

relatively young, chronic hydrocephalus in adults accounts for more than half of the 80,000

diagnoses of hydrocephalus in the US/year (Bondurant & Jimenez, 1995). This incidence is

2

likely underestimated since about 5-10% of demented patients have Normal Pressure

Hydrocephalus (NPH) (Hakim et al., 2001;Vale & Miranda, 2002). NPH is diagnosed

principally among individuals over the age of 60 years and is characterized clinically by urinary

incontinence, gait disturbances and dementia. The 'normal pressure' moniker is somewhat

misleading as continuous CSF pressure measurements can reveal waves of increased pressure

(Fishman R, 1992). NPH may develop from congenital hydrocephalus or arise in adult life

secondary to some known insult to the CSF system such as subarachnoid hemorrhage,

meningitis or head trauma (Edwards et al., 2004). In some cases, there is no known cause for the

disorder (idiopathic).

1.2.2 Clinical Treatment Problems

In an era in which innovative molecular and gene therapeutic approaches are increasingly being

entrenched in the clinical repertoire, the treatment of hydrocephalus remains primitive by

comparison. Current treatments are only partially effective and there is an urgent need to

reassess the conceptual foundation on which our understanding of this disease is based.

The management of this disorder involves diversion of cerebrospinal fluid (CSF) with

lumboperitoneal or ventriculoperitoneal shunts or the endoscopic third ventriculostomy

technique, a procedure in which the base of the third ventricle is surgically opened to permit the

movement of CSF into the basal cisterns. The lumboperitoneal or ventriculoperitoneal shunts

involve insertion of the peritoneal portion of a shunt catheter through a mini-laparotomy surgery

(Sekula, Jr. et al., 2009). Apart from the neurological complication due to the placement of the

shunt in the brain, this procedure has its own complications which involve incisional abdominal

herniation and CSF pseudocyst formation (Agha et al., 1983;Gaskill & Marlin, 1989). While

many children exhibit symptomatic and functional improvement with shunts, tragically, the

3

significant number of debilitating neurological deficits does not make such treatment an optimal

one. Indeed, a half century of research in shunt design has produced little improvement in the

rate of shunt survival. The majority of patients who receive a shunt need at least one revision

within the first 2 year after its initial placement (Drake et al., 1998;Drake et al., 2000;Kestle et

al., 2000). Shunts are often complicated by mechanical failure (Browd et al., 2006;McGirt et

al., 2002) and infection (McGirt et al., 2003;Kulkarni et al., 2001) requiring further surgical

procedures.

1.2.3 Defining and classifying the term “Hydrocephalus”

Hydrocephalus is generally thought to be caused by a disturbance in the CSF flow due to either

a deficit in CSF absorption or obstruction in the CSF pathway when CSF secretion rate remains

unchanged (Lorenzo et al., 1970;Levine, 1999). In terms of defining different types of

hydrocephalus, it is usually understood as communicating or non-communicating type. Non-

communicating hydrocephalus (non-CH) relates to the situation in which there is an obstruction

in the CSF system and no communication between the ventricular and subarchnoid space (SAS).

Communicating hydrocephalus (CH) is observed when there is some obstruction in the SAS but

not in the ventricular system. This is usually attributed to a blockage in CSF absorption.

However, the term “Hydrocephalus” does not always follow this clear-cut terminology. For

example, aqueductal stenosis, the prototypical non-communicating type of hydrocephalus, has

been suggested to occur secondary to communicating hydrocephalus in some cases (Nugent et

al., 1979;Raimondi et al., 1976;Williams, 1973). Additionally, it has been argued that the

classification of hydrocephalus should be considered from various standpoints. For example,

acute versus chronic, progressive versus arrested, congenital versus acquired, simple versus

complicated, hypertensive versus hypotensive, intra-uterine versus extrauterine, mild versus

4

severe, primary versus secondary, high intra-cranial pressure versus normal pressure, tumoral

versus non-tumoral, and infantile versus adult (Mori, 1990). The adult (hydrocephalus)

category may include congenital type with late onset, tumoral origin, slit ventricle syndrome,

non-CH, CH or normal pressure hydrocephalus (NPH) (Chahlavi et al., 2001;Bergsneider et al.,

2008). NPH is generally thought to be communicating type of hydrocephalus (Hurley et al.,

1999;Jellinger, 1976;Pickard, 1982;Tisell et al., 2006;Pena et al., 2002). The intracranial

pressure (ICP) in NPH is thought be normal due to some compensatory mechanism (Edwards et

al., 2004). Additionally, it can be defined generally as compensated or arrested congenital

hydrocephalus (Whittle et al., 1985) and may be subdivided into secondary NPH or idiopathic

NPH (Zemack & Romner, 2002;Hebb & Cusimano, 2001). Secondary NPH may arise after

subarachnoid hemorrhage, head trauma, or meningitis. However, it is difficult to pinpoint the

etiology of ventricular dilatation observed with idiopathic NPH i.e. whether it is due to brain

atrophy known as ex-vaccuo hydrocephalus (Kitagaki et al., 1998;Bradley, Jr. et al., 1996) or

because there is some disturbance in CSF dynamics (Silverberg et al., 2003;Silverberg et al.,

2002).

Hence collectively, the term “Hydrocephalus” encompasses many facets of this

condition. Clinically, in order to be useful diagnostically and with the hopes of urging future

investigations, Mori points out that this term should be considered at three levels (Mori, 1990).

First at a pathophysiological level, secondly at etiological level and thirdly at a pathological

level. Additionally, a more recent paper by Rekate proposed a new definition of hydrocephalus

as “an active distension of the ventricular system of the brain resulting from inadequate passage

of CSF from its point of production within the cerebral ventricles to its point of absorption into

the systemic circulation.” His definition excludes hydrocephalus ex vaccuo in which the

ventricles expands due to brain atrophy (thus not an active process) and benign intracranial

5

hypertension in which the ventricles are not enlarged (Rekate, 2009;Rekate, 2008).

Additionally, Rekate proposed a new classification system of hydrocephalus based on the use of

contemporary tools such as computed tomography (CT) and magnetic resonance imaging

(MRI). This classification system categorizes all types of hydrocephalus into obstructive types

based on the site of obstruction within the ventricular system.

This classification system however, raises two important issues. First, Rekate considers

hydrocephalus as an absorption problem at the level of arachnoid granulations; this is classically

defined as the communicating type. In this regard, as addressed in detail Part A - Chapter 2 of

this thesis, he does not addresses or consider the role of lymphatic CSF absorption. Secondly,

we will demonstrate in the studies outlined in this thesis, that hydrocephalus may occur without

any obstruction to CSF flow or absorption. We will develop the concept that a reduction in

parenchymal tissue pressure relative to that in the ventricles can cause pressure gradients

favourable to ventricular expansion. In any event, my thesis will address two critical issues of

CH. First, whether there is deficit in lymphatic CSF absorption and secondly, what role the brain

parenchymal interstitium may play in generating a pressure gradient favourable to

ventriculomegaly.

1.3 Proposed Theories of Hydrocephalus Induction

Since overproduction of CSF is relatively rare, most investigators have focused on CSF

malabsorption as the most likely cause of hydrocephalus. It is hard to ignore this possibility

since CSF outflow resistance is elevated in many human (Kosteljanetz, 1986) or experimental

cases (Mayfrank et al., 1997). Nonetheless, as will be discussed in more detail later, there are

theoretical objections to this concept and some investigators have looked elsewhere for

causative agents or events. Asymmetrical CSF pulsatility is one proposed mechanism (Egnor et

6

al., 2002) but some of the mathematical principles on which this idea is based have been

disputed (Tenti et al., 2002). Although, cilia dysfunction is believed to correlate with

hydrocephalus, a clear linkage between ventricular expansion and ciliary dynamics has yet to be

established (Daniel et al., 1995;Baas et al., 2006;Banizs et al., 2005). There are of course

several cytokines that have been implicated in causing hydrocephalus including TGF-β

(Galbreath et al., 1995;Kanaji et al., 1997) and FGF (Johanson et al., 1999;Johanson & Jones,

2001) as examples. Additionally, there are a myriad of mutations in various animal models that

give rise to a hydrocephalus phenotype but these are confusing, as in many cases, significant

other pathologies co-exist with the hydrocephalus phenotype (Philpot et al., 1999;Zhang et al.,

2006;Sunada et al., 1995).

1.3.1 A lymphatic CSF absorption deficit as a cause of hydrocephalus

The Johnston lab has developed the concept that a major pathway through which CSF is

absorbed from the subarachnoid space is via lymphatic vessels and not the arachnoid villi and

granulations. In this regard, there are no lymphatic vessels within the parenchyma of the brain.

CSF moves through the cribriform plate foramina in association with the olfactory nerves and is

taken up by an extensive network of lymphatic vessels located within the olfactory submucosa.

Once CSF has entered the absorbing lymphatics, it is conveyed in progressively larger ducts

through various lymph nodes and is deposited ultimately in the venous system at the base of the

neck (Zakharov et al., 2003). This concept is supported by studies in many mammalian species

including humans (Weller et al., 1992a;Lowhagen et al., 1994;Caversaccio et al., 1996) and

non-human primates (Yoffey & Drinker, 1939;Johnston et al., 2005). Overall, these experiments

demonstrated that lymphatics have the major function in removing CSF from the cranium

(Boulton et al., 1997;Boulton et al., 1998;Mollanji et al., 2001;Papaiconomou et al., 2002).

7

With this background, in Part A of this thesis we developed a model to begin to assess the role

of lymphatic function in hydrocephalus development.

1.3.2 A novel mechanism to explain the development of pressure gradients in

hydrocephalus

A study of inflammation in the skin has given rise to a fascinating concept related to matrix

function. The induction of skin inflammation results in a lowering of interstitial fluid pressure.

Remarkably, the reduction in tissue fluid pressure can range from a few mmHg to greater than

100 mmHg depending on the stimulus. This causes enhanced movement of water and solutes

into the tissue spaces by increasing the hydrostatic pressure gradient that favours capillary fluid

filtration (Reed & Rodt, 1991). In the studies conducted thus far, inflammation from a variety

of causes and several inflammatory mediators seem to induce this effect consistently but what is

most relevant to my studies, is the fact that a lowering of interstitial fluid pressure can be

produced by the injection of antibodies to the α2β1 or β1 integrins which appear to be the central

players in regulating this phenomenon (Reed et al., 1992). The critical factor here is that the

interstitial matrix is in a dynamic state and that disrupting the matrix integrity can be used to

modify tissue pressure. If the same were true of the brain parenchyma, we may be able to

develop a new view of hydrocephalus that incorporates perturbations in β1 integrin function as

one of the defining parameter that induces ventriculomegaly. In part B of this thesis, our recent

experience with a micropipette servo-null pressure measuring system encouraged us to

investigate whether changes in the parenchyma of the brain played an active role in inducing

ventriculomegaly.

8

PART A

CHAPTER 2:

INTRODUCTION TO CSF-LYMPHATIC ABSORPTION

9

2.1 Cerebrospinal Fluid (CSF)

From an historical perspective, CSF and hydrocephalus have been inseparable. In the 4th

Century BC, Hippocrates thought that this fluid was the pathological liquid that gets formed in

the brain with hydrocephalus (WOOLLAM, 1957). Interestingly, in the 3rd Century BC this

fluid was confused with “pneumas”, the animal spirits responsible for giving the whole body

energy and motion (Torack, 1982).

A small percentage, ~10-20% of CSF is produced by the capillary endothelial cells in the

brain parenchyma (Segal, 2001). It is also known as the brain interstitial fluid (ISF) or extra-

cellular fluid (ECF). However, the majority of CSF is produced by the choroid plexuses (CP)

(Table 2-1). CSF production is due to mainly three characteristics of the choroid plexuses. First,

the choroidal cells have multiple microvilli on the apical side (facing the ventricular surface)

along with a considerable amount of infolding at the basolateral (blood) side. These properties

increase the surface area of the CP. Second, the capillaries of the plexus are fenestrated

compared to the cerebral capillaries. This structural characteristic provides little resistance to

the movement of small molecules (Segal, 1993). Third, the CP has a very rich blood supply.

For example, the CP in rat lateral ventricles receives 3-4 ml/min/g which is 10 times the blood

supply to the cerebral cortex (Szmydynger-Chodobska et al., 1994). In terms of CSF volume, in

human, there is 164.5± 47 to 270±5 ml of this fluid, ~25% of which is in the ventricles and the

rest is extraventricular (Wright, 1978): in the subarachnoid space (SAS), in the basal cisterns of

the brain, and around the spinal cord (Segal, 2001). With regards to the flow of ISF, Cseff and

collogues have shown that there is a slow bulk flow of ISF towards the ventricular CSF (rather

than a simple diffusion) (Cserr et al., 1981). Ghersi-Egea and collogues showed a peri-vascular

pathway for ISF into (ventricular) CSF (Ghersi-Egea et al., 1996b;Ghersi-Egea et al., 1996a).

10

Table 2-1: CSF secretion, volume, and turnover rate in rat and human

Species Rat Human 3 months 30 months Young Older CSF secretion rate 1.21±0.27 0.65±0.16 410±240 190±70 (µl/min) CSF volume 155.5±12.2 307.8±57.7 165,000 - 270,000 (µl) CSF turnover 2.1 7.9 8.5 – 18.4 (Hrs to replace Total volume) Rat parameters - taken from Preston et al., 2001 Human parameter, CSF secretion rate - taken from May et al., 1990 Human CSF volume - taken from Wright et al., 1978 Human CSF turnover - calculated using CSF secretion rate taken from May et al., 1990 and averaged CSF volume taken from Wright et al., 1978

CSF is actively secreted by the choroid plexus. It is not a mere ultra-filtrate of blood plasma as

it was once thought to be because of its very low protein concentration (Ghersi-Egea et al.,

1996b;DAVSON & Welch, 1971;Kandel ER et al., 2000). The driving force of CSF secretion

is the unidirectional flux of ions due to the polarity of the CP epithelium (Brown et al., 2004).

In this regard, for the flux of ions, it has been shown that Na+-K+ ATPase, K+ channels, and Na+-

2Cl--K+ co-transporters are expressed in the apical membrane and Cl--HCO3 exchanger, a

variety of Na+ coupled HCO3- transporters and K+-Cl- co-transporters are expressed on the

basolateral side. The movement of ions through these various channels, transporters and co-

transporters creates an osmotic gradient allowing the movement of water cross the apical side of

choroid epithelium which is mediated by aquaporin-1 (Brown et al., 2004).

11

2.1.1 CSF Flow

It is important to briefly note the anatomical nomenclature of the brain ventricular system with

respect to CSF flow. CSF flows from the lateral ventricles through the foramina of Munro into

the third ventricle. It then passes down the aqueduct of Sylvius into the fourth ventricle.

Finally, it exits the ventricular system through the Foramina of Luschka and Magendie into the

subarachnoid spaces where it fills the spinal canal and the basal cisterns of the brain (Segal,

2001). From here, traditionally, it is believed that CSF is absorbed by arachnoid granulations

and villi into the superior sigittal sinus of the brain. However, the work of Dr. M Johnston has

demonstrated with qualitative and quantitative experimental data that the lymphatic vessels

located outside the brain play a primary role in CSF absorption in various animal models (they

used to address the concept).

2.1.2 CSF Function

The obvious function of CSF is to provide buoyancy and protection to the brain. It effectively

reduces the weight of the brain from ~1500g to ~50g (Davson H et al., 1987). The CSF also

acts to provide a “sink” to the brain extracellular fluid (ECF) (DAVSON et al., 1962). In this

regard, Cserr demonstrated that the ECF carries metabolized products and various peptides from

the brain into the CSF by bulk flow (Cserr et al., 1981).

2.2 Barriers of the Brain: CSF-Blood and Blood Brain Barrier

The blood-cerebrospinal fluid barrier (BCB) is formed by the epithelium of the choroid plexuses

and the arachnoid membrane, which are joined by tight junctions. At the level of choroid plexus,

the microstructure of the BCB is composed of a monolayer of polarized epithelial cells. The

12

apical side has abundant microvilli and the basolateral infoldings expand the surface area for

molecular exchange of various ions (Strazielle & Ghersi-Egea, 2000). It has been estimated that

in rats the total surface area of the choroid plexus is ~75cm2 (Keep & Jones, 1990;Zheng et al.,

2003).

The other barrier, the blood-brain barrier (BBB) is formed by a single layer of brain

capillary endothelial cells joined by tight junctions (Davson H & Segal M B, 1996). In the rat,

the surface area of the BBB is ~155 cm2 (Zheng et al., 2003). Structurally, the BBB is closely

associated with microglial, pericytes, and astrocytes in the external surface (Abbott,

2005;Pardridge et al., 1986). The main function of the BBB is to maintain the neuronal

microenvironment of the brain by being selective in transporting essential components across

the barrier. To restrict the free movement of solutes, it has been shown that the brain

endothelial cells have very low endocytic and transcytotic activity compared to the other tissue

endothelial cells (Vorbrodt & Dobrogowska, 2003).

In terms of efficiency of these barriers, the BBB is highly efficient compared to the

BCB. The BCB is functionally leaky (Zheng et al., 2003) because the capillaries of the plexus

are fenestrated (in contrast to the cerebral capillaries) (Redzic & Segal, 2004). These barrier

properties are of great interest in the pharmacology field.

2.3 A CSF Absorption Defect: Classical Way of Thinking about Hydrocephalus

The conventional view of hydrocephalus is founded on the assumption that ventriculomegaly is

essentially a 'plumbing' problem with the major defect associated with an impediment to

cerebrospinal fluid (CSF) absorption.

13

2.3.1 CSF Absorption into Lymphatic Vessels Vs Arachnoid Projections

In communicating hydrocephalus, the location of the transport deficit has never been established

but, since bulk CSF absorption is believed to occur through the arachnoid villi and granulations,

it has always been assumed that the impediment to CSF transport occurs at these structures or

somewhere within the extraventricular CSF system (hence the use of the term extraventricular

obstructive hydrocephalus by some investigators) (Redzic & Segal, 2004). However, the

function of the arachnoid projections is becoming increasingly unclear (Egnor et al.,

2002;Papaiconomou et al., 2002;Papaiconomou et al., 2004;Zakharov et al., 2004a).

While there is some in vitro data using isolated portions of dura that suggest a function for

arachnoid projections (Welch & Friedman, 1960;Welch & POLLAY, 1961), quantitative

experiments in cats, rabbits, monkeys (McComb et al., 1982;McComb JG & Hyman S,

1984;McComb & Hyman, 1990) and sheep (Papaiconomou et al., 2004;Zakharov et al.,

2004b;Zakharov et al., 2004a) indicate that very little CSF is transported into the cranial venous

system at normal CSF pressures. The available data suggests that CSF transport can occur into

the cranial venous system but only at high intracranial pressures (Papaiconomou et al.,

2004;Zakharov et al., 2004b). This suggests that a substantial portion of CSF absorption occurs

elsewhere and in this regard, the extracranial lymphatic system has received attention in recent

years (Koh et al., 2005).

The most important connections between the subarachnoid compartment and extracranial

lymphatics appear to exist at the level of the cribriform plate and the olfactory turbinates. The

specialized cells of olfaction (olfactory epithelium) are carried on turbinal bodies arising from

the ethmoid bone. The turbinates are supported by bone and are covered with a mucous

membrane originating at the cribriform plate. The olfactory turbinates contain an extensive

lymphatic network which is in communication with the subarachnoid space through specialized

14

connections with the olfactory nerves (Kida et al., 1993;Zakharov et al., 2003;Zakharov et al.,

2004b;Zakharov et al., 2004a).

Considerable evidence exists to support the notion that lymphatic vessels external to the

cranium play an important role in this process (Koh et al., 2005). While there are no lymphatic

vessels within the parenchyma of the brain, CSF moves through the cribriform plate foramina in

association with the olfactory nerves and is taken up by an extensive network of lymphatic

vessels located within the olfactory submucosa. Once CSF has entered the absorbing

lymphatics, it is conveyed in progressively larger ducts through various lymph nodes and is

deposited ultimately in the venous system at the base of the neck (Zakharov et al., 2003). This

concept is supported by studies in many mammalian species (Koh et al., 2005) including

humans (Weller et al., 1992b;Lowhagen et al., 1994;Caversaccio et al., 1996;Johnston et al.,

2004) and non-human primates (Yoffey & Drinker, 1939;McComb & Hyman, 1990;Brinker et

al., 1994;Johnston et al., 2005).

2.4 Objectives of the Experimental Studies Outlined in Part A of this Thesis

The main objective of Part A of this thesis is to examine the concept of a CSF absorption deficit

as a causative factor in ventricular expansion. For this, we first developed a quantitative method

to assess CSF lymphatic absorption and asked the following questions:

1. We hypothesize that there is a lymphatic CSF absorption deficit in the kaolin model (of

hydrocephalus) and we asked whether this impairment to lymphatic uptake of CSF

correlates with hydrocephalus?

2. Secondly, we hypothesize that the lymphatic CSF absorption deficit would affect the

global CSF absorption. For this, we measured CSF outflow resistance using the kaolin

15

model of hydrocephalus and asked whether a defect in lymphatic CSF absorption

correlates with a decline in global CSF absorption?

16

CHAPTER 3:

DEVELOPMENT OF MODEL TO ASSESS LYMPHATIC

CEREBROSPINAL FLUID ABSORPTION DEFICIT

17

3.1 Abstract

A major pathway by which cerebrospinal fluid (CSF) is removed from the cranium is transport

through the cribriform plate in association with the olfactory nerves. CSF is then absorbed into

lymphatics located in the submucosa of the olfactory epithelium (olfactory turbinates). In an

attempt to provide a quantitative measure of this transport, 125I-human serum albumin (HSA)

was injected into the lateral ventricles of adult Fisher 344 rats. The animals were sacrificed at

10, 20, 30, 40 and 60 minutes after injection and tissue samples including blood (from heart

puncture), skeletal muscle, spleen, liver, kidney and tail were excised for radioactive

assessment. The remains were frozen. To sample the olfactory turbinates, angled coronal tissue

sections anterior to the cribriform plate were prepared from the frozen heads. The average

concentration of 125I-HSA was higher in the middle olfactory turbinates than in any other tissue

with peak concentrations achieved 30 minutes after injection. At this point, the recoveries of

injected tracer (percent injected dose/gm tissue) were 9.4% middle turbinates, 1.6% blood,

0.04% skeletal muscle, 0.2% spleen, 0.3% liver, 0.3% kidney and 0.09% tail. The current belief

that arachnoid projections are responsible for CSF drainage fails to explain some important

issues related to the pathogenesis of CSF disorders. The rapid movement of the CSF tracer into

the olfactory turbinates further supports a role for lymphatics in CSF absorption and provides

the basis of a method to investigate the novel concept that diseases associated with the CSF

system may involve impaired lymphatic CSF transport.

3.2 Introduction

The true quantitative significance of any pathway in CSF transport can only be determined using

some forms of mass transport analysis, which involves quantifying the total mass of CSF tracer

over time. This has been the subject of previous studies from our group. In rats, we compared

18

the mass transport of radioactive HSA to plasma before and after obstruction of the downstream

cervical lymphatic vessels (Boulton et al., 1999). Plasma recoveries were reduced by

approximately 50% following the interruption of cervical lymph transport. Similar results were

observed in sheep (Boulton et al., 1997;Boulton et al., 1998). Additionally, support for

cribriform-lymphatic CSF absorption was obtained by combining infusion approaches with a

method to seal the cribriform plate extracranially (Mollanji et al., 2001;Papaiconomou et al.,

2002). Combined, these experiments demonstrated a significant role for lymphatic vessels in

cranial CSF transport. Indeed, at the theoretical opening pressure at which CSF absorption

would be initiated, the data suggested that over 80% of cranial drainage occurred through the

cribriform plate (Mollanji et al., 2001;Papaiconomou et al., 2002). We also assessed the direct

entry of a protein tracer into the cranial venous system in sheep and observed that tracer entry

into the superior sagittal sinus only occurred at high intracranial pressures (Papaiconomou et al.,

2004;Zakharov et al., 2004b). One possibility is that the arachnoid projections function to

divert CSF from the cranium when ICP is transiently or chronically elevated.

Most CSF studies in the Johnston lab have been performed in sheep (Boulton et al.,

1997;Boulton et al., 1998;Mollanji et al., 2001;Papaiconomou et al., 2002) and this could have

continued to provide important information on CSF parameters. However, studies in rodents

have several advantages. Apart from cost considerations, CSF dynamics in rats have been

studied extensively and (Mann et al., 1979;Cserr et al., 1981;Jones et al., 1987b;Ghersi-Egea et

al., 1996a;Johanson et al., 1999) a strong connection between CSF and extracranial lymph has

already been made in this species (Szentistvanyi et al., 1984;Kida et al., 1993;Johnston et al.,

2004;Koh et al., 2006). Additionally, in recent studies we used a rat model and found

quantitative data to support a lymphatic role in CSF absorption. We observed the impact of

elevated intracranial pressure being reflected in pressures measured in the deep cervical lymph

19

nodes; a study in which G. Nagra participated (Koh et al., 2007). Additionally, qualitative

support comes from a study in which we investigated the time point in development, at when the

CSF-lymphatic connection become established (Koh et al., 2006). Overall, these experiments

demonstrated that lymphatics have a major function in removing CSF from the cranium.

Therefore, having established the importance of the cribriform plate-lymphatic pathway in

previous studies, our first objective was to develop a more appropriate method for the routine

comparison of CSF movement across the cribriform plate. Our plan was to utilize this method

to address whether there was an impediment to CSF uptake by lymphatics in CH model (next

chapter) which was made available to us with our collaboration with Dr. McAllister’s group.

20

3.3 Materials and Methods

3.3.1 Animals

A total of 60 adult Fisher 344 rats (148-248 grams) and two 9-day-old pups were used for this

investigation (purchased from Charles River, Canada). The adult animals were fed lab rat chow

(LabDiet 5001) until sacrifice. All experiments were approved by the ethics committee at the

Sunnybrook Health Science Centre and conformed to the guidelines set by the Canadian

Council on Animal Care and the Animals for Research Act of Ontario.

3.3.2 Tracer Infusion and Tissue Sampling

Rats were anesthetized initially by placement in a custom built rodent anesthesia chamber using

halothane (4-5%) in oxygen. For the experimental procedure they were maintained with 2-2.5%

halothane in oxygen delivered by a nose cone (Rat Anesthesia Mask, KOPF, Model 906,

Tujunga, California). The animals were placed on a heating pad (Fine Science Tools,

Vancouver, BC) and fixed in position in a Small Animal Stereotaxic device (KOPF, Model 900,

Tujunga, California). The skin over the cranium was removed and the junction of the sagittal

and coronal sutures identified. A syringe with attached 22-gauge needle (rounded tip) was

positioned on top of the bregma. The coordinates were noted from the stereotaxic instrument

and adjusted according to the reference values from a rat brain atlas (Paxinos & Watson, 2007).

The syringe needle was lowered to make an impression on the skull, which was then marked

with a felt tip pen. A small high speed micro drill with a rounded tip (Fine Science Tools,

Vancouver) was used to grind away the bone to expose the dural membrane.

A 50 µl Hamilton syringe (Fisher Scientific, Toronto, Ontario) with a 30 gauge needle was

loaded with 125I-human serum albumin (HSA) and the tip lowered into one of the lateral

ventricles with the depth determined from the atlas (the right and left ventricles were used

21

randomly in this study). Fifty µl containing 500 µg of 125I-HSA (0.93 MBq/ml, 10 mg/ml, Drax

Image, Quebec) was injected into one of the lateral ventricles. The needle was removed after 1-2

minutes and the needle path sealed with bone wax. After 10, 20, 30, 40 or 60 minutes, the

animals were sacrificed by injection of 1.0 ml euthanyl i.p. Immediately after death, a blood

sample was taken from the heart. A kidney and the spleen were removed and samples from

liver, skeletal muscle and tail were collected. In five animals in the 30-minute group, the lymph

nodes in the neck region including the cervical nodes (6-8) as well as the popliteal (2) and

mesenteric nodes (2-3) were excised. The carcasses were then frozen for at least 24 hours in a

freezer.

3.3.3 Sectioning Olfactory Turbinate Sample and Radioactivity Assessment

To facilitate the assessment of radioactivity in the olfactory submucosa and to prevent potential

post-mortem tracer contamination from the CSF compartment, a portion of the turbinates were

cut from frozen tissue (illustrated schematically in Figure 3-1E). A diagonal line was marked on

each animal at a 45o angle relative to the orbitomeatus line rostral to the orbit. A second line was

drawn 1.0 cm anterior and parallel to the first line and the selected coronal tissue block was

excised with a fine tooth saw. The frozen tissue section was placed flat under a dissecting

microscope (Stereomaster, Cole-Parmer, Quebec) and measured top to bottom. This number was

divided by 4 and the top one-fourth, bottom one-fourth and middle half placed (along with the

other tissue samples collected) in pre-weighed glass test tubes for counting in a multi-channel

gamma spectrometer (Compugamma, LKB Wallac, Turku, Finland). The upper quarter

represented partial superior cells of the ethmoid turbinates along with cartilage and soft tissues

of the nasal wall. The middle portion contained the main portion of the turbinates (henceforth

22

termed middle turbinates). The bottom section consisted of a fraction of the posterior cells of the

turbinates and part of the hard palate.

3.3.4 Visualization of transport across the cribriform plate

We used 2 methods to visualize the movement of CSF across the cribriform plate. In 5 adult

rats, 100 µl Evans blue dye (Fisher Scientific, Toronto, 2% in heparinized sheep plasma) was

injected into the lateral ventricle using methods similar to those employed for the radioactive

tracer. After 20 minutes the rats were sacrificed and frozen. Following decapitation, the heads

were sectioned in either a sagittal, coronal or axial plane. The tissues were examined under a

dissecting microscope (Meiji EMZ-TR, Cole Palmer, Anjou, Quebec) and images were captured

on a Nikon digital camera (Coolpix 995, Henry's Camera, Toronto, ON).

In a second group of 5 adult and 2 pups, yellow Microfil® (MV-122, Flow Tech, MA) was

injected into the cisterna magna post-mortem. The best results were obtained using a preparation

that was more dilute than that recommended in the product literature. In adults, 3 ml of diluent

was used for every 1 ml of yellow Microfil and the material catalyzed with 10% (of total

volume) of MV Curing Agent. In 9-day-old rat pups, 2.5 ml diluent was used with 2.5 ml

Microfil. Immediately after sacrifice in adults, a laminectomy was performed at the C7 cervical-

thoracic level of the vertebral column and a 26 gauge angiocatheter inserted in the cisterna

magna. Microfil (2.5-3 ml) was infused into the cranial subarachnoid space. In pups, 0.2 ml was

injected suboccipitally.

After 20 minutes the animals were frozen and sagittal, coronal or axial slices were made of

the head in preparation for photographic and histological studies. Cranial slices for all studies

were performed to reveal the olfactory bulb and olfactory turbinates using a planar cooled with

dry ice clamped to a bench.

23

For histology, the tissues were harvested and fixed in 10% formalin. The tissues were

paraffin-embedded and cut into 4µm sections and stained with Haematoxylin and/or Eosin.

Histological assessments were performed using a Motic Digital Microscope (DMB5) and

images acquired using Motic Images Advanced 3.0 software (GENEQ Inc. Montreal, Canada).

Good sections of the cribriform plate area were difficult to achieve in adult animals because of

the proximity of bone and soft tissues that caused disruption of the area of interest.

Consequently, for histology we used 9-day-old rat pups. The soft cartilaginous cribriform plate

in these specimens proved much easier to section.

3.3.5 Analysis of Data

Tracer recoveries in all tissues were expressed as percent injected dose/gm tissue. All data were

expressed as the mean ± SD. The results were analyzed with a 2 Factor repeated measures

ANOVA (Grouptime by Tissue) followed by the post-hoc Tukey Studentized Range Test or

Paired t-test as appropriate. We interpreted P<0.05 as significant.

24

3.4 Results

3.4.1 Visualization of CSF transport across the cribriform plate

Twenty min after the injection of Evans blue dye into the CSF compartment, the dye was

observed to distribute throughout the CSF compartment and was found in the subarachnoid

space around the olfactory bulbs and adjacent to the cribriform plate (Figures 3-1 A, B). The

cribriform plate lies at the base of the brain and supports the olfactory bulbs. Figure 1F

illustrates the cribriform plate in an adult rat after removal of the brain and olfactory bulbs. In

addition, Evans blue dye was observed external to the cranium within the olfactory turbinates

(Figure 3-1 A, B). This indicated that material injected into the CSF space was transported

rapidly through the cribriform plate and formed the basis of the quantitative studies that were to

follow. We did not observe any obvious dye at other locations although it was likely that some

would be present within the venous system as was suggested by the vascular entry of the 125I-

HSA (see below). However, the presence of the dye in blood vessels could not be discerned in

the post-mortem state.

The dye could not be observed within individual lymphatic vessels macroscopically

presumably to the dissociation of the dye-protein complex. In addition, the tissues with Evans

blue were not amenable to microscopic analysis as the dye was lost during preparation for

histology. However, Microfil administered into the CSF space could be found in the olfactory

submucosa in an extensive lymphatic network (Figure 3-1 C). Individual lymphatic vessels

containing this contrast agent were clearly visible. On histological examination in young rats

(see Materials and Methods), Microfil appeared in lymphatic vessels throughout the submucosa

(example in Figure 3-1 D).

25

Figure 3-1 CSF transport into olfactory turbinates in the rat. Reference scales are provided either as a ruler (mm) or as a longitudinal bar.

26

Appearance of dye in turbinates (white arrows) after Evans blue dye injection into a lateral

ventricle (sagittal view – A, axial view - B). OB-olfactory bulb; cribriform plate denoted by red

arrows. (C) Filling of lymphatics in the olfactory turbinates after yellow Microfil was injected

into the subarachnoid space post-mortem. Individual lymphatic vessels containing the yellow

contrast agent are clearly visible. (D) Histological section of olfactory turbinates in a 9-day-old

rat pup demonstrating lymphatic vessels filled with Microfil. The yellow Microfil turns dark

brown during histological processing. (E) Schematic of rat head showing method to excise

portion of turbinate tissues. (F) Photograph of cribriform plate from above after removal of the

brain and olfactory bulbs. The cribriform plate lies at the base of the brain and supports the

olfactory bulbs. In rodents, the plates are relatively large structures. LV – lymphatic vessels.

27

3.4.2 Comparison of tracer concentrations in various tissues over time

We observed considerably higher tracer concentrations in the olfactory turbinates compared to

those measured in blood (Figure 3-2). The highest concentrations of 125I-HSA were found in

what we have termed the middle turbinate area, which represented the bulk of the olfactory

turbinates. The lower turbinates (which include some of the posterior cells of the turbinates and

part of the hard palate) also contained high concentrations of the tracer. The radioactivity in the

upper turbinates (representing partial superior cells of the ethmoid turbinates along with

cartilage and soft tissues of the nasal wall) was similar to that observed in blood. The recoveries

of CSF tracer in all olfactory turbinates peaked 30 minutes after injection and at this time, the

average radioactive recoveries in the middle (9.4%) and lower turbinates (5.9%) were 5.9 and

3.7 times higher than the blood average (1.6%) respectively.

In five of the animals in the 30 minute group, various lymph nodes were excised and

assessed for radioactivity. Averaged recoveries in the pooled cervical/neck lymph nodes were

high, being in the same range or even greater than those in the middle turbinates (inset in Figure

3-2). These nodes are interspersed along lymphatic CSF drainage pathways. In contrast, there is

no evidence to suggest that the popliteal and mesenteric lymph nodes collect lymph/CSF from

the subarachnoid compartment and tracer recoveries in these nodes were very low.

CSF ultimately drains into the blood through lymphatics and possibly to some extent

through arachnoid projections. In this regard, blood levels of the radioactive protein peaked 40

minutes after tracer installation but also showed an initial concentration spike at 10 minutes

(Figure 3-3; the blood recoveries from Figure 3-2 have been plotted here with adjustment of the

y-axis scale). Tracer recoveries in skeletal muscle, spleen, kidney, liver and tail were much

lower but seemed to mirror those in blood.

28

TIME (minutes)0 10 20 30 40 50 60

Per

cent

Inje

cted

/gm

Tis

sue

0

2

4

6

8

10

12

14 Bottom TurbinateMiddle TurbinateTop TurbinateBlood

* *

*

* *

*

* Percent Injected/gm Tissue0 5 10 15 20

Blood

Middle Turb

Cerv Nodes

Popliteal Nodes

Mesent Nodes30 Min

*

Figure 3-2 Average CSF tracer recoveries (percent injected dose/gm tissue) in various

tissues as a function of time (n=10 at each time).

29

ANOVA revealed that middle turbinate recoveries were significantly different from those in the

upper or bottom turbinates, blood, skeletal muscle, spleen, kidney, liver and tail with a

significant interaction effect in all cases with the exception of the bottom turbinate. The

concentration of tracer in the 3 turbinate groupings and blood changed significantly over time.

The Tukey’s test indicated that the 30 minute peak tracer levels were significantly different from

those at the other time points in both the bottom and middle turbinates. Significant differences

between the various turbinates and blood assessed with a Paired t-test are illustrated with

asterisks.

The inset represents data from a subgroup of these animals (n=5) in which tracer recoveries in

the middle turbinates and blood were compared with those in various lymph nodes (Mesent –

mesenteric; Cerv – cervical). Significant differences measured with Paired students t test

(asterisk).

30

TIME (minutes)0 10 20 30 40 50 60

Per

cent

Inje

cted

/gm

Tis

sue

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5 BloodSkeletal MuscleSpleenKidneyLiverTail *

*

*

*

*

Figure 3-3 Average CSF tracer recoveries (percent injected dose/gm tissue) in various

tissues as a function of time (n=10 at each time).

ANOVA revealed that blood recoveries were significantly different from those in skeletal

muscle, spleen, kidney, liver and tail with a significant interaction effect in all cases. The

concentration of tracer in blood changed significantly over time and the Tukey’s test indicated

that the peak tracer levels at 40 minutes were significantly different from those at the other

times. Paired t-tests revealed that blood recoveries were significantly different from all other

tissues at all time points (asterisks in Figure).

31

3.5 Conclusion

Our objective in this study was to develop a relatively simple quantitative approach to assess

lymphatic CSF absorption in rats. Once CSF is taken up by lymphatics adjacent to the

cribriform plate, it enters a network of vessels in the olfactory turbinates and consequently, it

seems reasonable to expect that the turbinate tissues would have high concentrations of a CSF

tracer. This assumption was supported by the studies outlined in this chapter and we concluded

that the turbinate protocol would provide a routine measure of lymphatic CSF absorption that

can be applied to studies of the pathophysiological implications of lymphatic CSF transport in

hydrocephalus.

The content presented in this chapter has been previously published: Nagra G, Koh L, Zakharov A, Armstrong D and Johnston M. Quantification of cerebrospinal fluid transport across the cribriform plate into lymphatics in rats. Am J Physiol Regul Integr Comp Physiol 291: R1383-R1389, 2006. doi:10.1152/ajpregu.00235.2006

32

CHAPTER 4:

DOES A CSF LYMPHATIC ABSORPTION DEFICIT EXIST IN A KAOLIN

MODEL OF HYDROCEPHALUS AND DOES THE LYMPHATIC DEFECT

CORRELATE WITH HYDROCEPHALUS?

33

4.1 Abstract

It has been assumed that the pathogenesis of hydrocephalus includes a cerebrospinal fluid (CSF)

absorption deficit. Since a significant portion of CSF absorption occurs into extracranial

lymphatics located in the olfactory turbinates, the purpose of this study was to determine if CSF

transport was compromised at this location in a kaolin-induced communicating

(extraventricular) hydrocephalus model in rats. Under 1-3% Halothane anesthesia, Kaolin

(n=10) or saline (n=9) was introduced into the basal cisterns of Sprague Dawley rats and the

development of hydrocephalus assessed 1 week later using MRI. Following injection of human

serum albumin (125I-HSA) into a lateral ventricle, the tracer enrichment in the olfactory

turbinates 30 minutes post-injection provided an estimate of CSF transport through the

cribriform plate into nasal lymphatics. Lateral ventricular volumes in the kaolin group (0.073 ±

0.014 ml) were significantly greater than those in the saline injected animals (0.016 ± 0.001 ml;

p = 0.0014). The CSF tracer enrichment in the olfactory turbinates (expressed as percent

injected/gm tissue) in the kaolin rats averaged 0.99 ± 0.39 and was significantly lower than that

measured in the saline controls (5.86 ± 0.32; p < 0.00001). The largest degree of

ventriculomegaly was associated with the lowest levels of lymphatic CSF uptake with lateral

ventricular expansion occurring only when almost all of the lymphatic CSF transport capacity

had been compromised. We conclude that lymphatic CSF absorption is impaired in a kaolin

communicating hydrocephalus model and that the degree of this impediment may contribute to

the severity of the induced disease.

34

4.2 Introduction

As a natural progression after developing a quantitative model to measure CSF absorption in rat,

we wanted to investigate if there is any evidence of an absorptive defect at the level of the

cribriform plate in a model of communicating hydrocephalus.

Many animal models of hydrocephalus have been developed (McAllister & Chovan,

1998;Hochwald, 1985;Johanson & Jones, 2001) but most mimic various forms of “obstructive”

hydrocephalus by blocking CSF flow at the 4th ventricle outlets. Communicating hydrocephalus

(CH) is characterized by impaired CSF flow only in the subarachnoid spaces and occurs

frequently, especially in older adults (Hoppe-Hirsch et al., 1998;Kosteljanetz, 1986). CH has

been difficult to model because the subarachnoid spaces are extremely small and difficult to

access. Several attempts have been made in the past 20 years to induce CH in dogs and rats, but

the methods used do not produce the disorder consistently and the site of obstruction has not

been determined unequivocally. CH has been induced in adult dogs by injecting kaolin (Deo-

Narine et al., 1994;James, Jr. et al., 1975;James, Jr. et al., 1978;Strecker et al., 1974) or silicone

(Price et al., 1976) into the subarachnoid space. Likewise, silastic has also been infused into the

basal cisterns of monkeys to produce CH (Diggs et al., 1986). In all of these early studies, the

severity of hydrocephalus was either not reported or was quite variable, and the survival periods

seldom exceeded 30 days. Non-mechanical induction methods such as viral (Davis, 1981) and

bacterial inoculations (Wiesmann et al., 2002) have also been used to produce CH, but all of

these procedures include the additional influence of the induction substances on the

pathophysiology, and thus are not true representations of the effects of hydrocephalus alone on

the brain. Growth factors such as TGF- and FGF-2 (Johanson et al., 1999;Moinuddin & Tada,

2000;Tada et al., 1994) and neurotoxins (Fiori et al., 1985) have all been successful to varying

degrees. Congenital and transgenic models also exist for CH (Dahme et al., 1997;das Neves L.

35

et al., 1999;Stoddart et al., 2000;Galbreath et al., 1995;Kume et al., 1998;Robinson et al.,

2002;Borit & Sidman, 1972;Moinuddin & Tada, 2000;Bruni et al., 1988;Jones, 1985), but these

all require the use of mice, which are too small for meaningful physiological investigation.

Kaolin, an inert silica derivative, is a well-accepted agent for inducing hydrocephalus in infant

and adult animals (mouse, rat, rabbit, hamster, cat, dog) via injections into the CSF, with no

evidence of direct pathological effects on structures distant to the injection site. While this

induction method is mechanical (surgical) and can produce hydrocephalus abruptly, which is not

always the time course found clinically, it is useful when more “natural” models are not

available. Indeed, in many applications kaolin administration is used to block the outlets of the

fourth ventricle (leading to obstructive or non-communicating hydrocephalus). These

approaches have yielded valuable data on the pathophysiology that appears secondary to

ventriculomegaly. In contrast, Dr. McAllister’s group has developed a CH model in rat by

injecting kaolin into the basal cisterns which does not obstruct the fourth ventricular outlets (Li

et al., 2008). We used this CH rat model as it was made available to us with our collaboration

with Dr. Pat McAllister to test the hypothesis that ventriculomegaly may be related to impaired

lymphatic function.

36


4.3.1 Animals

A total of 54 female Sprague Dawley rats with an average weight of 246.01 ± 2.41 were used

for this investigation (purchased from Harlan, Indiana, USA and Harlan, Canada). The animals

were fed lab rat chow (LabDiet 5001) until sacrifice.

The injections of kaolin or saline were performed in the laboratory of Dr. McAllister at the

Wayne State University School of Medicine according to the protocol approved by the

institutional Animal Investigation Committee. The lymphatic studies were carried out in Dr.

Johnston's laboratory at Sunnybrook Health Science Centre in Toronto. These experiments were

approved by the ethics committee at the Sunnybrook HSC and conformed to the guidelines set

by the Canadian Council on Animal Care and the Animals for Research Act of Ontario.

Magnetic Resonance Imaging (MRI) studies (to determine ventricle sizes) were performed

about one week after the injection of saline (average 8.7 ± 1.5 days) or kaolin (average 7.8 ± 0.1

days) into the basal cisterns. The animals were then sent to Toronto for lymphatic analysis about

one week after this (average time from induction of hydrocephalus to lymphatic studies were

17.3 ± 1.2 days for saline- and 15.0 ± 0.6 days for kaolin-injected rats).

4.3.2 Induction of communicating hydrocephalus

As described in detail previously (Li et al., 2008), the rats were anesthetized with a mixture of

1-3% Halothane with 40% oxygen, and using aseptic techniques the skin was incised along the

ventral midline of the neck. After the soft tissues were reflected to expose the base of the skull,

a 30-gauge needle, custom bent to 30º, was inserted into the sub-arachnoid space between the

clivus and the C1 vertebra. The needle was advanced 1.5 – 2.0 mm along the inner surface of

the cranial cavity, and a 50-µl sterile suspension of 25% kaolin in saline was injected at the rate

37

of about 6l/sec. The surgical incision was closed in layers using absorbable suture (Vicryl)

and the rats were allowed to recover. Post-operatively, Butorphenol was given subcutaneously

(0.05-2.0 mg/kg) every 4-8 hours as needed to control pain. Animals that experienced breathing

difficulty (coughing) or symptoms of life-threatening increases in intracranial pressure post-

operatively received Mannitol (1.5g/kg IV). Sham controls were prepared in a similar fashion

but received sterile saline only (308 mOsM/L, pH was 7.4).

4.3.3 Assessment of ventricular size

MRI was used at Wayne State University to measure ventricular size in vivo. After anesthesia

with a mixture of 87 mg/kg Ketamine plus 13 mg/kg Xylazine, the animal was placed into a

4.7T magnet and coronal and sagittal T1- and T2-weighted images were obtained (TE/TR =

20/700 ms and 67/5000ms, respectively) on 1.0 mm slice thickness. Ventricular volumes (lateral

and 3rd ventricles) were calculated from T2 images, starting from the center of the cerebral

aqueduct up to the anterior-most portion of the lateral ventricles. The volumetric calculations

were semi-automated as follows; an appropriate intensity threshold was first chosen to exclude

background tissue and to highlight the bright ventricles. This was followed by careful

inspection of each image, and manual tracing was used to correct any areas of the ventricle,

which had been incorrectly deleted, or to delete non-ventricular regions that had been

incorrectly included. This process resulted in a binary mask of ventricular pixels, which when

multiplied by the volume of each pixel and summed over all slices produced the net ventricular

volume in ml. The Evan’s ratio was taken at the level of the Foramen of Monroe as the

maximum width of both lateral ventricles divided by the maximum width of the brain at this

level.

38

4.3.4 Assessment of lymphatic CSF absorption

The method to assess lymphatic CSF uptake in the rat has been described in Chapter 3 of this

thesis. However, it is important to note that our collaborators used a different species of rats,

Sprague Dawley for the kaolin model of CH. Although, there is no reason to believe that the

quantitative CSF-lymphatic absorption analysis using Sprague Dawley rats should not be similar

to the Fisher 344, the species of rats which we used to develop the quantitative method but, we

did re-analyze the transport in normal control Sprague Dawley rats in order to figure out the

optimal time to sample the tissues. We found that this time was 30 minutes post radioactive

injection in both species.

4.3.5 Visualization of transport across the cribriform plate in kaolin and saline injected

animals

Evan’s blue dye (2%) was used to visualize the movement of CSF across the cribriform plate

post mortem in 2 kaolin and 2 saline injected rats. The animals were anesthetized as described

above. Following a midline incision the skin was reflected over the cranium. An Alzet rat brain

cannula (Direct Corporation, Cupertino, CA) was inserted into the cortex with the tip positioned

in the lateral ventricle. The cannula was secured to the skull with Surehold glue (mixture of

ethyl cyanoacrylate and polymethylmethacrylate, Surehold, Chicago, IL). Immediately after

sacrifice, 0.3ml of Evan’s blue dye was infused over 30 seconds into the lateral ventricle. The

animals were frozen overnight and following decapitation, the heads were sectioned in

preparation for photographic studies. The images were captured on a Nikon digital camera

(Coolpix 995, Henry’s Camera, Toronto, ON).

39


In the first group of experiments, we determined the most appropriate time after injection of the

radioactive tracer using 30 animals (6 rats at each time of 10, 20, 30, 40 and 60 minutes) to

assess the tracer enrichment in the olfactory turbinates. In a second group, we used Evans blue

dye in attempt to visualize CSF-lymphatic connections in 2 saline- and 2 kaolin-injected rats. In

the final group, we examined the lymphatic uptake of the tracer in 10 saline- or 10 kaolin-

injected animals (1 animal in the saline group died during the experiment and could not be used

in the data analysis). The enrichment of the CSF tracer in various tissues was expressed as

percent injected dose/gm tissue. All data were expressed as the mean ± SE. The data were

analyzed with an unpaired two-sample t-test and a Wilcoxon non-parametric equivalent. We

interpreted p<0.05 as significant.

40

4.4 Results

4.4.1 Development of hydrocephalus

Most rats undergoing kaolin injections exhibited relatively normal behavior post-operatively

although many of the kaolin-injected animals exhibited bloody nasal and orbital secretions and

coughed frequently. We presumed that these symptoms related to increases in intracranial

pressure although no pressures were measured in this study. The coughing either disappeared

normally within a few days or was alleviated by administration of mannitol, which is known to

reduce CSF pressure. Regardless of how severe ventriculomegaly had become, no animals

exhibited signs of cranial enlargement.

MRI analysis revealed that nearly all kaolin deposits on the ventral brainstem were

bilateral and contiguous. The largest formations extended from the medulla to the

interpeduncular fossa, and all covered a portion of the pons. Some kaolin deposits were small

and located unilaterally. No kaolin deposits blocked the foramina of Luschka grossly or

occupied the cisterna magna. Most importantly, no kaolin deposits were located anterior to the

optic chiasm, including the region of the cribriform plate.

Compared to saline-injected controls, the ventricular system expanded to relatively moderate

and severe levels in most kaolin-injected animals (examples in Figure 4-1). All portions of the

ventricular system exhibited expansion. Enlargement of the lateral ventricles was symmetrical

and proportionally greater than other regions of the ventricular system. The cerebral aqueduct

was patent and dramatically expanded posteriorly. This indicated that the hydrocephalus in this

model is of the communicating type. More details on the morphological changes that

accompany this model have been described previously (Li et al., 2008).

Figure 4-1 also illustrates the average ventricular volumes (C) and Evans Ratios (D) in the

two groups. In the kaolin injected rats the ventricular volumes were significantly greater

(ranging from 0.015 - 0.162 ml - average 0.073 ± 0.014 ml) than those observed in the saline

41

injected animals (ranging from 0.011 - 0.022 ml - average 0.016 ± 0.001 ml). In the kaolin

injected group the Evans ratios ranged from 0.36 - 0.58 (average 0.48 ± 0.02) and were

significantly greater than those in the saline injected animals (ranging from 0.25 - 0.39, average

0.35 ± 0.01).

42

Saline Kaolin

Lateral Ventricles

FM

3rd Ventricle

Lateral Ventricles

A B

0.0

0.1

0.2

0.3

0.4

0.5

0.6

Saline Kaolin

*

C D

Saline Kaolin

Vent

ricul

ar V

olum

e (m

l)

Evan

’s R

atio

Saline Kaolin

Lateral Ventricles

FM

3rd Ventricle

Lateral Ventricles

A B

0.0

0.1

0.2

0.3

0.4

0.5

0.6

Saline Kaolin

*

C D

Saline Kaolin

Vent

ricul

ar V

olum

e (m

l)

Evan

’s R

atio

Figure 4-1 Impact of saline or kaolin injection on the ventricular volumes.

43

Representative MRI images of saline (A) and kaolin (B) injected animals to illustrate the extent

of ventriculomegaly achieved in the experimental group. These coronal sections taken

approximately 1 week after injection show the bilateral enlargement of the frontal horns of the

lateral ventricles at the foramina of Monro (FM), as well as the expansion of the third ventricle

in the kaolin-injected animal. Ventricular volumes and Evans Ratios are illustrated in (C) and

(D) respectively. A 2-sample unpaired t-test (p = 0.0014) and a Two-Sided Wilcoxon non-

parametric test (p = 0.0042) indicated that the ventricular volumes for the kaolin group (n=10)

were significantly higher than those measured in the saline-injected animals (n=9). Similar

analysis of the Evans Ratios revealed significant differences in the saline and Kaolin-injected

rats (2-sample unpaired t-test, P= 0.0001; Two-Sided Wilcoxon non-parametric test (p =

0.0042).

44

4.4.2 Assessment of CSF transport through cribriform plate

Prior to the hydrocephalus experiments, we conducted a study to determine the most appropriate

time to assess tracer enrichment in the olfactory turbinates of Sprague Dawley rats. The data are

illustrated in Figure 4-2. The highest concentrations of 125I-HSA were found consistently in the

middle turbinate area (average weights 0.36 ± 0.03 gm), which represented the bulk of the

olfactory turbinates. For this reason, we chose to assess the lymphatic uptake of CSF using

tracer enrichment in the middle turbinates. The lower turbinates also contained high

concentrations of the tracer. The tracer enrichment at these locations was much greater than that

observed in blood (up to 40 min). The radioactivity in the top turbinates was similar to that

observed in blood. Tracer recoveries in skeletal muscle, spleen, kidney, liver and tail were much

lower than those observed in the lower and middle turbinates and likely were reflective of the

tracer within the vasculature of these tissues (not illustrated in Figure 4-2). On average, tracer

concentrations in the middle and bottom turbinates peaked at 30 minutes after ventricular

injection. This time was chosen to compare the transport of the CSF tracer across the cribriform

plate in the kaolin and saline injected animals.

45

Time (minutes)0 10 20 30 40 50 60 70

Enr

ichm

ent o

f Tra

cer

0

2

4

6

8

10

12Bottom TurbinatesMiddle TurbinatesTop TurbinatesBlood

Figure 4-2 Average CSF tracer enrichment (percent injected dose/gm tissue) in the

olfactory turbinates and blood as a function of time (n=6 at each time).

Tracer enrichment in the lower and middle olfactory turbinates was greater than that measured

in the upper turbinates or blood and peaked at 30 minutes after tracer injection into a lateral

ventricle. Tracer concentrations in skeletal muscle, spleen, kidney, liver and tail were very low

and are not illustrated in the figure.

46

4.4.3 Transport of CSF tracer into olfactory turbinates: comparison between kaolin and

saline injected animals

Initial studies were performed with Evan's blue dye injected into the lateral ventricles to

visualize the movement of the contrast agent within the CSF system and its possible entry into

the olfactory turbinates. Evan’s blue dye was observed in the lateral ventricles, and

subarachnoid space of both kaolin and saline injected animals. However, the appearance of dye

in the olfactory turbinates was much less prominent in the kaolin- injected animals compared

with the saline-injected group. Examples of the appearance of Evan’s blue dye in the kaolin and

saline injected animals are provided in Figure 4-3 A and B, respectively.

Figure 4-3 C illustrates two examples of tracer enrichment in one saline-injected (ventricular

volume - 0.020) and one kaolin-injected animal (ventricular volume - 0.099). The 125I-HSA

enrichment in the olfactory turbinates was much less in the kaolin-injected animal. While it

would appear that tracer enrichment in the other tissues was also less in this kaolin rat, this was

not a consistent pattern for all animals.

47

Figure 4-3 Comparison of lymphatic CSF absorption in the saline - and kaolin

injected animals.

48

Sagittal section of a saline (A) and kaolin (B) injected animal. Evan’s blue dye was injected into

a lateral ventricle post mortem. The turbinates of the saline injected animal are densely stained

with the dye. In contrast, the turbinates of the kaolin injected animal contained less dye. The

scale in mm is provided at the bottom of the images. C) Examples of the distribution of

radioactivity into various tissues after 125I-HSA injection into a lateral ventricle. Black bars

represent data from a saline-injected rat. White bars illustrate data from a kaolin-injected

animal. The ventricular volumes for both examples are provided in the inset. The enrichment of

the tracer in the turbinate tissues in the saline example was much greater than that in the kaolin-

injected rat. In the latter case, the ventricles were enlarged (0.099 cubic centimeters compared to

0.020 cubic centimeters for the saline animal).

49

Figure 4-4 A illustrates the average middle turbinate enrichment data assessed at 30 minutes

after tracer injection. The movement of the CSF tracer across the cribriform plate in the kaolin-

injected rats was significantly less that that measured in the saline group. Indeed, the tracer

enrichment in the kaolin animals was only 17 % of that observed in the controls. While the

tracer concentration in blood appears to be higher in the kaolin group (inset to Figure 4-4 A),

these differences were not significant. Additionally, there were no saline/kaolin related

significant differences in the radioactivity measured in any of the other tissues (data not

illustrated).

Figure 4-4 B illustrates the ventricular volumes for all saline (n=9, closed circles) and kaolin

injected animals (n=10, open circles) plotted against the corresponding 30-minute middle

turbinate enrichment data for these rats. The exponential-like relationship suggests that the

largest ventricles (most severe hydrocephalus) were associated with the lowest CSF transport

into ethmoidal lymphatic vessels. Indeed, ventricular volumes did not appear to increase until

almost all of the tracer movement across the cribriform plate had been abolished.

50

Figure 4-4 Comparison of lymphatic CSF absorption in the saline (n=9) and kaolin-

injected animals (n=10).

51

One saline-injected rat died before tracer assessment. A) Impact of Saline or Kaolin injection

on the middle turbinate enrichment of the CSF tracer. The movement of the CSF tracer across

the cribriform plate in the kaolin-injected rats was significantly less that that measured in the

saline group (2-sample unpaired t-test, p < 0.0001; Two-Sided Wilcoxon non-parametric test, p

< 0.0001). The tracer concentrations in the blood in the 2 groups are illustrated in the inset to A

(no significant differences). B) A plot of the turbinate enrichment of tracer versus ventricle size

yielded an exponential relationship (saline-injected, closed circles; kaolin-injected, open

circles). These data suggested that the largest ventricles (most severe hydrocephalus) were

associated with the lowest CSF transport into ethmoidal lymphatic vessels.

52

4.5 Conclusion

Using our quantitative method to assess CSF transport into the lympathics we investigated

whether an impediment to lymphatic uptake of CSF correlate with hydrocephalus. We found

that the largest degree of ventriculomegaly was associated with the lowest levels of lymphatic

CSF uptake with lateral ventricular expansion occurring only when almost all of the lymphatic

CSF transport capacity had been compromised. We conclude that lymphatic CSF absorption is

impaired in a kaolin communicating hydrocephalus model and that the degree of this

impediment may contribute to the severity of the disease.

The content presented in this chapter has been previously published: Nagra, G, Li J., Mcallister P, Miller J, Wagshul, M, Johnston, M. Impaired lymphatic cerebrospinal fluid absorption in a rat model of kaolin-induced communicating hydrocephalus: Am.J Physiol Regul.Integr.Comp Physiol, v. 294, p. R1752-R1759, 2008. doi:10.1152/ajpreg.00748.2007

53

CHAPTER 5:

DOES A REDUCTION IN LYMPHATIC CSF ABSORPTION CORRELATE WITH A

GLOBAL CSF ABSORPTION DEFICIT?

54

5.1 Abstract

We recently reported a lymphatic cerebrospinal fluid (CSF) absorption deficit in a kaolin model

of communicating hydrocephalus in rats with ventricular expansion correlating negatively with

the magnitude of the impediment to lymphatic function. However, it is possible that CSF

drainage was not significantly altered if absorption at other sites compensated for the lymphatic

defect. The purpose of this study was to investigate the impact of the lymphatic absorption

deficit on global CSF absorption (CSF outflow resistance). Kaolin was injected into the basal

cisterns of Sprague Dawley rats. The development of hydrocephalus was assessed using MRI. In

one group of animals at about 3 weeks after injection, the movement of intraventricularly

injected human serum albumin (125I-HSA) into the olfactory turbinates provided an estimate of

CSF transport through the cribriform plate into nasal lymphatics (n=18). Control animals

received saline in place of kaolin (n=10). In a second group at about 3.5 weeks after kaolin

injection, intraventricular pressure was measured continuously during infusion of saline into the

spinal subarachnoid space at various flow rates (n=9). CSF outflow resistance was calculated as

the slope of the steady-state pressure versus flow relationship. Control animals for this group

either received no injections (intact: n=11) or received saline in place of kaolin (n=8). Compared

to saline injected controls, lateral ventricular volume in the kaolin group was significantly

greater (0.087 ± 0.013 ml, n=27 versus 0.015 ± 0.001 ml, n=17) and lymphatic function was

significantly less (2.14 ± 0.72 % injected/gm, n=18 versus 6.38 ± 0.60 % injected/gm, n=10).

Additionally, the CSF outflow resistance was significantly greater in the kaolin group (0.46 ±

0.04 cm H2O.µL-1.min, n=9) than in saline injected (0.28 ± 0.03 cm H2O.µL-1.min, n=8) or

intact animals (0.18 ± 0.03 cm H2O.µL-1.min, n=11). There was a significant positive correlation

between CSF outflow resistance and ventricular volume. The data suggest that the impediment

to lymphatic CSF absorption in a kaolin-induced model of communicating hydrocephalus has a

55

significant impact on global CSF absorption. A lymphatic CSF absorption deficit would appear

to play some role (either direct or indirect) in the pathogenesis of ventriculomegaly.

56

5.2 Introduction

The aforementioned data suggest that a CSF absorption deficit contributed in some way to the

pathogenesis of hydrocephalus. However, in this kaolin-induced hydrocephalus model it is

possible that an obstruction to CSF transport at one location (the cribriform plate and

lymphatics) is met by enhanced CSF clearance through alternative absorption sites. These could

include other lymphatic vessels or the arachnoid granulations. To begin to answer this question,

it would be informative to determine if the global CSF outflow resistance is increased in the

kaolin hydrocephalus rat model. To address this issue, we report on measurements of global

CSF outflow resistance in this model of adult communicating hydrocephalus. These studies

show that outflow resistance is elevated significantly and that a global CSF absorption deficit

correlates negatively with ventricular volume.

57


5.3.1 Animals

A total of 56 female Sprague Dawley rats (234-287 g, average 261.9 gm) were used for this

investigation (purchased from Harlan, Indiana, USA and Charles River, Canada). The animals

were fed lab rat chow (LabDiet 5001) until sacrifice.

5.3.2 Details of collaboration

The injections of kaolin or saline were performed in the laboratories of Dr. McAllister at the

Wayne State University School of Medicine and MRI analysis to measure the ventricular

volume were performed at Dr. Mark Wagshul's laboratory at Stony Brook University according

to the protocol approved by each institutional Animal Investigation Committee. The lymphatic

studies were carried out in Dr. Johnston's laboratory at Sunnybrook HSC in Toronto. These

experiments were approved by the ethics committee at the Sunnybrook Health Science Centre

and conformed to the guidelines set by the Canadian Council on Animal Care and the Animals

for Research Act of Ontario.

At approximately 2.5 weeks following kaolin or saline injection, Magnetic Resonance

Imaging (MRI) studies were performed to determine ventricle size. Two separate groups of

animals were used to study lymphatic CSF absorption and CSF outflow resistance. Animals

were sent to Toronto following MRI measurements at approximately 3 weeks post injection to

perform these analyses.

5.3.3 Induction of communicating hydrocephalus

The details on how the CH was induced is briefly described in the previous chapter of this thesis

and elsewhere (Li et al., 2008).

58

5.3.4 Assessment of ventricular size

MRI was used at Wayne State University to measure ventricular size in vivo. This has been

described in detail, in the previous chapter of this thesis.

5.3.5 Assessment of lymphatic CSF absorption

The method to assess lymphatic CSF uptake in the rat has also been described in the previous

chapter of this thesis.

5.3.6 Outflow Resistance

At about 3.5 weeks post injection, the animals were anesthetized and the skull stabilized in a

stereotaxic device as described above. These animals were either injected with kaolin (n=9),

saline (n=8), or not injected (intact, n=11). Following a midline incision, the skin was reflected

over the cranium and a hole was drilled into the skull for eventual insertion of a brain cannula

into a lateral ventricle for CSF pressure measurement (same stereotaxic co-ordinates as

radioactive tracer injection). Animals were then taken out of the stereotaxic frame and a

tracheotomy was performed. A three-way stopcock supplying 1.5% oxygen mixed isofluorane

and air exhaust (Bickford Inc., Wallis Centre, New York, USA) was connected to a 14-gauge

catheter (Infusion Therapy System Inc., CE0086, Sandy, USA), which was inserted into the

trachea tube. The tracheal catheter was secured with Surehold glue (mixture of ethyl

cyanoacrylate and polymethylmethacrylate, Surehold, Chicago, USA) and the skin sutured with

4.0 silk (suture).

Access to the SAS for infusion of artificial CSF was achieved with a laminectomy to expose

the SAS at the lumbar level. A 24-gauge needle containing a 1.9 cm catheter (Terumo, Surflo

I.V. Catheter 2227) was inserted ~0.25 cm into the SAS using a surgical microscope (Carl Zeiss

OPMI 1-FC). Once the catheter was in place the needle was removed. When CSF had filled the

59

catheter, it was connected via an adapter (Lot 99575890 Argon Medical Devices, Athens, USA)

to a 5 ml syringe containing sterile Ringer's solution mounted in a syringe pump (Stoelting Co.,

catalogue # 53200, Wood Dale, USA).

At this point, an Alzet rat brain cannula (Direct Corporation, Cupertino, USA) was inserted

into the burr hole with the tip positioned in the lateral ventricle. The cannula was secured to the

skull with Surehold glue. Intraventricular pressure was monitored with the brain cannula

connected to a pressure transducer (Custom CDX3 with stop cock, Lot 99471637, Richmond

Hill, Canada). This system was calibrated first using a water reservoir at the beginning of the

surgical procedure. The data were recorded using a data acquisition system (Daq Software, A-

Tech Instuments, Toronto, Canada).

A stable baseline pressure was recorded for each experiment before the infusion of Ringer's

solution into the lumbar SAS. Flow rates were adjusted incrementally (10, 22, 34, 50, 100 and

153 µL/min) and steady-state pressures were established for 3-6 minutes at each flow rate before

increasing the rate to the next level.


Out of the total of 56 animals, we have included data from 19 rats that were used for the study

outlined in Chapter 4. These include measures of lymphatic function and ventricular volumes

following kaolin and saline injections that were collected in experiments conducted over a 3-

year period. All data are expressed as the mean + SEM. The data were assessed as suited with

the t-test, linear regression, ANOVA, or the Kruskal-Wallis test and as appropriate, post-hoc

Bonferroni using SPSS 17 software.

60

5.4 Results

5.4.1 Development of hydrocephalus

MRI analysis revealed that nearly all kaolin deposits on the ventral brainstem were bilateral and

contiguous. The largest formations extended from the medulla to the interpeduncular fossa, and

all covered a portion of the pons. No kaolin deposits blocked the foramina of Luschka or

occupied the cisterna magna. Additionally, no kaolin deposits were located anterior to the optic

chiasm, including the region of the cribriform plate. Compared to saline-injected controls, the

ventricular system expanded to relatively moderate and severe levels in most kaolin-injected

animals (a post-mortem example is illustrated in Figure 5-1 B). MRI revealed that the cerebral

aqueduct was patent and dramatically expanded posteriorly. This indicated that the

hydrocephalus in this model is of the communicating type. More details on the morphological

changes that accompany this model have been described previously (Li et al., 2008).

Figure 5-2 A illustrates the average ventricular volumes in the saline and kaolin-injected

groups. In the kaolin injected rats the ventricular volumes were significantly greater (0.087 ±

0.013 ml, n=27) than those observed in the saline injected animals (0.015 ± 0.001 ml, n=17)

(p<0.0001).

5.4.2 Transport of CSF tracer into olfactory turbinates: comparison between kaolin and

saline injected animals

In past studies, we determined that the highest concentrations of 125I-HSA were found

consistently in the middle turbinate area, which represented the bulk of the olfactory turbinates.

For this reason, we chose to assess the lymphatic uptake of CSF using tracer enrichment in the

middle turbinates.

61

Figure 5-2 B illustrates the average middle turbinate enrichment data assessed at 30 minutes

after tracer injection. The movement of the CSF tracer across the cribriform plate in the kaolin-

injected rats (2.14 ± 0.72, n=18) was significantly less that that measured in the saline group

(6.38 ± 0.60, n=10) (p<0.0001). Indeed, the tracer enrichment in the kaolin animals was only

34% of that observed in the saline controls.

62

Figure 5-1 Example of hydrocephalus induced with administration of kaolin into the

basal cisterns. (A) saline injected, (B) kaolin injected

63

Figure 5-2 Ventricle size and lymphatic CSF uptake.

64

(A) Ventricular volumes in the kaolin injected rats were significantly greater than those in the

saline injected animals (p<0.0001; independent t-test).

(B) The enrichment of radioactive protein tracer in the olfactory turbinates (lymphatic CSF

uptake) was significantly less in the animals receiving kaolin (p<0.0001; independent t-test).

The numbers of animals in each group are indicated below each histogram. The numbers within

the brackets represent data taken from Chapter 4 (Nagra et al., 2008).

65

5.4.3 CSF outflow resistance

CSF outflow resistance was calculated from the slopes of the pressure versus inflow infusion

rate data. At each infusion rate, pressure increases until a steady-state is attained; at this point

the infusion rate is increased. At the end of the experiment, the average steady-state pressures

were plotted against the infusion rate.

The average slopes of pressure/flow relationships (outflow resistances) are illustrated in

Figure 5-3. The CSF outflow resistance for the kaolin group (0.46 ± 0.04 cm H2O/µL/min) was

significantly greater than that observed in the saline-injected animals (0.28 ± 0.03 cm

H2O/µL/min) (p=0.004) or in the intact, non-injected rats (0.18 ± 0.03 cm H2O/µL/min) (p<

0.0001).

Figure 5-4 illustrates a significant positive correlation between CSF outflow resistances and

ventricular volume (r2=0.583, p=0.001), with the largest ventricular volumes associated with the

highest levels of outflow resistance.

66

Figure 5-3 Average CSF outflow resistance in intact, saline or kaolin-injected animals.

Analysis by both the Kruskal Wallis (p<0.0001) test and ANOVA (p<0.0001) revealed that the

3 groups were significantly different. Post-hoc Bonferroni indicated that the outflow resistance

in the kaolin group was significantly greater than that of the saline (p=0.004) and intact groups

(p<0.0001) but no significant differences were observed between saline and intact animals

(p=0.082).

67

Figure 5-4 Relationship between CSF outflow resistance and ventricular volumes with

both parameters measured in the same animals (n=16).

There was a significant positive correlation between CSF outflow resistance and ventricular

volume (p = 0.001, Pearson correlation; r2 = 0.583).

68

5.5 Conclusion

We found that the lateral ventricular volumes in the kaolin group were significantly greater and

lymphatic function significantly less than those in the saline injected animals. Additionally, the

CSF outflow resistance in the kaolin group was significantly greater than that in saline injected

or intact animals. A plot of the outflow resistance versus ventricular volumes revealed a

significant relationship with the largest volumes associated with the highest resistances.

The data suggest that the impediment to lymphatic CSF absorption observed in a kaolin-

induced model of communicating hydrocephalus has a significant impact on global CSF

absorption. The elevation in CSF outflow resistance appears to relate to an impediment to CSF

clearance through the cribriform plate into extracranial lymphatic vessels. It would seem

therefore, that a lymphatic CSF absorption deficit plays some role in the pathogenesis of

ventriculomegaly. However, whether association between CSF absorption and ventricle size

originates from a direct cause and effect relationship or arises secondarily from a complex

interplay of other unknown physiological parameters is not known.

69

CHAPTER 6:

LYMPHATIC CSF ABSORPTION AND HYDROCEPHALUS DISCUSSION

70

6.1 Discussion of Results

6.1.1 Quantifying lymphatic CSF absorption

With a quantitative method, we hoped to capture a time when the injected radioactive tracer

would give a ‘signal’ that would be sufficiently high to warrant the development of a routine

method to assess CSF movement across the cribriform plate. This was done by sampling a

tissue through which lymphatic vessels are transporting CSF. Focusing on the extracranial

olfactory tissues seems to have achieved this goal since the concentration of the CSF tracer was

much higher in the turbinates than in the other tissues monitored. However, there are several

limitations in using turbinate concentrations of a CSF tracer to assess the transport of CSF by

lymphatics.

6.1.1.1 Limitations of Quantitative method

First, we could not measure lymphatic uptake of the protein tracer directly with the methods

employed, as described in chapter 3. However, past studies indicate that the olfactory turbinates

contain an extensive network of lymphatic vessels and many of these ducts play an important

role in CSF absorption in mice, rats, rabbits, sheep, pigs, non-human primates and humans

(Johnston et al., 2004;Johnston et al., 2005). Furthermore, the evidence suggests that CSF

convects directly into lymphatic vessels adjacent to the cribriform plate rather than disperse

throughout the interstitium of the olfactory submucosa. Our experience with Microfil in a

variety of species would indicate some form of direct linkage exists between the CSF and

extracranial lymph (Johnston et al., 2004;Johnston et al., 2005;Koh et al., 2006;Zakharov et al.,

2004a). Additionally, the turbinate lymphatic vessels taking up yellow Microfil that was

injected into the CSF space could be distinguished clearly from blood vessels after blue Microfil

had been infused into the vasculature. In the studies of Kida and colleagues, the injection of

71

Indian ink into the cisterna magna of rats resulted in carbon particles passing from the

subarachnoid space beneath the olfactory bulbs into discrete channels that crossed the cribriform

plate into nasal submucosal lymphatic vessels (Kida et al., 1993). The carbon particles were not

observed to spread diffusely through the interstitium of the nasal submucosa. The images

illustrated in Figures 3-1C, D) would support this concept. For technical reasons (as described in

Materials and Methods in chapter 3) we found it difficult to assess Microfil distribution with

histological methods in the adult rats. Consequently, in this report we included images from 9

day-old postnatal pups (Figure 3-1 D). However, in non-processed tissues (Figure 3-1C)

Microfil particles were noted within discrete lymphatic vessels but were not commonly

observed in the surrounding tissues.

A second limitation of using CSF tracer concentrations in the olfactory turbinates to reflect

lymphatic uptake, relates to the nature of the measurements themselves. Ideally, we would like

to quantify the total mass of CSF tracer that had entered the olfactory submucosa over time. For

example, we could have estimated the mass of tracer in each tissue by multiplying the cpm/gm

by tissue weight. However, this approach would have been misleading. Over time, CSF

transports from the subarachnoid compartment into the turbinate tissues and is conveyed

ultimately to the venous system. At any given time, the amount of tracer in the turbinates would

simply represent the tracer that was ‘caught’ in transit through this tissue and would in no way

reflect the total mass of tracer that had transited from the CSF space through this intermediate

compartment on its passage to venous blood.

6.1.1.2 Physiological characteristics of tracer movement across the cribriform plate

Based on the available data, the most important connections between the subarachnoid

compartment and extracranial lymphatics appear to exist at the level of the cribriform plate.

Lymphatic vessels within the olfactory submucosa form a distinctive association with the

72

olfactory nerves and this anatomical relationship serves to facilitate CSF removal from the

subarachnoid space (Johnston et al., 2004;Kida et al., 1993).

Once instilled in ventricular CSF, the radioactive albumin entered the olfactory turbinates

rapidly with peak concentrations being achieved 30 minutes after injection. At this time,

radioactivity in the middle turbinates was nearly 6 times that of blood. After 30 minutes,

turbinate tracer concentrations declined but levels of 125I-HSA were always greater than those

monitored in blood and other tissues. Presumably, the relatively small radioactive signal in the

skeletal muscle, spleen, liver, kidney and tail simply represents the presence of the tracer within

the blood of these tissues.

The rapid appearance of albumin in the turbinates is consistent with relatively quick CSF

disappearance rates from the brain measured in other studies. For instance, the half-lives in CSF

of soluble amyloid β peptide, sucrose or PEG4000 were ~8 minutes, ~60 minutes and ~60

minutes respectively (Ghersi-Egea et al., 1996a;Ghersi-Egea et al., 1996b). About 80% of IGF-

1 was cleared from the brain 30 minutes after injection into the ventricles (Nagaraja et al.,

2005). It is of interest to note that the rapid removal of IGF-1 was not temporally related to its

appearance in blood. The authors speculated that the delay in plasma appearance might have

been due to the entry of IGF-1 into the lymphatics with some time required for transport to

blood through the cervical lymphatic vessels. Additionally, experiments in rats using X-ray

microscopy have indicated that the X-ray contrast medium reached the cribriform plate about 7

minutes after installation in the cisterna magna followed by the appearance of the agent in the

nasal cavities (Brinker et al., 1997). In this same study, the authors noted that Indian ink

similarly administered, reached the cervical lymph nodes 20 minutes after injection.

The appearance of the tracer in blood is of course expected since all CSF removed from the

cranium is ultimately transported to the bloodstream either through a lymphatic/cribriform plate

route, drainage through the perineurial spaces of other cranial/spinal nerves and/or clearance

73

across the arachnoid villi and granulations into the cranial veins. The initial 'spike' in blood

tracer concentrations may have been due to direct transport into the cranial veins, as the

intracranial pressure would be expected to increase temporarily due to the tracer injection into

the ventricle. In past studies, we observed some transient direct CSF-cranial venous transport

when ICP was abruptly elevated to high levels (Papaiconomou et al., 2004;Zakharov et al.,

2004b).

However, the 10-minute delay in the peak concentration of tracer in blood (40 min)

compared to the turbinates (30 min), seems consistent with the view that a significant amount of

the tracer made its way into blood after first traversing the lymphatic network in the olfactory

turbinates, moving through the cervical lymphatics and finally transporting into the venous

system at the base of the neck. This is supported by the data that indicates high levels of CSF

tracer in the lymph nodes in the neck region (Figure 3-2). Presumably the leveling off of blood

tracer concentrations reflects the balance between the declining tracer entry from lymphatics

(and possibly other sources) and tracer removal from blood as albumin disappears continuously

from the vasculature by filtering into the myriad non-central nervous system interstitial

compartments throughout the body.

6.1.2 Lymphatic absorption in kaolin model of hydrocephalus

With the quantitative method to access CSF absorption, we were able to assess lymphatic CSF

absorption in a kaolin-induced rat model of communicating hydrocephalus. Our data showed

that the transport of a CSF across the cribriform plate was reduced significantly in this

hydrocephalus model.

Based on assessments with MRI, the injection of kaolin into the basal cisterns induced a

communicating form of hydrocephalus in the majority of animals. This model appears to

replicate neonatal or infantile forms of hydrocephalus in that ventriculomegaly developed within

74

days of induction and moderate to severe levels of enlargement were observed in most animals.

Kaolin administration in adult animals with fixed skulls does not normally produce rapid or

severe ventricular expansion; rather, such progression is usually found only in young animals

with expandable skulls (Hale et al., 1992;McAllister et al., 1991).

The relationship between a CSF absorption deficit and ventricular expansion is no doubt,

very complex. Figure 4-4 B demonstrates an unusual non-linear relationship between lymphatic

function and ventricular volume. It would seem that bulk flow could be perturbed significantly

before there was any evidence of ventricular expansion. Indeed, from the graph it would appear

that almost all of the lymphatic bulk flow capacity must be compromised before significant

hydrocephalus develops. This implies that the CSF absorption system has significant excess

capacity and that pathology does not develop until this capacity is largely consumed. Perhaps

there is a critical breakpoint in the bulk flow-ventricular volume relationship that has not been

appreciated in the past.

It seems likely however, that the host can afford to lose a certain (unknown) number of CSF

absorption sites as presumably, other locations or mechanisms can compensate up to a certain

extent. The absorption of CSF from the spinal subarachnoid compartment would be a good

example (Bozanovic-Sosic et al., 2001). In support of this, we noted previously that CSF

outflow resistance increased when CSF transport through the cribriform plate was obstructed

(Silver et al., 2002) but this increase was much more evident when CSF absorption from the

spinal cord was prevented (Mollanji et al., 2001;Papaiconomou et al., 2002). As more and more

CSF absorption sites become blocked, the diminished CSF absorptive capacity would be

reflected by reduced CSF (and thus tracer) clearance to plasma. As can be observed in Figure 4-

4 A (inset), the average blood levels of the tracer were slightly higher than in the saline group

although not significantly so. This suggests that other pathways might have compensated for the

diminished absorption at the cribriform plate.

75

One possibility in this regard is the arachnoid villi and granulations. While a role for these

elements in CSF absorption under normal conditions is increasingly being challenged, there is

evidence to suggest that CSF may be transported from the subarachnoid space into the cranial

veins at high intracranial pressures (Papaiconomou et al., 2004;Zakharov et al., 2004b).

Additionally, if we assume that the arachnoid projections have functional significance, it is

theoretically possible that their role was diminished after kaolin administration. However, in this

regard, it is interesting to compare the impact of kaolin injection into the basal cisterns with the

effects of kaolin administration over the cerebral convexities. In both cases, the ventricles

expanded but the ventriculomegaly associated with convexity injections was less severe and

much more protracted, requiring 3-4 months to develop compared to the ventriculomegaly

produced by basil cistern injections which developed quickly (Li et al., 2008). With this in

mind, we think it unlikely that CSF absorption through the arachnoid projections was

compromised in the studies reported here.

From a lymphatic perspective, we can obtain some idea of the impact of reduced CSF

transport through the cribriform plate from studies in which the plate has been sealed on the

extracranial surface. These reports indicate that experimental obstruction of the cribriform plate

reduces cranial CSF absorption (Papaiconomou et al., 2002;Mollanji et al., 2001), elevates

cranial CSF outflow resistance (Silver et al., 2002) and increases intracranial pressure (Mollanji

et al., 2002). However, it is unclear how impaired lymphatic function may contribute to

ventricular expansion. One might expect that intracranial pressure would rise with any

obstruction to CSF outflow. Indeed, previous studies have indicated that ICP is elevated when

the cribriform plate is sealed (Mollanji et al., 2002). However, it is not apparent how disruption

of CSF transport through the cribriform plate would lead to a transmantle pressure gradient

favoring enlargement of the ventricles since CSF pressure would likely rise equally in all CSF

spaces within the cranium. One possibility is that impaired lymphatic CSF uptake could affect

76

compliance and cause a redistribution of pulsatility within the cranium. One group has

postulated that this could lead to ventriculomegaly (Egnor et al., 2002) but this concept will

need to be tested in future experiments looking at changes in pulsatility in the kaolin model.

Further evidence suggesting some relationship between hydrocephalus and lymphatic

function comes from mouse experiments in which intrathecal injections of TGF-β1 were used to

induce hydrocephalus. In these studies, ink was administered into the lateral ventricles and the

time taken for the ink to stain the cervical lymph nodes was considerably lengthened compared

to the non-hydrocephalic animals (Moinuddin & Tada, 2000;Tada et al., 2006). This suggested

that the cribriform-lymphatic connection is also disrupted in the TGFß1-infused hydrocephalus

model.

Therefore, in two different models of hydrocephalus, there is evidence for a suppression of

CSF transport into extracranial lymphatic vessels. Nevertheless, many questions remain. The

lymphatic absorption defect could be due to restricted access of CSF to absorption sites at the

base of the brain, some blockage in the foramina of the cribriform plate or the impact of kaolin

directly on the lymphatic vessels themselves.

It is possible that kaolin particles were dislodged from the original injection site and

migrated to other areas to form multiple foci of obstruction (fibrosis) or inflammation.

Certainly, collagen deposits have been observed in other rat models of hydrocephalus. When

FGF-2 is introduced into the ventricular system of rats, an ex vacuuo type of hydrocephalus

develops characterized by elevations in CSF pressure and CSF outflow resistance (Johanson et

al., 1999). These changes were attributed in part, to fibrosis in the arachnoid membrane. Along

these lines, a kaolin-induced inflammatory response might generate pro-inflammatory

substances that elicit fibrosis in the subarachnoid space or in the cribriform plate foramina. If

this were the case, CSF movement into extracranial lymphatics might be compromised.

However, kaolin deposits in the area of the olfactory bulb and cribriform plate were never

77

observed in gross post-mortem examinations of the animals used in this study, and thus we do

not believe that kaolin had a direct effect on the foramina of the cribriform plate or on the nasal

lymphatics themselves.

Venous hypertension has been postulated as a contributing factor in hydrocephalus

development (Portnoy et al., 1994). It has been assumed that an increase in intracranial venous

pressure would alter the CSF-venous pressure gradient and thus compromise CSF absorption

through the arachnoid granulations and villi (Jones & Gratton, 1989). Assuming that the venous

hypertension in the cranium would be reflected to the veins in the base of the neck, this

phenomenon could also theoretically, reduce lymphatic CSF transport since lymph flow would

have to overcome the elevated outflow pressures expressed in the venous drainage basis.

However, water is removed from lymph as it transits from the base of the brain to the veins

located at the base of the neck (Koh et al., 2007). This tends to ‘depressurize’ the lymphatic

system and reduce the impact of elevated inflow to the vessels or increased outflow resistance.

Therefore, while we do not know if venous pressures are elevated in the kaolin model, we think

it unlikely that venous hypertension will have any long-term impact on the transport of CSF into

the lymphatics associated with the olfactory turbinates.

78

6.1.3 Outflow resistance in kaolin model of hydrocephalus

Our data demonstrated that an absorption deficit occurs at a discrete anatomical location in a

kaolin-induced model of communicating hydrocephalus in the rat (Nagra et al., 2008). At

present, we do not know the mechanism by which lymphatic CSF uptake is impaired.

The lymphatic CSF absorption deficit appears to correlate with ventricular volumes.

However, it is possible that the impediment to lymphatic CSF transport may be a local

phenomenon with CSF clearance increasing at other absorption sites to compensate, such that

global CSF absorption is relatively unchanged. These sites could include other lymphatic

vessels, absorption from the spinal subarachnoid space into peri-spinal lymphatics, drainage

through the arachnoid projections into the superior sagittal sinus or clearance via the capillary

system. If marked compensation were to occur through these other pathways, we would expect

that CSF outflow resistance would be close to normal in the kaolin group. However, we

observed that outflow resistance was elevated significantly suggesting that the impediment to

lymphatic CSF absorption had an impact on global CSF drainage.

This conclusion is supported by several other considerations. First, we know that the

movement of CSF through the cribriform plate into nasal lymphatic vessels accounts for at least

50% of global CSF absorption in the rat (Boulton et al., 1999). Second, in sheep, we also know

that about 25% of absorption occurs from the spinal subarachnoid compartment and there is

every reason to believe that the majority of this transport is via peri-spinal lymphatic vessels

(Bozanovic-Sosic et al., 2001). While not measured in our study, CSF removal from the spinal

subarachnoid compartment in the kaolin rat model may have been compromised as well. Third,

as mentioned before in sheep in which a similar proportion of CSF absorption occurs through

this pathway, sealing the cribriform plate on the extracranial surface leads to reduction in CSF

absorption (Papaiconomou et al., 2002;Mollanji et al., 2001), elevation of intracranial pressure

(Mollanji et al., 2002) and a significant increase in CSF outflow resistance (Silver et al., 2002).

79

Taken together, the data suggest that the kaolin-induced reduction in lymphatic CSF uptake

plays a major role in the increased outflow resistance.

Literature values for CSF outflow resistance measurements in the rat vary considerably.

Values from 0.63 to 2.4 cm H2O.µL-1.min have been reported (Jones et al., 1987a;Mann et al.,

1978;Meier et al., 2002;Kiefer et al., 2000;Kondziella et al., 2002;Botel & Brinker,

1994;Luedemann et al., 2002) and these are greater than the average value we report in this

study (0.18 ± 0.03 cm H2O.µL-1.min) in intact animals. However, it is difficult to compare

values due to methodological differences. Nonetheless, in relative terms, our results agree with

previous studies showing elevated outflow resistance with hydrocephalus induction (Luedemann

et al., 2002;Kondziella et al., 2002;Kiefer et al., 2000;Meier et al., 2002).

80

PART B

CHAPTER 7:

ROLE OF CELL β1-INTEGRIN - MATRIX INTERACTIONS IN GENERATING A

TRANSPARENCHYMAL GRADIENT FAVOURING THE DEVELOPMENT OF

HYDROCEPHALUS

81

7.1 Abstract

In some tissues, the injection of antibodies to the β1 integrins leads to a reduction in interstitial

fluid pressure indicating an active role for the extracellular matrix in tissue pressure regulation.

If perturbations of the matrix occur in the periventricular area of the brain, a comparable

lowering of interstitial pressures may induce transparenchymal pressure gradients favoring

ventricular expansion. To examine this concept, we measured periventricular (parenchymal) and

ventricular pressures with a servo-null micropipette system (2 µm tip) in adult Wistar rats before

and after anti-integrin antibodies, or IgG/IgM isotype controls were injected into a lateral

ventricle. In a second group, the animals were kept for 2 weeks after similar injections and upon

sacrifice, the brains were removed and assessed for hydrocephalus. In experiments in which

antibodies to β1 integrins were injected (n=10) but not isotype control IgG/IgM (n=7), we

observed a decline in periventricular pressures relative to the pre-injection values. Under similar

circumstances, ventricular pressures were elevated (n=10) and were significantly greater than

those in the periventricular interstitium. We estimated ventricular to periventricular pressure

gradients of up to 4.3 cm H2O. In the chronic preparations, we observed enlarged ventricles in

many of the animals that received injections of anti-integrin antibodies (21 of 29 animals; 72 %)

but not in any animal receiving the isotype controls. We conclude that modulation/disruption of

β1 integrin-matrix interactions in the brain generates pressure gradients favoring ventricular

expansion suggesting a novel mechanism for hydrocephalus development.

82

7.2 Introduction

It is not clear how impaired CSF clearance could lead to a dilation of the ventricles since the

ventricular and subarachnoid compartments are in communication with on another and pressure

would likely increase in both compartments equally if the outflow to CSF was obstructed. It is

of course possible that very small transmantle pressure gradients (1 mm Hg or less) induced by

some impediment to CSF flow expand the ventricles as postulated by Levine (Levine, 2008).

Levine has argued that the pressure gradients favouring ventricular expansion are very small

since pressures are diminished towards the periphery of the brain due to the absorption of

interstitial fluid into the brain capillaries. The capillary absorption of water becomes a critical

element in his mathematical formulation. If Levine’s suppositions are correct, then a decline in

lymphatic CSF absorption with a concomitant increase in global CSF outflow resistance could

play an important role in hydrocephalus development. However, the postulated small gradient of

pressure has never been verified directly. In addition, while hydrocephalus is associated with

elevations in CSF outflow resistance, the opposite is not necessarily true as elevations in CSF

outflow resistance (inferring reductions in CSF absorption) do not always correlate with

hydrocephalus. In the disease pseudotumor cerebri for example, CSF outflow resistance is

higher than normal without hydrocephalus being present (Wall, 2008).

Furthermore, in previous studies from our group, it was clear that a lymphatic CSF

absorption deficit occurred in ageing rats and yet no hydrocephalus was present in these animals

(Nagra & Johnston, 2007). In additional studies in 3 sheep, Dr. Johnston’s group sealed the

cribriform plate and even though that this procedure elevates intracranial pressure and CSF

outflow resistance in this species (Mollanji et al., 2002;Silver et al., 2002), we did not observe

any ventricular expansion over 3 months (unpublished observations). In conclusion, it is not yet

apparent whether an impediment to CSF drainage represents a pivotal event in hydrocephalus

development or whether it is a ‘co-conspirator’ or a secondary finding in the pathogenesis of

83

ventricular enlargement with some other factor denoting the definitive cause. In this regard it is

of interest to note that kaolin is known to have an inflammatory effect (Wall, 2008;Rubin et al.,

1976). This introduces another factor into the hydrocephalus equation.

7.3 An alternative explanation for ventricular expansion

Dr. Pickard’s Cambridge group has developed an interesting sidebar to the question of

hydrocephalus. Based on a mathematical model employing finite element analysis and poro-

elastic concepts, these authors postulated that ventricular expansion may result from a relative

reduction in interstitial fluid pressure in the periventricular area leading to the formation of an

intra-mantle rather than a transmantle pressure gradient (Pena et al., 2002). No mechanism that

could cause this phenomenon was identified or proposed but studies carried out in non-CNS

tissues may provide a clue as to how this might occur. Rubin and colleagues have proposed that

fibroblasts in the skin regulate interstitial fluid pressure by exerting a tensile force on matrix

elements, which restrains the interstitial gel from swelling. Various types of inflammatory

reactions, certain prostaglandins, and Cytochalasin D can induce a lowering of interstitial fluid

pressure. The data with the F actin-disrupting agent Cytochalasin D supported a role for

extracellular and intracellular cytoskeletal linkages in pressure regulation (Berg et al., 2001).

These factors appear to regulate the balance between grip and release by altering matrix-integrin

interactions leading to compaction or tissue swelling which in turn affects interstitial pressure

(Rubin et al., 1995). This phenomenon appears to occur in the trachea as well (Woie et al.,

1993). Furthermore, injections of anti β1 integrin antibodies into skin simulated the

inflammatory effects by inducing a significant reduction of interstitial fluid pressure (Wiig et

al., 2003). Indeed, the anti-inflammatory effects of alpha-trinositol appear to relate to its ability

to modulate β1 integrin function with a concomitant reversal of the interstitial pressure lowering

effect (Rodt et al., 1994). Therefore, integrin-matrix interactions appear to be prime candidates

84

for regulating this phenomenon. We may then ask if the brain parenchyma can modulate tissue

pressure similarly.

7.4 Objective of Part B of this Thesis

As a first step, we decided to test the hypothesis that the intraventricular injection of anti β1

integrin antibodies in rats will lower periventricular interstitial fluid pressure relative to that

measured in the ventricular system and induce ventricular enlargement.

85


7.5.1 Animals

All experiments were approved by the animal ethics committee at Sunnybrook Health Science

Centre. Additionally, all studies conformed to the guidelines set by the Canadian Council on

Animal Care and the Animals for Research Act of Ontario. A total of 31 adult Wistar rats (208-

274 grams) were used for this investigation (Charles River, Canada).

7.5.2 Surgical procedures for pressure measurements

Rats were anesthetized initially by placement in a custom-built rodent anesthesia chamber using

isofluorane (5%) in oxygen. For the experimental procedure they were maintained with 2-2.5%

isofluorane in oxygen delivered by a nose cone. The animals were placed on a heated water pad

and the heart rate and oxygenation status were monitored by a pulse oximeter placed on the hind

foot (Nonin 8600V, Benson Medical, Markham, Ont.). The animal was then fixed into position

in a small animal stereotaxic device (KOPF, Model 900, Tunjunga, Calif., USA). All stereotaxic

coordinates were taken from a rat brain atlas (Paxinos & Watson, 2007).

The skin over the cranium was removed and the junction of the sagittal and coronal sutures

identified using a stereomicroscope (Carl Zeiss OPMI 1-FC). A small high-speed microdrill

with a rounded tip (Fine Science Tools, Vancouver, Canada) was used to grind away the bone to

expose the dural membrane at two locations; 0.92 mm posterior/1.4 mm lateral right to the

bregma and 1.7 mm anterior/1.6 mm lateral right to the bregma. The former hole was used for

the injection of anti-β1 antibodies into the lateral ventricle and the latter hole was used for the

insertion of the micropipette into the parenchymal interstitium for pressure measurement. For

ventricular pressure measurements, 2 holes were made at 0.92 mm posterior and 1.4 mm left and

86

right lateral to the bregma for the antibody injection and ventricular pressure tip insertion

respectively.

7.5.3 Measurement of pressures using the Servonull System

Pressures were measured using a servo-nulling pressure measuring system (Vista Electronics,

Ramona, CA). Borosilicate glass capillaries (World Precision Instruments, Sarasota, Florida) 1

mm OD and 0.75 mm ID were pulled with a two-step pipette puller (Narshige PP-830 Puller,

East Meadow, NY) to yield a tapered tip that had a 2 µm diameter. A pipette grinder (Narshige

EG-400 Micropipette Grinder, East Meadow, NY) was used to grind the pipette tips to a 40˚

angle to facilitate tissue penetration. Each tip was examined under the microscope to ensure

uniform tip diameters. The micropipettes were then filled with a 1 M NaCl solution using a

nonmetallic syringe needle (Microfil MF34G, World Precision Instruments, Sarasota, Florida).

For the experiment, the micropipette was connected to a pipette holder, which was mounted

on a micromanipulator (MM-33B Precision Micromanipulator, Fine Science Tools, North

Vancouver, BC). The servo-null system was calibrated to zero before the micropipette puncture

by immersing the tip in a pool of Ringer’s solution at the site of the hole in the cranium. For

parenchymal pressures, the micropipette was then advanced 3.0 mm beyond the dura into the

corpus callosum. For intraventricular pressures, the micropipette tip was inserted at a depth of

3.4 mm with respect to the dura. Figure 7-1 illustrates the location of the micropipettes relative

to the lateral ventricles. The pipette tip was approximately 500-600 µm from the anterior horn of

the lateral ventricle.

After achieving a stable reading, the antibodies were injected slowly into one of the lateral

ventricles (approximately 5 µl/second). The parenchymal interstitial pressure was monitored

continuously on a computer-based data-acquisition system (Daq Software; A-Tech Instruments,

Toronto, Canada). All data was collected at a frequency of 10 Hz. Pressure recordings had to

87

meet the following criteria: baseline stability for at least 10 minutes and an unchanged zero of

the system upon withdrawal of the pipette from the tissue and re-immersion in the pool of

Ringers lactate.

Figure 7-1 Sections of a rat head illustrating the position of the micropipette tip used to

measure periventricular interstitial fluid pressures (asterix in circle).

(A) Sagittal section, (B) Axial section. For these illustrations, Evans blue dye was injected into

the cisterna magna to aid in the visualization of the ventricles. The inset in B illustrates the

location from which a block of tissue was removed for Western blot analysis.

88

7.5.4 Antibody injection for pressure measurements

Antibodies against β1, α2β1 integrins or non-immune isotype controls (IgG and IgM) were

injected into a lateral ventricle (outlined in Table 7-1). For injections, a 250 µL Hamilton

syringe (Fisher Scientific, Toronto, Canada) with a 27-gauge needle was used. The syringe

needle was loaded with 50 µL of the antibody solution and the needle tip lowered into one of the

lateral ventricles 3.4 mm deep from the dura. The mass of protein injected was as follows: anti

β1 Integrin IgG (10 μg), anti β1 Integrin IgM (50 μg), anti α2β1 Integrin IgG (25-50 µg), rabbit

control IgG (10 µg), purified hamster IgM (25 µg), rat control IgG (20 µg).

7.5.5 Chronic experiments

Rats (56 animals, 206-285 grams) were anesthetized and fixed in position as described earlier.

Lube (Refresh Lacri-Lube, Allergan Inc, Markham, Ont., Canada) was applied to eyes to

prevent dryness and alcohol and iodine was applied to their shaved heads. Also, 0.2 mL of

Duplocillin (intramuscular.), 0.1 mL of Temgesic (subcutaneous) and 4-5 mL of saline

(subcutaneously at the paralumbar fossa) were injected. The skin over the cranium was reflected

and the junction of the sagittal and coronal sutures identified using the stereomicroscope. A

small high-speed microdrill described earlier was used to grind away the bone to expose the

dural membrane at 0.92 mm posterior and 1.4 mm lateral right to the bregma. The hole was used

for the injection of the antibodies or isotype controls into a lateral ventricle (Table 7-1).

A 250 µL Hamilton syringe with a 30-gauge needle was loaded with the injectate and the

needle tip lowered into the lateral ventricle 3.4 mm deep from the dura. The injections (25, 50 or

100 µL) were performed at a rate of about 5 µl/second and the needle retracted 1 minute after

the injection had finished. The hole was covered with bone wax. Bupivacaine hydrochloride

(Sensorcaine, Astra Zeneca, Mississauga, Ont., Canada) was applied in the area of the incision

89

for pain control. The deeper skin layer over the cranium was sutured with polysorb 4-0 sutures

and the more superficial layer closed with 4-0 silk sutures.

The rats were fed lab rat chow (LabDiet 5001) until sacrifice with Euthanyl at 2 weeks post

surgery. At this point, the brains were removed and fixed in 10% formalin. The weight of the

rats immediately after sacrifice ranged between 225-303 grams. A coronal section of the brain

was made 6 mm from the frontal tip of the cortex.

90

Table 7-1

91

7.5.6 Assessment of function blocking status of the anti β1 Integrin IgG antibody

To determine if the anti β1 integrin IgG antibody was a function-blocking antibody (Table 7-1,

#1) we assessed its impact on the integrin mediated adhesion of U937 cells to Vascular Cell

Adhesion Molecule 1 (VCAM-1). U937 cells are a monocytic cell line that expresses

constitutive levels of α4 β1 integrins in a low affinity state. To induce a high affinity

conformation in this system, U937 cells were treated with manganese (Mn++). Cells were

pretreated with the anti integrin antibody (10 µg) or IgG-isotype control (10 µg) prior to

adhesion. Using a parallel plate flow chamber system, cells were then introduced onto VCAM-1

coated tissue culture plates and allowed to adhere for a period of 2 minutes. An incremental wall

shear stress (0, 2, 4, 10, 20 dynes/cm2) was applied and cell adhesion quantified by video

microscopy.

7.5.7 Penetration of immunoglobins into parenchymal tissues

Animals received 100 µL intraventricular injections of the same mass (20 µg) of antibodies

directed against β1 integrins (rabbit IgG, n=3), control isotype rabbit IgG (n=3) or hamster IgG

(n=1) using methods described earlier. Two hours after injection the animals were perfused

transcardially with 80-100 mL of saline, the brains were removed and an approximately 1.5

mm3 block from the cortical areas dorsal to the injection site in the ventricles was dissected. The

location of the tissue harvested (superficial to the location of the pipette tip) is illustrated

schematically in the insert to Figure 7-1 B (circled). The tissues were placed on dry ice before

their storage in an 80C freezer. Brain samples were sonicated in a lysis buffer (10X w/v; 50mM

Tris HCL pH 7.4, 10mM EDTA, 4M Urea, 3% Triton X100, 1X protease inhibitor, Calbiochem,

539131), and 200 µM phosphatase inhibitor sodium vandate.

Immunoblots were performed according to standard procedures. Briefly, the samples (40 µg

total protein) were boiled, centrifuged for 5 minutes, separated by sodium dodecyl sulfate-

92

polyacrylamide gel electrophoresis (SDS-PAGE, 10%) and transferred to nitrocellulose

membranes (BioRad). Membranes were blocked for 1 hour in Tris-buffered saline (TBS)

containing 0.1% Tween and 5% skim milk powder. Following a 1 hour incubation with anti-

rabbit IgG secondary antibodies conjugated to horseradish peroxidase (1:5000; Jackson

ImmunoResearch), the membranes were washed with TBS-tween buffer and the bands revealed

using Immobilon Chemiluminescence (Millipore, WBKL50500). To assure that equal amounts

of protein were loaded in each lane, the blots were stripped and reprobed with an antibody

against glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (1:2000; Biodesign, Saco, ME)

for 1 hour followed by a horseradish peroxidase conjugated antibody and chemiluminescence

detection of the GAPDH bands.

7.5.8 Data Analysis

Pressure Data. To normalize the pressure values, the data were averaged over 120 second

intervals. Subsequently, a baseline value was derived for each trace by averaging a minimum of

10 minutes of a stable pressure recording immediately prior to the injection time. Each post-

injection data point was expressed as a ratio by dividing each point by the average baseline

value.

Cell adhesion data. These data were normalized by expressing each point as a percentage

with respect to total number of adherent cells at 0 shear stress.

All averaged data are represented as the mean ± SE. The data has been assessed with the

Student's t-test, ANOVA or repeated measures regression analysis adjusted for within

correlations over time. We interpreted P<0.05 as significant.

93

7.6 Results

7.6.1 Acute experiments

Parenchymal pressures. In 8 out of 10 cases, we noted a bolus injection effect of variable

magnitude in the periventricular interstitium when IgG (n=5) or IgM antibodies to the β1

integrins (n=5) were injected into the ventricles. However, following this increase in interstitial

fluid pressure, in every case the parenchymal pressures declined below baseline at some point

during the experiment. In 7 of 10 cases, the pressures declined steadily after antibody injection

(example with IgG anti-β1 integrin antibody in Figure 7-2 A). In 3 of 10 examples, the pressures

fell below baseline rapidly, and then returned to pre-injection levels (example with IgG anti-β1

integrin antibody illustrated in Figure 7-2 B). In 2 of 10 cases, no bolus effect was observed

with pressures decreasing immediately after injection (example with IgM anti-β1 integrin

antibody illustrated in Figure 7-2 C). In addition to the aforementioned studies, we performed

experiments using antibodies to α2β1 integrin (n=3). In each case, pressures declined below pre-

injection levels (data not illustrated). It is of interest to note that in all experiments in which the

appropriate isotype controls were injected into a lateral ventricle (n=7), intra-parenchymal

pressures increased above pre-injection values (example with isotype Hamster IgM in Figure 7-

3 A).

Ventricular pressures. Following injections of the anti-integrin antibodies (IgG based, n=5;

IgM based, n=5), a bolus pressure increase of variable magnitude was observed in the ventricles.

In some cases, pressures returned to baseline, however on average, pressures remained above

pre-injection levels over the course of the experiment (example after injection of Anti β1 integrin

IgG antibody in Figure 7-3 B). A similar pattern was observed after injection of the isotype

rabbit IgG (n=2) or hamster IgM (n=2) controls (an example after injection of control isotype

hamster IgM is illustrated in Figure 7-3 C).

94

C

B

A

Figure 7-2

95

Figure 7-2

Examples of periventricular interstitial fluid pressures. Times of injection are illustrated

with arrows. The dashed line represents baseline pressure.

A) Injection of anti-β1 integrin antibody (IgG; number 1 in Table 1). In this example, pressure

increased transiently due to the bolus injection effect and then declined steadily below the pre-

injection level.

B) Injection of anti β1-integrin antibody (IgG; number 1 in Table 1). After the transient bolus

effect, pressure declined for a period and then increased to approximately pre-injection levels.

C) Injection of anti β1-integrin antibody (IgM; number 2 in Table 2). In this example, there was

a minimal bolus effect and immediately, periventricular pressures fell below baseline values.

96

A

B

C

Figure 7-3

97

Figure 7-3

Examples of periventricular and intraventricular pressures. Times of injection are

illustrated with arrows. The dashed line represents baseline pressure.

A) Periventricular pressure following injection of control isotype hamster IgM (number 5 in

Table 1). In this example there was essentially no bolus injection effect with pressures rising and

then reaching a plateau. Pressures remained above baseline for the duration of the experiment.

B) Intraventricular pressure after injection of anti β1-integrin antibody (IgG; number 1 in Table

1). There was a large bolus effect and pressures increased steadily over the course of the study.

C) Intraventricular pressure following injection of control isotype Hamster IgM (number 5 in

Table 1). A marked bolus injection effect was observed and pressures remained considerably

elevated.

98

7.6.1.1 Pressure analysis

In every experiment in which IgG or IgM antibodies to the β1 integrins were injected, interstitial

fluid pressures declined below baseline. Therefore, we pooled the results for the ventricular

(n=10) and parenchymal tissue fluid pressures (n=10). We also averaged the data from all the

isotype control injections for parenchymal (n=7) and ventricular pressures (n=4). The averaged

data are illustrated in Figure 7-4.

7.6.1.2 Parenchymal pressures following injection of anti β1 Integrin antibodies or their

isotype controls (Figure 7-4 A)

Approximately 20 minutes after injection of the anti β1 integrin antibodies, the parenchymal

pressures decreased below pre-injection (baseline) levels and remained below baseline for the

duration of the experiment. By the end of the monitoring period, interstitial fluid pressure had

declined to around 70% of the baseline value (solid circles). In contrast, the parenchymal

pressures associated with the injection of the IgG/IgM isotype controls were elevated above

baseline and remained so for the duration of the study (open circles). Pressures were

significantly different between the 2 groups (p<0.0001; repeated measures regression analysis).

7.6.1.3 Comparison of ventricular and parenchymal pressures after the injection of

antibodies to β1 Integrins (Figure 7-4 B)

The pressures measured in the lateral ventricles induced with injections of the specific anti β1

Integrin antibodies (open circles) remained above baseline during the experiments and were

significantly greater than pressures monitored in the periventricular interstitium (p=0.038;

repeated measures regression analysis). This indicated a pressure gradient between the ventricles

and surrounding tissues.

99

7.6.1.4 Comparison of ventricular pressures after injections of anti β1 Integrin antibodies

or their isotype controls (Figure 7-4 C)

Ventricular pressures associated with the isotype control injections were higher than those

induced by the specific antibodies to the β1 Integrins. These differences were significant

(p=0.0002; repeated measures regression analysis).

7.6.1.5 Comparison of pressures in the ventricles and parenchyma after injection of the

IgG/IgM isotype controls (Figure 7-4 D)

Pressures in the ventricles and parenchyma following the injection of the isotype controls were

not significantly different (p=0.067; repeated measures regression analysis).

100

Figure 7-4 Comparison of the averaged pressures following injection of antibodies or

isotype controls. The normalization of the data and the method for averaging have been

described in the Materials and Methods. Injections occurred at time 0.

A) Parenchymal interstitial fluid pressures following the injection of the IgG/IgM isotype

controls (open circles, n=7) or the antibodies to β1 integrins (closed circles, IgG/IgM, n=10).

Pressures associated with the administration of the specific anti β1 integrin antibodies were

significantly less than those associated with the injection of the isotype controls (p<0.0001).

B) Ventricular (open circles, n=10) and parenchymal pressures (closed circles, n=10) after

injection of the antibodies to β1 integrins. Pressures in the periventricular interstitium were

significantly lower than those monitored in the ventricles following the administration of the

specific anti β1 Integrin antibodies (p=0.038).

C) Ventricular pressures after injecting antibodies to the β1 Integrins (open circles, n=10) were

significantly less (p=0.0002) than those observed after the injection of the isotype controls

(closed circles, n=4).

D) Pressures in the ventricles (n=4, closed circles) and parenchyma (n=7, open circles)

following the injection of the isotype controls were not significantly different (p=0.067;

repeated measures regression analysis).

102

7.6.1.6 Pressure gradients

The data from Figure 7-4 can be used to estimate potential pressure gradients between

ventricular CSF and periventricular interstitial fluid under the conditions of our experiments.

The average baseline (pre-injection) ventricular and parenchymal pressures in these studies were

6.6 ± 0.5 cm H2O and 5.6 ± 0.5 cm H2O respectively (not significantly different; students t-test).

Averaging all baseline pressures we obtain 6.0 ± 0.3 cm H2O which is only slightly higher than

the normal servonull-based values of CSF pressure noted in the studies of Wiig (3.4 mm Hg or

4.7 cm H2O) (Wiig & Reed, 1983). Taking the data from Figure 7-4B, at 24, 48 and 72 minutes

after injection of the specific anti β1 integrin antibodies, the average ventricular pressures had

increased 24, 27 and 42% above baseline. In contrast, the average parenchymal pressures at the

same times had declined to 88, 80 and 70% of baseline levels. Therefore, we can estimate

average ventricular pressures of 7.4, 7.6 and 8.5 cm H2O and parenchymal pressures of 5.3, 4.8

and 4.2 cm H2O at the 3 times. This would give theoretical ventricle to parenchymal pressure

gradients of 2.1, 2.8 and 4.3 cm H2O at the various times after injection. Using a similar

approach we estimated the average parenchymal pressure differences caused by injection of

specific anti-integrin antibodies relative to the isotype controls (data from Figure 7-4A). At 24,

48 and 72 minutes, the pressures induced with the specific anti-integrin antibodies were 4.3, 3.6

and 4.8 cm H2O lower than those induced with the appropriate IgG/IgM controls.

7.6.2 Chronic experiments

Over the course of the 2-week period following the injection of specific antibodies or isotype

controls, we did not notice any obvious behavioral or weight changes that might be indicative of

hydrocephalus development. However, on post-sacrifice analysis, we observed hydrocephalus in

103

many of the animals that received antibodies to the β1 integrins. Table 7-2 summarizes the

results of these experiments. If we consider the results from injections of the antibodies to β1

Integrins (IgG and IgM) and to α2β1 Integrin, 21 out of 29 animals developed some degree of

ventriculomegaly. The magnitude of hydrocephalus was considerably greater in the animals that

received IgG antibodies to β1 or α2β1 integrins in comparison to those having been injected with

the IgM antibodies (examples provided in Figure 7-5 C-F). Indeed, the IgM anti β1 integrin

antibodies induced only minimal expansion. In contrast, none of the animals injected with

isotype IgG/IgM controls exhibited hydrocephalus (examples in Figure 7-5 A, B).

104

Table 7-2

105

Figure 7-5

Coronal sections of rat brains for

assessment of hydrocephalus 2

weeks after antibody/isotype

control injection into a lateral

ventricle. A ruler in 1 mm

increments is illustrated in each

image.

A) Rabbit IgG isotype control; 50

µL (number 4 in Table 1). No

ventricular enlargement is evident.

B) Hamster IgM isotype control; 50

µL (number 5 in Table 1). No

ventricular enlargement is evident.

C) Anti-β1 integrin antibody (IgG);

50 µL (number 1 in Table 1).

Ventricles are expanded

considerably.

D) Anti-β1 integrin antibody (IgG);

100 µL (number 1 in Table 1).


considerably.

E) Anti-β1 integrin antibody (IgM);

50 µL (number 2 in Table 1). A

minor increase in ventricle size was

noted.

F) Anti-α2β1 integrin antibody

(IgG); 50 µL (number 3 in Table 1).


considerably.

106

7.6.3 Assessing the characteristics of the IgG anti-β1 integrin

The functional status of the IgG anti β1 integrin antibody (listed #1 in Table 7-1) was not

apparent from the manufacturer's data sheet. As this antibody induced marked hydrocephalus in

many of the rats and lowered parenchymal interstitial fluid pressure, we suspected that it was a

functional blocking antibody with respect to integrin function. To investigate this issue using an

independent measure, a cell adhesion assay was employed. Adherent cells were counted in a

flow system that permitted the application of variable shear stress forces on the cells (Figure 7-

6). We observed that the percentage of adherent U937 cells treated with the anti β1 antibody was

significantly less than that noted with the isotype-treated control (2 factor ANOVA, Group X

Shear stress; p< 0.0001). This result provided additional evidence that the IgG anti-β1 integrin

antibody was function blocking with respect to the β1 integrins.

107

Figure 7-6 Adhesion of U937 cells to VCAM-1.

Manganese alone (open circles, n=4), Manganese + istotype control IgG (closed circles, n=4),

and manganese + antibodies to β1 integrin (closed triangles, n=4) at incremental wall shear

stresses (dynes/cm2). ANOVA revealed that the introduction of the β1 integrin antibodies

impaired adhesion significantly relative to the isotype control (p< 0.0001) indicating that this

antibody has function blocking abilities.

108

7.6.4 Penetration of immunoglobulins into the peri-ventricular parenchyma following

intraventricular administration

Our servonull pressure experiments implied that the antibodies to the β1 integrins penetrated at

least 500-600 µM from the anterior horn of the lateral ventricle (position of the micropipette tip)

since we observed a pressure effect after injecting the antibodies. Additionally, in the chronic

studies some of the ventricles seemed to expand beyond the pressure measurement point

suggesting that the antibodies may have penetrated even further into the parenchyma. This

supposition was supported by preliminary experiments.

Western blotting was performed on brain samples from animals injected with rabbit anti β1

integrin, isotype control rabbit IgG and anti-hamster IgG (example in Figure 7-7 A-C). A 50

kDa band was detected in animals injected with anti β1 integrin IgG (7-7A) and the isotype

control IgG (7-7 B) but not following the administration of anti-hamster IgG antibodies (7-7 C).

These data indicated that the immunoglobulins injected into the lateral ventricles penetrated a

considerable distance into the brain parenchyma. In all experiments, the β1 integrin bands

(example in Figure 7-7 A) were more intense than those exhibited by the isotype rabbit IgG

controls (7-7 B). Whether the antibodies to the β1 integrins were retained in the brain more than

the isotype controls (perhaps due to binding integrins in the tissues) needs further evaluation.

109

Figure 7-7 Penetration of the antibodies into the periventricular parenchyma after

administration into a lateral ventricle.

Animals were injected with rabbit anti β1 integrin antibody (IgG - A), non-immune isotype

rabbit IgG (B) and non-immune hamster IgG (C). Brain samples were taken from the cortex,

dorsal to the injection site. Western blots probed with an anti-rabbit IgG secondary antibody

revealed a 50 kDa band corresponding to the heavy chains of the (A) anti β1 integrin rabbit IgG

and (B) the isotype rabbit IgG control. The non-immune hamster IgG antibody was not detected

(C) demonstrating the specificity of the anti rabbit secondary antibody. Glyceraldehyde 3-

phosphate dehydrogenase (GAPDH) was used as a loading control for all animals with a typical

band at 37 kDa (A-C).

110

7.7 Conclusion

In experiments in which antibodies to β1 integrins were injected (n=10) but not isotype control

IgG/IgM (n=7), we observed a decline in periventricular pressures relative to the pre-injection

values. Under similar circumstances, ventricular pressures were elevated (n=10) and were

significantly greater than those in the periventricular interstitium. We estimated ventricular to

periventricular pressure gradients of up to 4.3 cm H2O. In the chronic preparations, we observed

enlarged ventricles in many of the animals that received injections of anti-integrin antibodies

(21 of 29 animals; 72 %) but not in any animal receiving the isotype controls. We conclude that

disruption of β1 integrin-matrix interactions in the brain generates pressure gradients favoring

ventricular expansion suggesting a novel mechanism for hydrocephalus development.

The content presented in this chapter has been previously published: Nagra G, Koh L, Aubert I, Kim M and Johnston M. Intraventricular injection of antibodies to beta1-integrins generates pressure gradients in the brain favoring hydrocephalus development in rats. Am J Physiol Regul Integr Comp Physiol 297: R1312-R1321, 2009. doi:10.1152/ajpreg.00307.2009

111

CHAPTER 8:

THE BRAIN INTERSTITIUM AND HYDROCEPHALUS

DISCUSSION

112

8.1 Summary of Conclusions Contained in Part B of this thesis It seems unlikely that an absorption defect alone could induce ventricular expansion observed in

hydrocephalus since this defect is unlikely to induce a transmantle pressure gradient. Hence we

searched the literature to find a potential initiator of hydrocephalus. In this regard, studies in

non-CNS tissues have provided a clue as to how this may occur and in doing so, have

transformed our view of the interstitium (Wiig et al., 2003) Rather than simply being a passive

participant, the extracellular matrix assumes an active role in regulating interstitial fluid

pressure. Our data would suggest that matrix components provide a similar dynamic function

within the brain parenchyma. Especially important are the β1 integrins.

In the rat paw, a polyclonal anti-β1 integrin IgG that inhibited fibroblast mediated collagen

adhesion in vitro, lowered the interstitial fluid pressure and also induced edema formation,

similar to that observed in inflammatory reactions (Reed et al., 1992) or following burn injury

(Lund et al., 1988;Lund et al., 1989). Further studies implicated the α2β1 integrins (which bind

collagen and laminin) in interstitial pressure regulation (Rodt et al., 1996). The observation that

the F actin-disrupting agent Cytochalasin D also lowered interstitial fluid pressure, supports a

role for extracellular and intracellular cytoskeletal linkages in pressure regulation (Berg et al.,

2001). In contrast, the pressure effect was not mediated by fibronectin as polyclonal anti-rat

fibronectin IgG and the fibronectin receptor binding protein Arg-Gly-Asp (RGD) had no impact

on interstitial pressures.

Integrins are cell surface glycoproteins that mediate cell-matrix interactions by providing a

physical transmembrane link between the extracellular matrix and the cell cytoskeleton (Hynes,

2002). Rubin and colleagues have proposed that fibroblasts in the skin regulate interstitial fluid

pressure by exerting a tensile force on the collagen matrix, which restrains the interstitial gel

from swelling. The β1 integrins provide the link between the extracellular matrix and the

cytoskeletal contractile apparatus. Integrin function can be modulated by cytokines and other

113

factors that regulate the balance between grip and release leading to compaction or tissue

swelling which in turn affects interstitial fluid pressure (Rubin et al., 1995).

8.2 Discussion of Results

8.2.1 Integrins and matrix elements in the brain

The extracellular matrix of the brain is different from that of the skin. Relatively speaking, in the

brain there are lesser amounts of fibrous proteins such as collagens, fibronectins and vitronectins

(Gladson & Cheresh, 1991;Rutka et al., 1988;Stallcup et al., 1989). Prominent brain matrix

elements include lectican, hyaluronic acid and the tenascin family (Ruoslahti, 1996). At this

point, the implication of the differences in the matrix composition of the brain relative to skin is

unknown. It is possible that the modulation of more brain-specific cell-matrix interactions may

have a similar or even greater effect on hydrocephalus development.

Nonetheless, β1 integrins are believed to have many important roles in CNS function (Milner

& Campbell, 2002) and are expressed on choroidal and ependymal cells and throughout the

neuropil on glial cells and vascular structures (Grooms et al., 1993;Paulus et al., 1993;Del

Zoppo & Milner, 2006). Additionally, astrocyte fibers surrounding select microvessels in the

adult primate express β1 integrins as do the endothelial cells themselves (Del Zoppo & Milner,

2006). Endothelial cells and astrocytes maintain the basal lamina and support the barrier

properties associated with the blood brain barrier. At this point, we do not know if injections of

the anti β1 integrin antibodies affected barrier function. However, if this was the case, one might

expect that the integrity of the barrier would be lost and the interstitial pressure would increase

as fluid and solutes convected from the blood into the parenchymal interstitium. Of course, it is

possible that this effect blunted the reduction in pressure we observed and future experiments

will examine the impact of the antibodies on blood brain barrier function.

The ependymal cells lining the ventricles would have been exposed to the anti β1 integrin

114

antibodies. While these cells appear to provide a metabolic barrier at the CSF-brain interface

(Del Bigio, 1995), it is unlikely they restrict significantly the movement of immunoglobins or

water between the ventricles and peri-ventricular tissues. Nonetheless, the ependymal cell cilia

play a role in CSF homeostasis and deficiency of regulatory factors necessary for ciliogenesis is

associated with abnormal differentiation of ependymal cells and hydrocephalus in mice (Baas et

al., 2006). Therefore, we cannot discount some unknown effect of the antibodies on the

ependymal cell layer.

When β1 integrin expression is ablated specifically in precursors of neuronal and glial cells

using a cre-lox genetic strategy, the mice exhibited a strikingly convoluted cortex and a reduced

brain size (Graus-Porta et al., 2001). This suggested that the β1 integrins had an important role

in determining the structural integrity of the brain. Additionally, disorganization of the cerebral

cortex is observed in humans and mice with mutations in the laminin alpha 2 chain (Miyagoe-

Suzuki et al., 2000). Children with mutations of the LAMA2 gene have reduced or absent

laminin alpha 2 chain. MRIs of these individuals show structural cortex abnormalities and

ventricular dilation (Sunada et al., 1995;Philpot et al., 1999). Since laminin is an important

ligand for the β1 integrins, it is possible that the disruption of the β1 integrin-laminin interactions

may be a contributor to the pressure patterns and hydrocephalus development noted in this

thesis.

In this regard, it is of interest to note that the dystroglycans share laminin as a primary matrix

ligand with a number of β1 integrin receptors (Satz et al., 2008). The dystroglycans are

heterodimeric transmembrane receptors that link the intracellular cytoskeleton of select cells

with the matrix elements. Enlargement of the lateral ventricles was observed in 45% of

dystroglycan null mice (Satz et al., 2008). The authors speculated that this could be due to the

stenosis of the cerebral aqueduct and/or blockage of the arachnoid granulations. However, one

might speculate that the epiblast-specific loss of dystroglycan might affect cell matrix

115

interactions in a way analogous to that observed with the injection of β1 integrin antibodies.

8.2.2 Antibody Issues

The original concept of matrix tissue pressure regulation arose from studies on inflammation

(Reed & Rodt, 1991;Wiig et al., 2003). The antibodies we used were foreign proteins and as

such, might induce some inflammation in the brain. However, we injected three different

antibodies to the integrins and only those specifically directed against β1 or α2β1 had the

pressure/hydrocephalus effect. If a non-specific inflammation were responsible for the pressure

effects observed, we should have seen the effects with all antibody injections including the

isotype controls but this was not the case.

Based on servo-null measurements, we estimated average ventricular to parenchymal

pressure gradients up to 4.3 cm H2O. In this context, it would seem that the antibodies had

passed successfully through the ependyma and had penetrated some distance into the peri-

ventricular tissues to achieve this effect. This conclusion was supported by the molecular

experiments. However, at this point, we have no way of knowing whether the antibody

concentrations we chose were optimal or even if we had targeted the most appropriate matrix

interactions.

In our experience, both the IgG and IgM anti-integrin antibodies reduced parenchymal

pressures approximately the same degree. However, the ventriculomegaly with the anti integrin

IgG antibodies was considerably greater than that observed with the IgM counterparts, which

induced modest ventricular expansion at best. There are a number of possible explanations for

these findings. First, IgM is larger than IgG and it is possible that less of this antibody entered

the parenchyma and consequently, the volume of brain affected may have been smaller.

Additionally, relatively less penetration of the IgM antibodies may have made more available

for CSF absorption via lymphatic or arachnoid CSF drainage mechanisms (Koh et al., 2005).

116

Second, the full time course of the pressure effects of the 2 antibodies is unknown as our

servonull pressure experiments were of relatively short duration. It is possible that the pressure

effects of the IgM anti β1 integrin antibodies were exerted over a shorter period compared with

those of its IgG counterpart. If so, a short-lived pressure gradient may have been insufficient to

permit the development of marked hydrocephalus.

8.2.3 Dissociation of Ventricular and Parenchymal Interstitial pressures

Under normal conditions, ventricular and parenchymal pressures are closely coupled suggesting

fluid continuity between the two compartments (Wiig & Reed, 1983). In our studies, ventricular

and periventricular (parenchymal) pressures were virtually identical following the injection of

IgG/IgM isotype controls into the ventricles (Figure 7-4 D). Nonetheless, we observed that

pressures could become uncoupled under certain circumstances. The most obvious example of

this relates to the differences between the ventricles and periventricular parenchyma after

injection of the antibodies to the β1 integrins (Figure 7-4 B). Unexpectedly however, we also

noted some other pressure differences.

It would seem that there was a tendency for the ventricular pressures to be greater after

injection of isotype controls compared with injection of the specific anti integrin antibodies

(Figure 7-4 C). If the integrin-matrix interactions in the periventricular area are disrupted, it

seems possible that the matrix will expand and water will enter this tissue from the ventricular

system. This may have the effect of reducing the ventricular pressure transiently in comparison

with injections of the isotype controls, which are expected to have little impact on the brain

interstitium.

117

8.3 Pressure gradients and hydrocephalus

At this point, we do not know the molecular mechanisms associated with the tissue pressure

drop after the injection of the antibodies to the β1 integrins. This might be due to disruption of

cell integrin-matrix interactions as noted earlier or possibly due to some as yet-uncharacterized

consequence of antibody-β1 integrin binding.

Additionally, it will be important to determine if the integrin-matrix concept has relevance to

recognized animal models of hydrocephalus. In a kaolin-based dog model of hydrocephalus,

Penn and colleagues were unable to measure any pressure gradients between the ventricle, brain

and subarachnoid space with pressure measurements taken from each location continuously

before and after kaolin administration (Penn et al., 2005). They used an InSite pressure

monitoring system (a capacitive-based sensor at the end of a pacemaker lead) for the

measurements. Whether these differences relate to the technology to measure pressure or to

unknown issues related to the complexity of hydrocephalus development remain to be

elucidated.

We were surprised initially that the injection of antibodies to the integrins induced

hydrocephalus. In the servo-null studies we had established the principle that injection of anti

integrin antibodies could lower interstitial fluid pressure in the brain. Nevertheless, we assumed

that the immunoglobulins would be removed from the CSF system rapidly and as a

consequence, would not be present long enough for a sustained pressure gradient to form.

Indeed, we had evidence to suggest that the antibody-induced reductions in parenchymal

pressures were transient in nature as in some cases, parenchymal pressures returned to baseline

rapidly (example in Figure 7-2B). Since we observed hydrocephalus in many animals, one

might argue that the pressure gradients need only be very transiently applied across the

ventricles into the periventricular tissues to cause ventriculomegaly. If this is the case, this could

provide the basis for understanding why many investigators have failed to observe pressure

118

gradients.

8.4 Perspective and Significance of cell-matrix concept

We observed a role for cell intregrin – matrix interactions in hydrocephalus development. What

is exciting regarding these studies is that in experimental setting, the reduction in tissue pressure

in skin at least can be reversed with selected drugs such as the anti-inflammatory agent α –

trinositol (D-myo-inositol 1,2,6 triphosphate) (Lund & Reed, 1994;Rodt et al., 1994) and an

isoform of platelet-derived growth factor (PDGF-BB) (Rodt et al., 1996). If this concept is

applicable to the brain, communicating hydrocephalus may ultimately be treatable with

pharmacological agents thus reducing the dependence on problematic shunts.

119

CHAPTER 9:

GENERAL DISCUSSION

120

9.1 Summary of Conclusions Contained in this Thesis

1) It has been commonly assumed that a CSF absorption defect occurs in communicating

hydrocephalus. However, apart from global measures of CSF outflow resistance, this has

never been measured directly. Our data demonstrates clearly that an absorption deficit occurs

at a discrete anatomical location. It is of importance to note that this deficiency relates to

absorption via extracranial lymphatic vessels.

2) The lymphatic CSF absorption deficit that exists in a kaolin model of hydrocephalus appears

to correlate with ventricular volumes. Additionally, measures of CSF outflow resistance in

this model also indicate that this impediment has a significant impact on global CSF drainage

(outflow resistance measurements). There is a correlation between CSF outflow resistance

and ventricular volumes.

3) Evidence from recent experiments suggests a new concept for hydrocephalus development. In

this notion, the extracellular matrix is a dynamic contributor to interstitial fluid pressure

regulation via integrin-matrix interactions. It seems that disruption of these connections

lowers brain parenchymal fluid pressure relative to that in the ventricles. This forms an

intramantle or intraparenchymal pressure gradient favouring ventricular expansion.

4) Due to the myriad ways in which integrin function is known to be impeded, we hypothesize

that disruption of matrix-integrin interactions represents a fundamental event in the

development of communicating hydrocephalus. In some cases, this may be related to

inflammation.

121

9.2 The development of hydrocephalus: Can one reconcile the 'classical' CSF absorption

deficit concept with the notion that the brain interstitium may be the 'epi-centre' of

dysfunction in ventricular expansion?

At this stage, there is no obvious answer to the question posed above. We continue to feel that

the pathogenesis of ventriculomegaly is unlikely to be reduced to a simple 'plumbing problem'

with our reasons for this view provided earlier. Is it possible therefore, that anomalies related to

CSF clearance represent ‘co-conspirators’ in hydrocephalus development but by themselves, do

not represent the initiating cause of the disorder? In this regard, could the 2 concepts be related

in any way? Along this line of reasoning, is it possible that pathological disturbances that

interfere with integrin-matrix interactions might also impact CSF clearance?

Inflammation is known to affect β1 integrin-matrix interactions and induce the interstitial

pressure lowering effects in skin (Wiig et al., 2003;Reed et al., 1992). In addition, based on

earlier work from the Johnston lab, inflammatory processes have a profound impact on

lymphatic function with many inflammatory mediators and cytokines inhibiting lymphatic

pumping activity (Johnston & Gordon, 1981;Elias et al., 1987;Elias & Johnston, 1988;Hanley et

al., 1989;Elias & Johnston, 1990) .

Of course, we have never tested whether the lymphatic vessels within the olfactory

turbinates can be affected similarly under inflammatory conditions but if so, this would link

matrix interactions and downstream lymphatic function in an interesting way. Kaolin clearly

induces local inflammatory responses in the subarachnoid spaces, such as arachnoiditis, which

contribute to the mechanical obstruction seen in models of both obstructive (McAllister et al.,

1985) and communicating hydrocephalus (Balasubramaniam & Del Bigio, 2002). Thus, one

could speculate that kaolin first induces matrix changes that lead to intra-mantle pressure

gradients favouring ventriculomegaly.

If this supposition has merit, one might develop a ‘two-hit’ hypothesis (Schematic Figure 9-

122

1) for communicating hydrocephalus in which the CSF absorption deficit is secondary to some

primary insult affecting the brain interstitium.

In the kaolin model of CH, one might speculate that kaolin first induces matrix changes that

lead to intra-mantle pressure gradients favouring ventriculomegaly. A concurrent CSF

absorption deficit induced by released inflammatory mediators/cytokines may alter downstream

CSF lymphatic drainage and in doing so, affect compliance and possibly intensify the magnitude

of this pressure gradient. We recently performed a small number of experiments approximately

2 weeks after kaolin or saline injection. Using the servo null device, we could find no evidence

of pressure gradients in 4 of the animals. However, in the two rats with the largest ventricles,

ventricular-parenchymal pressure differences of ~3.0 cmH2O were observed (example in Figure

Inflammation

Cytokines

Hemorrhage

2 - Hit Hypothesis for CH

Intra-parenchymal

pressure gradients and

Hydrocephalus

CSF absorption deficit

Elevate ICP

Beta-1 Integrin – Matrix interactions

Schematic Figure 9-1: Two-Hit hypothesis for CH

123

9-2). These data are still very preliminary, but if supported by additional studies would suggest

that a transparenchymal pressure gradient may co-exist with a lymphatic CSF absorption deficit.

Another pathophysiological insult to the system that may have relevance to this issue is

hemorrhage. There is a strong association between subarachnoid haemorrhage and the

development of hydrocephalus in the literature (van et al., 2007;Nieuwkamp et al., 2000). It is

of interest to note that, in a very limited number of preliminary experiments, we injected 100µl

heparinised blood into a ventricle and measured periventricular interstitial fluid pressures (n=1)

and ventricular pressures (n=1). We observed a decline in the parenchymal pressure with respect

to the ventricular pressure under these conditions (examples in Figure 9-3). The pressure

patterns were very similar to those observed following the injection of antibodies to the β1

integrins. There is some suggestion in the literature regarding a linkage between hemorrhage

and integrins (Del Zoppo, 1997). Furthermore, we must also consider that when blood enters a

lymphatic vessel it reduces pumping activity significantly (Elias et al., 1990;Wandolo et al.,

1992;Elias et al., 1992).

Hence, at first glance, it would appear that a matrix-centric view of ventricular expansion

and the CSF absorption impediment concept are independent and unrelated. However, it is

possible that a reduction in CSF absorption exacerbates the pressure differential caused by

matrix disruption. This might suggest a 'two-hit' hypothesis for ventriculomegaly; i.e. a

molecular disturbance in matrix integrity in the periventricular area may be an initiating event in

the development of some forms of hydrocephalus. A concurrent CSF absorption deficit may, in

some way, intensify the magnitude of the pressure gradient.

124

Figure 9-2 An example of ventricular and parenchymal pressure measured

simultaneously in a 2 weeks post kaolin injection animal with evens ratio of 0.51.

A ventricular-parenchymal pressure differences of ~3.0 cmH2O were observed

Evans ratio 0.51 Evans ratio 0.51

Ventricular Pressure

Parenchymal Pressure

125

A

B

C

A

B

C

A

B

C

Figure 9-3 Example of (A, B)

parenchymal and (C)

ventricular pressure

measured after (A, C) 100µl

heparinised blood and (B)

100µl heparinised saline

injection.

The parenchyamal pressure

declines with respect to the

baseline, after the injection of

100µl heparinised blood but the

ventricular pressure does not.

The parenchymal pressure after

100µl heparinised saline

injection does not below

baseline.

126

9.3 Issues for future investigation

One of the most important outcomes of our servo null-based studies is that we have been able to

uncouple ventricular CSF pressures from the pressures within the periventricular interstitium.

The resultant pressure gradients may represent an important component of ventricular expansion

in hydrocephalus of the non-obstructive type. If this were to be true, a number of physiological

conditions would need to be met. First, if the pressures were to decline in the periventricular

tissues, one would expect that CSF would be drawn into the parenchyma from the ventricle due

to the pressure gradient. Recent consultations with members of the Centre for Mathematical

Medicine at the Fields Institute (Arciero J et al., 2009) have indicated that, in order for CSF

pressure to be equal in the ventricles and the subarachnoid space but be lower in the

parenchyma, some removal of fluid must occur from the tissues. This concept is echoed in the

papers of Levine (Levine, 1999;Levine, 2008) as noted earlier and interestingly, is a necessary

condition for ventricular enlargement in the Cambridge model as well (Pena et al., 2002). The

potential absorption of brain interstitial fluid into the capillaries would seem an important

concept to investigate further. This question brings up several issues. First, it seems appropriate

in future studies to consider a role for the aquaporins as they have been implicated in playing a

role in tissue water management in a variety of brain pathophysiological states (Papadopoulos et

al., 2004;Verkman et al., 2006;Bloch & Manley, 2007) including hydrocephalus (Tourdias et

al., 2009). Second, whether hydrostatic or osmotic forces would control this water movement is

not known. Third, it is not clear whether the antibodies to the β1 integrins affected the integrity

of the blood-brain-barrier. Four, the molecular mechanisms associated with β1 integrin function

also require attention. At present, we do not know the specific molecular mechanisms associated

with the tissue pressure drop after the injection of the antibodies to the β1 integrins. This might

be due to disruption of cell integrin-matrix interactions or possibly due to some as yet,

uncharacterized consequence of antibody- β1 integrin binding. In addition, we need to

127

determine which cell-matrix interactions are affected and which subgroups of integrin receptors

are involved in generating the pressure lowering effect. These questions could be addressed

using a combination of classical physiological approaches and innovative new molecular

technologies and strategies.

128

Reference List

Abbott NJ (2005). Dynamics of CNS barriers: evolution, differentiation, and modulation. Cell Mol Neurobiol 25, 5-23.

Agha FP, Amendola MA, Shirazi KK, Amendola BE, & Chandler WF (1983). Unusual abdominal complications of ventriculo-peritoneal shunts. Radiology 146, 323-326.

Arciero J, Begg R, & Wilkie K. http://www.math.uwaterloo.ca/~kpwilkie/OCCAM_Hydrocephalus_Report.pdf . 2009. Ref Type: Electronic Citation

Baas D, Meiniel A, Benadiba C, Bonnafe E, Meiniel O, Reith W, & Durand B (2006). A deficiency in RFX3 causes hydrocephalus associated with abnormal differentiation of ependymal cells. Eur J Neurosci 24, 1020-1030.

Balasubramaniam J & Del Bigio MR (2002). Analysis of age-dependant alteration in the brain gene expression profile following induction of hydrocephalus in rats. Exp Neurol 173, 105-113.

Banizs B, Pike MM, Millican CL, Ferguson WB, Komlosi P, Sheetz J, Bell PD, Schwiebert EM, & Yoder BK (2005). Dysfunctional cilia lead to altered ependyma and choroid plexus function, and result in the formation of hydrocephalus. Development 132, 5329-5339.

Berg A, Rubin K, & Reed RK (2001). Cytochalasin D induces edema formation and lowering of interstitial fluid pressure in rat dermis. Am J Physiol Heart Circ Physiol 281, H7-13.

Bergsneider M, Miller C, Vespa PM, & Hu X (2008). Surgical management of adult hydrocephalus. Neurosurgery 62 Suppl 2, 643-659.

Bloch O & Manley GT (2007). The role of aquaporin-4 in cerebral water transport and edema. Neurosurg Focus 22, E3.

Bondurant CP & Jimenez DF (1995). Epidemiology of cerebrospinal fluid shunting. Pediatr Neurosurg 23, 254-258.

Borit A & Sidman RL (1972). New mutant mouse with communicating hydrocephalus and secondary aqueductal stenosis. Acta Neuropathol 21, 316-331.

129

Botel C & Brinker T (1994). Measurement of the dynamics of the cerebrospinal fluid system in the rat. J Exp Anim Sci 36, 78-83.

Boulton M, Flessner M, Armstrong D, Hay J, & Johnston M (1997). Lymphatic drainage of the CNS: effects of lymphatic diversion/ligation on CSF protein transport to plasma. Am J Physiol 272, R1613-R1619.

Boulton M, Flessner M, Armstrong D, Hay J, & Johnston M (1998). Determination of volumetric cerebrospinal fluid absorption into extracranial lymphatics in sheep. Am J Physiol 274, R88-R96.

Boulton M, Flessner M, Armstrong D, Mohamed R, Hay J, & Johnston M (1999). Contribution of extracranial lymphatics and arachnoid villi to the clearance of a CSF tracer in the rat. Am J Physiol 276, R818-R823.

Bozanovic-Sosic R, Mollanji R, & Johnston MG (2001). Spinal and cranial contributions to total cerebrospinal fluid transport. Am J Physiol Regul Integr Comp Physiol 281, R909-R916.

Bradley WG, Jr., Scalzo D, Queralt J, Nitz WN, Atkinson DJ, & Wong P (1996). Normal-pressure hydrocephalus: evaluation with cerebrospinal fluid flow measurements at MR imaging. Radiology 198, 523-529.

Brinker T, Botel C, Rothkotter HJ, Walter GF, & Samii M (1994). The perineural pathway of cerebrospinal fluid absorption into the cervical lymphatic system. Morphological findings in rats, cats, dogs and monkeys. In Intracranial Pressure IX, eds. Nagai H, Kamiya K, & Ishii K, pp. 132-135. Springer, Tokyo.

Brinker T, Ludemann W, Berens von RD, & Samii M (1997). Dynamic properties of lymphatic pathways for the absorption of cerebrospinal fluid. Acta Neuropathol 94, 493-498.

Browd SR, Gottfried ON, Ragel BT, & Kestle JR (2006). Failure of cerebrospinal fluid shunts: part II: overdrainage, loculation, and abdominal complications. Pediatr Neurol 34, 171-176.

Brown PD, Davies SL, Speake T, & Millar ID (2004). Molecular mechanisms of cerebrospinal fluid production. Neuroscience 129, 957-970.

Bruni JE, Del Bigio MR, Cardoso ER, & Persaud TV (1988). Neuropathology of congenital hydrocephalus in the SUMS/NP mouse. Acta Neurochir 92, 118-122.

Caversaccio M, Peschel O, & Arnold W (1996). The drainage of cerebrospinal fluid into the lymphatic system of the neck in humans. ORL J Otorhinolaryngol Relat Spec 58, 164-166.

130

Chahlavi A, El-Babaa SK, & Luciano MG (2001). Adult-onset hydrocephalus. Neurosurg Clin N Am 12, 753-60, ix.

Cserr HF, Cooper DN, Suri PK, & Patlak CS (1981). Efflux of radiolabeled polyethylene glycols and albumin from rat brain. Am J Physiol 240, F319-F328.

Dahme M, Bartsch U, Martini R, Anliker B, Schachner M, & Mantei N (1997). Disruption of the mouse L1 gene leads to malformations of the nervous system. Nature Genetics 17, 346-349.

Daniel GB, Edwards DF, Harvey RC, & Kabalka GW (1995). Communicating hydrocephalus in dogs with congenital ciliary dysfunction. Dev Neurosci 17, 230-235.

das Neves L., Duchala CS, Tolentino-Silva F, Haxhiu MA, Colmenares C, Macklin WB, Campbell CE, Butz KG, Gronostajski RM, & Godinho F (1999). Disruption of the murine nuclear factor I-A gene (Nfia) results in perinatal lethality, hydrocephalus, and agenesis of the corpus callosum. Proc Natl Acad Sci U S A 96, 11946-11951.

Davis LE (1981). Communicating hydrocephalus in newborn hamsters and cats following vaccinia virus infection. J Neurosurg 54, 767-772.

Davson H & Segal M B (1996). Physiology of the CSF and Blood Brain Barriers CRC, Boca Raton, FL.

Davson H, Welch K, & Segal M B (1987). Physiology and pathobiology of cerebrospinal fluid Chichill Livingstone, Edinburgh.

DAVSON H, KLEEMAN CR, & LEVIN E (1962). Quantitative studies of the passage of different substances out of the cerebrospinal fluid. J Physiol 161, 126-142.

DAVSON H & Welch K (1971). The permeation of several materials into the fluids of the rabbit's brain. J Physiol 218, 337-351.

Del Bigio MR (1995). The ependyma: a protective barrier between brain and cerebrospinal fluid. Glia 14, 1-13.

Del Zoppo GJ (1997). Microvascular responses to cerebral ischemia/inflammation. Ann N Y Acad Sci 823, 132-147.

Del Zoppo GJ & Milner R (2006). Integrin-matrix interactions in the cerebral microvasculature. Arterioscler Thromb Vasc Biol 26, 1966-1975.

131

Deo-Narine V, Gomez DG, Vullo T, Manzo RP, Zimmerman RD, Deck MD, & Cahill PT (1994). Direct in vivo observation of transventricular absorption in the hydrocephalic dog using magnetic resonance imaging. Invest Radiol 29, 287-293.

Diggs J, Price AC, Burt AM, Flor WJ, McKanna JA, Novak GR, & James AE, Jr. (1986). Early changes in experimental hydrocephalus. Invest Radiol 21, 118-121.

Drake JM, Kestle JR, Milner R, Cinalli G, Boop F, Piatt J, Jr., Haines S, Schiff SJ, Cochrane DD, Steinbok P, & MacNeil N (1998). Randomized trial of cerebrospinal fluid shunt valve design in pediatric hydrocephalus. Neurosurgery 43, 294-303.

Drake JM, Kestle JR, & Tuli S (2000). CSF shunts 50 years on--past, present and future. Childs Nerv Syst 16, 800-804.

Edwards RJ, Dombrowski SM, Luciano MG, & Pople IK (2004). Chronic hydrocephalus in adults. Brain Pathol 14, 325-336.

Egnor M, Zheng L, Rosiello A, Gutman F, & Davis R (2002). A model of pulsations in communicating hydrocephalus. Pediatr Neurosurg 36, 281-303.

Elias RM, Eisenhoffer J, & Johnston MG (1992). Role of endothelial cells in regulating hemoglobin-induced changes in lymphatic pumping. Am J Physiol 263, H1880-H1887.

Elias RM & Johnston MG (1988). Modulation of fluid pumping in isolated bovine mesenteric lymphatics by a thromboxane/endoperoxide analogue. Prostaglandins 36, 97-106.

Elias RM & Johnston MG (1990). Modulation of lymphatic pumping by lymph-borne factors after endotoxin administration in sheep. J Appl Physiol 68, 199-208.

Elias RM, Johnston MG, Hayashi A, & Nelson W (1987). Decreased lymphatic pumping after intravenous endotoxin administration in sheep. Am J Physiol 253, H1349-H1357.

Elias RM, Wandolo G, Ranadive NS, Eisenhoffer J, & Johnston MG (1990). Lymphatic pumping in response to changes in transmural pressure is modulated by erythrolysate/hemoglobin. Circ Res 67, 1097-1106.

Fiori MG, Sharer LR, & Lowndes HE (1985). Communicating hydrocephalus in rodents treated with beta,beta'-iminodipropionitrile (IDPN). Acta Neuropathol (Berl) 65, 209-216.

132

Fishman R (1992). Cerebrospinal fluid in diseases of the nervous system, 2nd ed. W.B. Saunders Co, Philadelphia.

Galbreath E, Kim SJ, Park K, Brenner M, & Messing A (1995). Overexpression of TGF-beta 1 in the central nervous system of transgenic mice results in hydrocephalus. J Neuropathol Exp Neurol 54, 339-349.

Gaskill SJ & Marlin AE (1989). Pseudocysts of the abdomen associated with ventriculoperitoneal shunts: a report of twelve cases and a review of the literature. Pediatr Neurosci 15, 23-26.

Ghersi-Egea JF, Finnegan W, Chen JL, & Fenstermacher JD (1996a). Rapid distribution of intraventricularly administered sucrose into cerebrospinal fluid cisterns via subarachnoid velae in rat. Neuroscience 75, 1271-1288.

Ghersi-Egea JF, Gorevic PD, Ghiso J, Frangione B, Patlak CS, & Fenstermacher JD (1996b). Fate of cerebrospinal fluid-borne amyloid beta-peptide: rapid clearance into blood and appreciable accumulation by cerebral arteries. J Neurochem 67, 880-883.

Gladson CL & Cheresh DA (1991). Glioblastoma expression of vitronectin and the alpha v beta 3 integrin. Adhesion mechanism for transformed glial cells. J Clin Invest 88, 1924-1932.

Graus-Porta D, Blaess S, Senften M, Littlewood-Evans A, Damsky C, Huang Z, Orban P, Klein R, Schittny JC, & Muller U (2001). Beta1-class integrins regulate the development of laminae and folia in the cerebral and cerebellar cortex. Neuron 31, 367-379.

Grooms SY, Terracio L, & Jones LS (1993). Anatomical localization of beta 1 integrin-like immunoreactivity in rat brain. Exp Neurol 122, 253-259.

Gupta N, Park J, Solomon C, Kranz DA, Wrensch M, & Wu YW (2007). Long-term outcomes in patients with treated childhood hydrocephalus. J Neurosurg 106, 334-339.

Hakim CA, Hakim R, & Hakim S (2001). Normal-pressure hydrocephalus. Neurosurg Clin N Am 12, 761-73, ix.

Hale PM, McAllister JP, Katz SD, Wright LC, Lovely TJ, Miller DW, Wolfson BJ, Salotto AG, & Shroff DV (1992). Improvement of cortical morphology in infantile hydrocephalic animals after ventriculoperitoneal shunt placement. Neurosurgery 31, 1085-1096.

133

Hanley CA, Elias RM, Movat HZ, & Johnston MG (1989). Suppression of fluid pumping in isolated bovine mesenteric lymphatics by interleukin-1: interaction with prostaglandin E2. Microvasc Res 37, 218-229.

Hebb AO & Cusimano MD (2001). Idiopathic normal pressure hydrocephalus: a systematic review of diagnosis and outcome. Neurosurgery 49, 1166-1184.

Hochwald GM (1985). Animal models of hydrocephalus: recent developments. Proc Soc Exp Biol Med 178, 1-11.

Hoppe-Hirsch E, Laroussinie F, Brunet L, Sainte-Rose C, Renier D, Cinalli G, Zerah M, & Pierre-Kahn A (1998). Late outcome of the surgical treatment of hydrocephalus. Childs Nervous System 14, 97-99.

Hurley RA, Bradley WG, Jr., Latifi HT, & Taber KH (1999). Normal pressure hydrocephalus: significance of MRI in a potentially treatable dementia. J Neuropsychiatry Clin Neurosci 11, 297-300.

Hynes RO (2002). Integrins: bidirectional, allosteric signaling machines. Cell 110, 673-687.

James AE, Jr., Burns B, Flor WF, Strecker EP, Merz T, Bush M, & Price (1975). Pathophysiology of chronic communicating hydrocephalus in dogs (Canis familiaris). Experimental studies. J Neurol Sci 24, 151-178.

James AE, Jr., Flor WJ, Novak GR, Strecker EP, & Burns B (1978). Evaluation of the central canal of the spinal cord in experimentally induced hydrocephalus. J Neurosurg 48, 970-974.

Jellinger K (1976). Neuropathological aspects of dementias resulting from abnormal blood and cerebrospinal fluid dynamics. Acta Neurol Belg 76, 83-102.

Johanson CE & Jones HC (2001). Promising vistas in hydrocephalus and cerebrospinal fluid research. Trends Neurosci 24, 631-632.

Johanson CE, Szmydynger-Chodobska J, Chodobski A, Baird A, McMillan P, & Stopa EG (1999). Altered formation and bulk absorption of cerebrospinal fluid in FGF-2-induced hydrocephalus. Am J Physiol 277, R263-R271.

Johnston M, Zakharov A, Koh L, & Armstrong D (2005). Subarachnoid injection of Microfil reveals connections between cerebrospinal fluid and nasal lymphatics in the non-human primate. Neuropathol Appl Neurobiol 31, 632-640.

134

Johnston M, Zakharov A, Papaiconomou C, Salmasi G, & Armstrong D (2004). Evidence of connections between cerebrospinal fluid and nasal lymphatic vessels in humans, non-human primates and other mammalian species. Cerebrospinal Fluid Res 1, 2.

Johnston MG & Gordon JL (1981). Regulation of lymphatic contractility by arachidonate metabolites. Nature 293, 294-297.

Jones HC (1985). Cerebrospinal fluid pressure and resistance to absorption during development in normal and hydrocephalic mutant mice. Exp Neurol 90, 162-172.

Jones HC, Deane R, & Bucknall RM (1987a). Developmental changes in cerebrospinal fluid pressure and resistance to absorption in rats. Brain Res 430, 23-30.

Jones HC, Deane R, & Bucknall RM (1987b). Developmental changes in cerebrospinal fluid pressure and resistance to absorption in rats. Brain Res 430, 23-30.

Jones HC & Gratton JA (1989). The effect of cerebrospinal fluid pressure on dural venous pressure in young rats. J Neurosurg 71, 119-123.

Kanaji M, Tada T, & Kobayashi S (1997). A murine model of communicating hydrocephalus: Role of TGF-beta1. J Clin Neurosci 4, 51-56.

Kandel ER, Schwartz JH, & Jessell TM (2000). Principles of Neural Science McGraw Hill, New York.

Keep RF & Jones HC (1990). A morphometric study on the development of the lateral ventricle choroid plexus, choroid plexus capillaries and ventricular ependyma in the rat. Brain Res Dev Brain Res 56, 47-53.

Kestle J, Drake J, Milner R, Sainte-Rose C, Cinalli G, Boop F, Piatt J, Haines S, Schiff S, Cochrane D, Steinbok P, & MacNeil N (2000). Long-term follow-up data from the Shunt Design Trial. Pediatr Neurosurg 33, 230-236.

Kida S, Pantazis A, & Weller RO (1993). CSF drains directly from the subarachnoid space into nasal lymphatics in the rat. Anatomy, histology and immunological significance. Neuropathol Appl Neurobiol 19, 480-488.

Kiefer M, Eymann R, & Steudel WI (2000). The dynamic infusion test in rats. Childs Nerv Syst 16, 451-456.

135

Kitagaki H, Mori E, Ishii K, Yamaji S, Hirono N, & Imamura T (1998). CSF spaces in idiopathic normal pressure hydrocephalus: morphology and volumetry. AJNR Am J Neuroradiol 19, 1277-1284.

Koh L, Nagra G, & Johnston M (2007). Properties of the lymphatic cerebrospinal fluid transport system in the rat: impact of elevated intracranial pressure. J Vasc Res 44, 423-432.

Koh L, Zakharov A, & Johnston M (2005). Integration of the subarachnoid space and lymphatics: is it time to embrace a new concept of cerebrospinal fluid absorption? Cerebrospinal Fluid Res 2, 6.

Koh L, Zakharov A, Nagra G, Armstrong D, Friendship R, & Johnston M (2006). Development of cerebrospinal fluid absorption sites in the pig and rat: connections between the subarachnoid space and lymphatic vessels in the olfactory turbinates. Anat Embryol (Berl) 211, 335-344.

Kondziella D, Ludemann W, Brinker T, Sletvold O, & Sonnewald U (2002). Alterations in brain metabolism, CNS morphology and CSF dynamics in adult rats with kaolin-induced hydrocephalus. Brain Res 927, 35-41.

Kosteljanetz M (1986). CSF dynamics and pressure-volume relationships in communicating hydrocephalus. J Neurosurg 64, 45-52.

Kulkarni AV, Drake JM, & Lamberti-Pasculli M (2001). Cerebrospinal fluid shunt infection: a prospective study of risk factors. J Neurosurg 94, 195-201.

Kume T, Deng KY, Winfrey V, Gould DB, Walter MA, & Hogan BL (1998). The forkhead/winged helix gene Mf1 is disrupted in the pleiotropic mouse mutation congenital hydrocephalus. Cell 93, 985-996.

Levine DN (1999). The pathogenesis of normal pressure hydrocephalus: a theoretical analysis. Bull Math Biol 61, 875-916.

Levine DN (2008). Intracranial pressure and ventricular expansion in hydrocephalus: have we been asking the wrong question? J Neurol Sci 269, 1-11.

Li J, McAllister JP, Shen Y, Wagshul ME, Miller JM, Egnor MR, Johnston MG, Haacke EM, & Walker ML (2008). Communicating hydrocephalus in adult rats with kaolin obstruction of the basal cisterns or the cortical subarachnoid space. Exp Neurol 211, 351-361.

Lorenzo AV, Page LK, & Watters GV (1970). Relationship between cerebrospinal fluid formation, absorption and pressure in human hydrocephalus. Brain 93, 679-692.

136

Lowhagen P, Johansson BB, & Nordborg C (1994). The nasal route of cerebrospinal fluid drainage in man. A light-microscope study. Neuropathol Appl Neurobiol 20, 543-550.

Luedemann W, Kondziella D, Tienken K, Klinge P, Brinker T, & Berens von RD (2002). Spinal cerebrospinal fluid pathways and their significance for the compensation of kaolin-hydrocephalus. Acta Neurochir Suppl 81, 271-273.

Lund T, Onarheim H, Wiig H, & Reed RK (1989). Mechanisms behind increased dermal imbibition pressure in acute burn edema. Am J Physiol 256, H940-H948.

Lund T & Reed RK (1994). Alpha-Trinositol inhibits edema generation and albumin extravasation in thermally injured skin. J Trauma 36, 761-765.

Lund T, Wiig H, & Reed RK (1988). Acute postburn edema: role of strongly negative interstitial fluid pressure. Am J Physiol 255, H1069-H1074.

Mann JD, Butler AB, Johnson RN, & Bass NH (1979). Clearance of macromolecular and particulate substances from the cerebrospinal fluid system of the rat. J Neurosurg 50, 343-348.

Mann JD, Butler AB, Rosenthal JE, Maffeo CJ, Johnson RN, & Bass NH (1978). Regulation of intracranial pressure in rat, dog, and man. Ann Neurol 3, 156-165.

Mayfrank L, Kissler J, Raoofi R, Delsing P, Weis J, Kuker W, & Gilsbach JM (1997). Ventricular dilatation in experimental intraventricular hemorrhage in pigs. Characterization of cerebrospinal fluid dynamics and the effects of fibrinolytic treatment. Stroke 28, 141-148.

McAllister JP & Chovan P (1998). Neonatal hydrocephalus. Mechanisms and consequences. Neurosurg Clin N Am 9, 73-93.

McAllister JP, Cohen MI, O'Mara KA, & Johnson MH (1991). Progression of experimental infantile hydrocephalus and effects of ventriculoperitoneal shunts: an analysis correlating magnetic resonance imaging with gross morphology. Neurosurgery 29, 329-340.

McAllister JP, Maugans TA, Shah MV, & Truex RC, Jr. (1985). Neuronal effects of experimentally induced hydrocephalus in newborn rats. J Neurosurg 63, 776-783.

McComb JG & Hyman S (1984). Lymphatic drainage of cerebrospinal fluid in the cat. In Hydrocephalus, eds. Shapiro K, Marmarou A, & Portnoy H, Raven Press, New York.

137

McComb GJ & Hyman S (1990). Lymphatic drainage of cerebrospinal fluid in the primate. In Pathophysiology of the blood-brain barrier, eds. Johansson BB & Widner H, pp. 421-437. Elsevier Science Publishers.

McComb JG, DAVSON H, Hyman S, & Weiss MH (1982). Cerebrospinal fluid drainage as influenced by ventricular pressure in the rabbit. J Neurosurg 56, 790-797.

McGirt MJ, Leveque JC, Wellons JC, III, Villavicencio AT, Hopkins JS, Fuchs HE, & George TM (2002). Cerebrospinal fluid shunt survival and etiology of failures: a seven-year institutional experience. Pediatr Neurosurg 36, 248-255.

McGirt MJ, Zaas A, Fuchs HE, George TM, Kaye K, & Sexton DJ (2003). Risk factors for pediatric ventriculoperitoneal shunt infection and predictors of infectious pathogens. Clin Infect Dis 36, 858-862.

Meier U, Kiefer M, & Bartels P (2002). The ICP-dependency of resistance to cerebrospinal fluid outflow: a new mathematical method for CSF-parameter calculation in a model with H-TX rats. J Clin Neurosci 9, 58-63.

Milner R & Campbell IL (2002). The integrin family of cell adhesion molecules has multiple functions within the CNS. J Neurosci Res 69, 286-291.

Miyagoe-Suzuki Y, Nakagawa M, & Takeda S (2000). Merosin and congenital muscular dystrophy. Microsc Res Tech 48, 181-191.

Moinuddin SM & Tada T (2000). Study of cerebrospinal fluid flow dynamics in TGF-beta 1 induced chronic hydrocephalic mice. Neurol Res 22, 215-222.

Mollanji R, Bozanovic-Sosic R, Silver I, Li B, Kim C, Midha R, & Johnston M (2001). Intracranial pressure accommodation is impaired by blocking pathways leading to extracranial lymphatics. Am J Physiol Regul Integr Comp Physiol 280, R1573-R1581.

Mollanji R, Bozanovic-Sosic R, Zakharov A, Makarian L, & Johnston MG (2002). Blocking cerebrospinal fluid absorption through the cribriform plate increases resting intracranial pressure. Am J Physiol Regul Integr Comp Physiol 282, R1593-R1599.

Mori K (1990). Hydrocephalus--revision of its definition and classification with special reference to "intractable infantile hydrocephalus". Childs Nerv Syst 6, 198-204.

138

Nagaraja TN, Patel P, Gorski M, Gorevic PD, Patlak CS, & Fenstermacher JD (2005). In normal rat, intraventricularly administered insulin-like growth factor-1 is rapidly cleared from CSF with limited distribution into brain. Cerebrospinal Fluid Res 2, 5.

Nagra G & Johnston MG (2007). Impact of ageing on lymphatic cerebrospinal fluid absorption in the rat. Neuropathol Appl Neurobiol 33, 684-691.

Nagra G, Li J, McAllister JP, Miller J, Wagshul M, & Johnston M (2008). Impaired lymphatic cerebrospinal fluid absorption in a rat model of kaolin-induced communicating hydrocephalus. Am J Physiol Regul Integr Comp Physiol 294, R1752-R1759.

Nieuwkamp DJ, de GK, Rinkel GJ, & Algra A (2000). Treatment and outcome of severe intraventricular extension in patients with subarachnoid or intracerebral hemorrhage: a systematic review of the literature. J Neurol 247, 117-121.

Nugent GR, Al-Mefty O, & Chou S (1979). Communicating hydrocephalus as a cause of aqueductal stenosis. J Neurosurg 51, 812-818.

Papadopoulos MC, Manley GT, Krishna S, & Verkman AS (2004). Aquaporin-4 facilitates reabsorption of excess fluid in vasogenic brain edema. FASEB J 18, 1291-1293.

Papaiconomou C, Bozanovic-Sosic R, Zakharov A, & Johnston M (2002). Does neonatal cerebrospinal fluid absorption occur via arachnoid projections or extracranial lymphatics? Am J Physiol Regul Integr Comp Physiol 283, R869-R876.

Papaiconomou C, Zakharov A, Azizi N, Djenic J, & Johnston M (2004). Reassessment of the pathways responsible for cerebrospinal fluid absorption in the neonate. Childs Nerv Syst 20, 29-36.

Pardridge WM, Oldendorf WH, Cancilla P, & Frank HJ (1986). Blood-brain barrier: interface between internal medicine and the brain. Ann Intern Med 105, 82-95.

Patwardhan RV & Nanda A (2005). Implanted ventricular shunts in the United States: the billion-dollar-a-year cost of hydrocephalus treatment. Neurosurgery 56, 139-144.

Paulus W, Baur I, Schuppan D, & Roggendorf W (1993). Characterization of integrin receptors in normal and neoplastic human brain. Am J Pathol 143, 154-163.

Paxinos G & Watson C (2007). The rat brain, in stereotaxic coordinates, 6th ed. Academic Press, London, UK.

139

Pena A, Harris NG, Bolton MD, Czosnyka M, & Pickard JD (2002). Communicating hydrocephalus: the biomechanics of progressive ventricular enlargement revisited. Acta Neurochir Suppl 81, 59-63.

Penn RD, Lee MC, Linninger AA, Miesel K, Lu SN, & Stylos L (2005). Pressure gradients in the brain in an experimental model of hydrocephalus. J Neurosurg 102, 1069-1075.

Philpot J, Cowan F, Pennock J, Sewry C, Dubowitz V, Bydder G, & Muntoni F (1999). Merosin-deficient congenital muscular dystrophy: the spectrum of brain involvement on magnetic resonance imaging. Neuromuscul Disord 9, 81-85.

Pickard JD (1982). Adult communicating hydrocephalus. Br J Hosp Med 27, 35, 37-8, 40, passim.

Portnoy HD, Branch C, & Castro ME (1994). The relationship of intracranial venous pressure to hydrocephalus. Childs Nerv Syst 10, 29-35.

Price DL, James AE, Jr., Sperber E, & Strecker EP (1976). Communicating hydrocephalus. Arch Neurol 33, 15-20.

Raimondi AJ, Clark SJ, & McLone DG (1976). Pathogenesis of aqueductal occlusion in congenital murine hydrocephalus. J Neurosurg 45, 66-77.

Redzic ZB & Segal MB (2004). The structure of the choroid plexus and the physiology of the choroid plexus epithelium. Adv Drug Deliv Rev 56, 1695-1716.

Reed RK & Rodt SA (1991). Increased negativity of interstitial fluid pressure during the onset stage of inflammatory edema in rat skin. Am J Physiol 260, H1985-H1991.

Reed RK, Rubin K, Wiig H, & Rodt SA (1992). Blockade of beta 1-integrins in skin causes edema through lowering of interstitial fluid pressure. Circ Res 71, 978-983.

Rekate HL (2008). The definition and classification of hydrocephalus: a personal recommendation to stimulate debate. Cerebrospinal Fluid Res 5, 2.

Rekate HL (2009). A contemporary definition and classification of hydrocephalus. Semin Pediatr Neurol 16, 9-15.

140

Robinson ML, Allen CE, Davy BE, Durfee WJ, Elder FF, Elliott CS, & Harrison WR (2002). Genetic mapping of an insertional hydrocephalus-inducing mutation allelic to hy3. Mammalian Genome: Official Journal Of The International Mammalian Genome Society 13, 625-632.

Rodt SA, Ahlen K, Berg A, Rubin K, & Reed RK (1996). A novel physiological function for platelet-derived growth factor-BB in rat dermis. J Physiol 495 ( Pt 1), 193-200.

Rodt SA, Reed RK, Ljungstrom M, Gustafsson TO, & Rubin K (1994). The anti-inflammatory agent alpha-trinositol exerts its edema-preventing effects through modulation of beta 1 integrin function. Circ Res 75, 942-948.

Rubin K, Sundberg C, Ahlen K, & Reed RK (1995). Integrins: transmembrane links between the extracellular matrix and the cell interior. In Interstitium, Connective Tissue and Lymphatics, eds. Reed RK, McHale NG, Bert JL, Winlove CP, & Laine GA, pp. 29-40. Portland Press Ltd., London.

Rubin RC, Hochwald GM, Tiell M, & Liwnicz BH (1976). Hydrocephalus: II. Cell number and size, and myelin content of the pre-shunted cerebral cortical mantle. Surg Neurol 5, 115-118.

Ruoslahti E (1996). Brain extracellular matrix. Glycobiology 6, 489-492.

Rutka JT, Apodaca G, Stern R, & Rosenblum M (1988). The extracellular matrix of the central and peripheral nervous systems: structure and function. J Neurosurg 69, 155-170.

Satz JS, Barresi R, Durbeej M, Willer T, Turner A, Moore SA, & Campbell KP (2008). Brain and eye malformations resembling Walker-Warburg syndrome are recapitulated in mice by dystroglycan deletion in the epiblast. J Neurosci 28, 10567-10575.

Segal MB (1993). Extracellular and cerebrospinal fluids. J Inherit Metab Dis 16, 617-638.

Segal MB (2001). Transport of nutrients across the choroid plexus. Microsc Res Tech 52, 38-48.

Sekula RF, Jr., Marchan EM, Oh MY, Kim DK, Frederickson AM, Pelz G, & Uchal M (2009). Laparoscopically assisted peritoneal shunt insertion for hydrocephalus. Br J Neurosurg 23, 439-442.

Silver I, Kim C, Mollanji R, & Johnston M (2002). Cerebrospinal fluid outflow resistance in sheep: impact of blocking cerebrospinal fluid transport through the cribriform plate. Neuropathol Appl Neurobiol 28, 67-74.

141

Silverberg GD, Levinthal E, Sullivan EV, Bloch DA, Chang SD, Leverenz J, Flitman S, Winn R, Marciano F, Saul T, Huhn S, Mayo M, & McGuire D (2002). Assessment of low-flow CSF drainage as a treatment for AD: results of a randomized pilot study. Neurology 59, 1139-1145.

Silverberg GD, Mayo M, Saul T, Rubenstein E, & McGuire D (2003). Alzheimer's disease, normal-pressure hydrocephalus, and senescent changes in CSF circulatory physiology: a hypothesis. Lancet Neurol 2, 506-511.

Simon TD, Riva-Cambrin J, Srivastava R, Bratton SL, Dean JM, & Kestle JR (2008). Hospital care for children with hydrocephalus in the United States: utilization, charges, comorbidities, and deaths. J Neurosurg Pediatr 1, 131-137.

Stallcup WB, Pytela R, & Ruoslahti E (1989). A neuroectoderm-associated ganglioside participates in fibronectin receptor-mediated adhesion of germinal cells to fibronectin. Dev Biol 132, 212-229.

Stoddart JHJ, Ladd D, Bronson RT, Harmon M, Jaworski J, Pritzker C, Lausen N, & Smith BD (2000). Transgenic mice with a mutated collagen promoter display normal response during bleomycin-induced fibrosis and possess neurological abnormalities. Journal of Cellular Biochemistry 77, 135-148.

Strazielle N & Ghersi-Egea JF (2000). Choroid plexus in the central nervous system: biology and physiopathology. J Neuropathol Exp Neurol 59, 561-574.

Strecker EP, James AE, Jr., Konigsmark B, & Merz T (1974). Autoradiographic observations in experimental communicating hydrocephalus. Neurology 24, 192-197.

Sunada Y, Edgar TS, Lotz BP, Rust RS, & Campbell KP (1995). Merosin-negative congenital muscular dystrophy associated with extensive brain abnormalities. Neurology 45, 2084-2089.

Szentistvanyi I, Patlak CS, Ellis RA, & Cserr HF (1984). Drainage of interstitial fluid from different regions of rat brain. Am J Physiol 246, F835-F844.

Szmydynger-Chodobska J, Chodobski A, & Johanson CE (1994). Postnatal developmental changes in blood flow to choroid plexuses and cerebral cortex of the rat. Am J Physiol 266, R1488-R1492.

Tada T, Kanaji M, & Kobayashi S (1994). Induction of communicating hydrocephalus in mice by intrathecal injection of human recombinant transforming growth factor-beta 1. J Neuroimmunol 50, 153-158.

142

Tada T, Zhan H, Tanaka Y, Hongo K, Matsumoto K, & Nakamura T (2006). Intraventricular administration of hepatocyte growth factor treats mouse communicating hydrocephalus induced by transforming growth factor beta1. Neurobiol Dis 21, 576-586.

Tenti G, Sivaloganathan S, & Drake JM (2002). The synchrony of arterial and CSF pulsations is not due to resonance. Pediatr Neurosurg 37, 221-222.

Tisell M, Hellstrom P, hl-Borjesson G, Barrows G, Blomsterwall E, Tullberg M, & Wikkelso C (2006). Long-term outcome in 109 adult patients operated on for hydrocephalus. Br J Neurosurg 20, 214-221.

Torack RM (1982). Historical aspects of normal and abnormal brain fluids. I. Cerebrospinal fluid. Arch Neurol 39, 197-201.

Tourdias T, Dragonu I, Fushimi Y, Deloire MS, Boiziau C, Brochet B, Moonen C, Petry KG, & Dousset V (2009). Aquaporin 4 correlates with apparent diffusion coefficient and hydrocephalus severity in the rat brain: a combined MRI-histological study. Neuroimage 47, 659-666.

Vale FA & Miranda SJ (2002). Clinical and demographic features of patients with dementia attended in a tertiary outpatient clinic. Arq Neuropsiquiatr 60, 548-552.

van GJ, Kerr RS, & Rinkel GJ (2007). Subarachnoid haemorrhage. Lancet 369, 306-318.

Verkman AS, Binder DK, Bloch O, Auguste K, & Papadopoulos MC (2006). Three distinct roles of aquaporin-4 in brain function revealed by knockout mice. Biochim Biophys Acta 1758, 1085-1093.

Vorbrodt AW & Dobrogowska DH (2003). Molecular anatomy of intercellular junctions in brain endothelial and epithelial barriers: electron microscopist's view. Brain Res Brain Res Rev 42, 221-242.

Wall M (2008). Idiopathic intracranial hypertension (pseudotumor cerebri). Curr Neurol Neurosci Rep 8, 87-93.

Wandolo G, Elias RM, Ranadive NS, & Johnston MG (1992). Heme-containing proteins suppress lymphatic pumping. J Vasc Res 29, 248-255.

Welch K & Friedman V (1960). The cerebrospinal fluid valves. Brain 83, 454-469.

143

Welch K & POLLAY M (1961). Perfusion of particles through arachnoid villi of the monkey. Am J Physiol 201, 651-654.

Weller RO, Kida S, & Zhang ET (1992a). Pathways of fluid drainage from the brain--morphological aspects and immunological significance in rat and man. Brain Pathol 2, 277-284.

Weller RO, Kida S, & Zhang ET (1992b). Pathways of fluid drainage from the brain--morphological aspects and immunological significance in rat and man. Brain Pathol 2, 277-284.

Whittle IR, Johnston IH, & Besser M (1985). Intracranial pressure changes in arrested hydrocephalus. J Neurosurg 62, 77-82.

Wiesmann M, Koedel U, Bruckmann H, & Pfister HW (2002). Experimental bacterial meningitis in rats: demonstration of hydrocephalus and meningeal enhancement by magnetic resonance imaging. Neurol Res 24, 307-310.

Wiig H & Reed RK (1983). Rat brain interstitial fluid pressure measured with micropipettes. Am J Physiol 244, H239-H246.

Wiig H, Rubin K, & Reed RK (2003). New and active role of the interstitium in control of interstitial fluid pressure: potential therapeutic consequences. Acta Anaesthesiol Scand 47, 111-121.

Williams B (1973). Is aqueduct stenosis a result of hydrocephalus? Brain 96, 399-412.

Woie K, Koller ME, Heyeraas KJ, & Reed RK (1993). Neurogenic inflammation in rat trachea is accompanied by increased negativity of interstitial fluid pressure. Circ Res 73, 839-845.

WOOLLAM DH (1957). The historical significance of the cerebrospinal fluid. Med Hist 1, 91-114.

Wright EM (1978). Transport processes in the formation of the cerebrospinal fluid. Rev Physiol Biochem Pharmacol 83, 3-34.

Yoffey JM & Drinker CK (1939). Some observations on the lymphatics of the nasal mucous membrane in the cat and monkey. J Anat 74, 45-52.

Zakharov A, Papaiconomou C, Djenic J, Midha R, & Johnston M (2003). Lymphatic cerebrospinal fluid absorption pathways in neonatal sheep revealed by subarachnoid injection of Microfil. Neuropathol Appl Neurobiol 29, 563-573.

144

Zakharov A, Papaiconomou C, & Johnston M (2004a). Lymphatic vessels gain access to cerebrospinal fluid through unique association with olfactory nerves. Lymphat Res Biol 2, 139-146.

Zakharov A, Papaiconomou C, Koh L, Djenic J, Bozanovic-Sosic R, & Johnston M (2004b). Integrating the roles of extracranial lymphatics and intracranial veins in cerebrospinal fluid absorption in sheep. Microvasc Res 67, 96-104.

Zemack G & Romner B (2002). Adjustable valves in normal-pressure hydrocephalus: a retrospective study of 218 patients. Neurosurgery 51, 1392-1400.

Zhang J, Williams MA, & Rigamonti D (2006). Genetics of human hydrocephalus. J Neurol 253, 1255-1266.

Zheng W, Aschner M, & Ghersi-Egea JF (2003). Brain barrier systems: a new frontier in metal neurotoxicological research. Toxicol Appl Pharmacol 192, 1-11.

145

Appendix

Our paper “Impact of ageing on lymphatic cerebrospinal fluid absorption in the rat” was

published in Neuropathology and Applied Neurobiology in 2007 and is included in this

Appendix section. Early in my studies, the original intention was to focus on CSF absorption

issues in ageing but as time progressed, we became more interested in the biomechanics of

hydrocephalus.

146

Impact of ageing on lymphatic cerebrospinal fluidabsorption in the rat

G. Nagra and M. G. Johnston

Neuroscience Program, Department of Laboratory Medicine and Pathobiology, Sunnybrook Health Sciences Centre,University of Toronto, Toronto, Ontario, Canada

G. Nagra and M. G. Johnston (2007) Neuropathology and Applied Neurobiology 33, 684–691Impact of ageing on lymphatic cerebrospinal fluid absorption in the rat

Several parameters associated with the cerebrospinal fluid(CSF) system show a change in the later stages of life,including elevated CSF outflow resistance. The latterimplies a CSF absorption deficit. As a significant portion ofCSF absorption occurs into extracranial lymphatic vesselslocated in the olfactory turbinates, the purpose of thisstudy was to determine whether any age-related impedi-ments to CSF absorption existed at this location. In previ-ous studies, we observed rapid movement of the CSF tracerinto the olfactory turbinates in young rats (peaking30 min after injection), with the concentration of thetracer being much higher in the turbinates than in anyother tissue measured. In the study reported here, 125I-

human serum albumin was injected into the lateral ven-tricles of 3-, 6-, 12- and 19-month-old Fisher 344 rats.The animals were sacrificed at various times after injec-tion of the radioactive tracer, and appropriate tissuesamples were extracted. At 30 min post injection, theaverage tracer values expressed as per cent injected/gtissue were 6.68 � 0.42 (n = 9, 3 months), 4.78 � 0.67(n = 9, 6 months), 2.49 � 0.31 (n = 9, 12 months) and2.42 � 0.72 (n = 9, 19 months). We conclude that lym-phatic CSF transport declines significantly with age. Inconcert with the known drop in CSF formation, the reduc-tion in lymphatic CSF absorption may contribute to adecrease in CSF turnover in the elderly.

Keywords: Alzheimer’s disease, arachnoid granulations and villi, cribriform plate, normal pressure hydrocephalus,olfactory turbinates

Introduction

Diseases of the cerebrospinal fluid (CSF) system are notuncommon, with perhaps hydrocephalus representingthe prototypical CSF disorder. However, this system can beperturbed in more subtle ways, and we are beginning torealize that altered CSF function may contribute to diseasein a manner not conceived of in the past. In this regard,several of the important parameters associated with theCSF system show a change in the later stages of life. These

include a fall in CSF production [1] and elevated CSFoutflow resistance [2]. While the implications of thesechanges are not fully understood, the resultant decline inCSF turnover is believed to contribute to the accumulationof toxic material in the ventricular and subarachnoidcompartments and, in some patients possibly, to facilitateneurological impairment, including dementia [3].

The fact that CSF outflow resistance increases withage suggests that CSF absorption declines in the elderly.However, estimates of outflow resistance do not provideany information on the nature or location of the impedi-ment to CSF drainage. Most would probably assume that ifan impediment to CSF outflow exists in the older popula-tion, the obstruction would occur at the arachnoid villiand granulations, preventing CSF transport into thecranial venous sinuses. While there may be some merit in

Correspondence: Dr Miles G. Johnston, Department of LaboratoryMedicine and Pathobiology, Neuroscience Research, SunnybrookHealth Sciences Centre, University of Toronto, Research Building,S-111, 2075 Bayview Avenue, Toronto, Ontario, M4N 3M5, Canada.Tel: +1 416 480 5700; Fax: +1 416 480 5737; E-mail:[email protected]

684 © 2007 Blackwell Publishing Ltd

Neuropathology and Applied Neurobiology (2007), 33, 684–691 doi: 10.1111/j.1365-2990.2007.00857.x

147

mailto:[email protected]

this concept, the accepted views of the normal mecha-nisms and pathways responsible for CSF transport havebeen challenged in recent years and new conceptshave emerged. Most notably, quantitative experimentshave demonstrated that CSF absorption by the arachnoidprojections under normal circumstances may have beenexaggerated [4–6]. In contrast, quantitative and qualita-tive evidence increasingly supports the view that a signifi-cant portion of CSF is drained from the subarachnoidcompartment by extracranial lymphatic vessels. WhileCSF may be absorbed by lymphatic vessels at numerousanatomical locations, the majority appears to convectthrough the foramina of the cribriform plate in associa-tion with the olfactory nerves [7,8]. This CSF is absorbedby an extensive complex of lymphatic vessels in the olfac-tory turbinates and is conveyed through a variety oflymph nodes in the head and neck region to the cervicallymphatics [9–12].

If a decline in CSF absorption is a significant contributorto the CSF circulatory disturbances in the elderly, it will beimportant to establish where the impediment to CSF trans-port occurs. As a first step in addressing this issue, ourobjective in this study was to quantify CSF uptake intoethmoidal lymphatic vessels in rats of various ages. Wereport that lymphatic CSF transport declines significantlywith ageing in the Fisher 344 rat.

Materials and methods

A total of 54 male Fisher 344 rats (from 3 to 19 months ofage) were used for this investigation (purchased fromHarlan, Madison, WI, USA). The average weights of theanimals used in this study were 248 � 5, 329 � 9,375 � 11 and 414 � 6 g at 3, 6, 12 and 19 months,respectively. The adult animals were fed lab rat chow(LabDiet 5001) until sacrifice. All experiments wereapproved by the ethics committee at the SunnybrookHealth Science Centre and conformed to the guidelines setby the Canadian Council on Animal Care and the Animalsfor Research Act of Ontario.

The development of the method to assess lymphaticCSF uptake in the rat has been described in a previouspublication [13]. The rapid movement of CSF tracers(dyes or radioactive proteins) into the olfactory turbi-nates supports a role for lymphatics in CSF absorptionand provides the basis of the method utilized in thisreport to assess the impact of ageing on lymphatic CSFtransport.

Rats were anaesthetized initially by placement in acustom-built rodent anaesthesia chamber using hal-othane (4–5%) in oxygen. For the experimental procedure,they were maintained with 2–2.5% halothane in oxygendelivered by a nose cone (Rat Anaesthesia Mask, KOPF,Model 906, Tujunga, CA, USA). The animals were placedon a heating pad (Fine Science Tools, Vancouver, BC,Canada) and fixed in position in a Small Animal Stereo-taxic device (KOPF, Model 900). The skin over the craniumwas removed and the junction of the sagittal and coronalsutures identified. A small high-speed micro drill with arounded tip (Fine Science Tools) was used to grind awaythe bone to expose the dural membrane.

A 50-mL Hamilton syringe (Fisher Scientific, Toronto,Ontario, Canada) with a 30-gauge needle was used at0.92 mm posterior and 1.4 mm lateral to the bregma, and3.3 mm deep to the dural membrane at the left or righthemisphere. The needle was loaded with 125I-humanserum albumin (HSA) and the tip lowered into one of thelateral ventricles (the right and left ventricles were usedrandomly in this study). The coordinates were noted fromthe stereotaxic instrument and adjusted according to thereference values from a rat brain atlas [14]. In total, 50 mLcontaining 500 mg of 125I-HSA (0.93 MBq/mL, 10 mg/mL, Drax Image, Quebec, Canada) was injected into one ofthe lateral ventricles. The needle was removed after1–2 min and the needle path sealed with bone wax. Aftervariable periods, the animals were sacrificed by injection of1.0 mL euthanyl i.p. Immediately before death, a bloodsample was taken from the heart. After death, the kidneyand the spleen were removed, and samples from liver, skel-etal muscle and tail were collected. All tissue samples wereplaced into preweighed glass test tubes for weight determi-nation using a Mettler BB2400 balance (Fisher Scientific,Unionville, ON, Canada). The carcasses were then frozenfor at least 24 h in a freezer.

To facilitate the assessment of radioactivity in the olfac-tory submucosa and to prevent potential post mortemtracer contamination from the CSF compartment, aportion of the turbinates were cut from frozen tissue. Theupper, lower and middle olfactory turbinates were excisedwith a fine tooth saw as described in detail in our previouspaper and assessed separately for radioactivity [13]. Theupper portion represented partial superior cells of theethmoid turbinates along with cartilage and soft tissues ofthe nasal wall. The middle portion contained the mainportion of the turbinates (henceforth termed middle tur-binates). The lower portion consisted of a fraction of the

Impact of ageing on lymphatic CSF absorption 685

© 2007 Blackwell Publishing Ltd, Neuropathology and Applied Neurobiology, 33, 684–691

148

posterior cells of the turbinates and part of the hardpalate. The samples were weighed as described aboveand were monitored for radioactivity in a multichannelgamma spectrometer (Compugamma, LKB Wallac, Turku,Finland). To orient the reader, Figure 1 illustrates theolfactory turbinates in a rat. In this example, Evans bluedye has been introduced into a lateral ventricle and the

animal sacrificed after 20 min. The Evans blue (dark area)can be observed in the olfactory turbinates.

Analysis of data

The enrichment of the CSF tracer in various tissues wasexpressed as per cent injected dose/g tissue. All data wereexpressed as the mean � SE. The results were analysedwith a one-way anova, followed by the post-hoc Tukeystudentized range test. We interpreted P < 0.05 assignificant.

Results

In 3-month-old rats the highest concentrations of 125I-HSA were found in the middle turbinate area, which rep-resented the bulk of the olfactory turbinates. An exampleis provided in Figure 2 (black bars). The lower turbinates(which include some of the posterior cells of the turbi-nates and part of the hard palate) also contained highconcentrations of the tracer. The radioactivity in theupper turbinates (representing partial superior cells of theethmoid turbinates along with cartilage and soft tissues ofthe nasal wall) was similar to that observed in blood.Tracer recoveries in skeletal muscle, spleen, kidney, liverand tail were much lower and likely were reflective of thetracer within the vasculature of these tissues. Similar

Brain

Olfactory BulbCribriform

Plate

OlfactoryTurbinates

Figure 1. Sagittal section of rat head illustrating the olfactoryturbinates and attendant structures. Evans blue dye was injectedinto a lateral ventricle 20 min before the animal was sacrificed. Theturbinates appear dark in this image due to the transport of the dyeacross the cribriform plate into the olfactory turbinates. A scaleruler is provided at the bottom of the image in mm.

Per cent injected/gm tissue

0 1 2 3 4 5 6 7

Lower turb

Middle turb

Upper turb

Blood

Skeletal muscle

Spleen

Kidney

Liver

Tail

Figure 2. Examples of the distribution of radioactivity into various tissues after 125I-HSA injection into a lateral ventricle. Black barsrepresent data from a 3-month-old rat. Grey bars illustrate data from a 19-month-old animal. The recovery of the tracer in the turbinatetissues was much greater in the young animal.

686 G. Nagra and M. Johnston


149

assessments in older animals revealed that the upper,middle and lower turbinate concentrations were consider-ably less than those observed at 3 months. An example isprovided in Figure 2 (grey bars).

Figure 3A illustrates all of the 30-min middle turbinateenrichment data (n = 9 for each group) plotted as a func-tion of age. It is clear that the transport of the CSF tracerinto the middle olfactory turbinates declined with age.Figure 3B illustrates the averaged data assessed at 30 minafter tracer injection. The impediment to transport acrossthe cribriform plate appeared to plateau at 12 months, asthe tracer recoveries at 12 and 19 months were not sig-nificantly different from one another. At 30 min postinjection, the average tracer values expressed as per cent

injected/g tissue were 6.68 � 0.42 (n = 9, 3 months),4.78 � 0.67 (n = 9, 6 months), 2.49 � 0.31 (n = 9,12 months) and 2.42 � 0.72 (n = 9, 19 months). anova

revealed a significant effect of age on tracer enrichment inthe middle turbinates. The tracer concentrations in bloodare provided in the inset to Figure 3B. Overall, the bloodlevels of the CSF tracer did not change significantly overtime, although the concentration of 125I-HSA seemed to behighest in the 6-month-old rats.

To determine whether the peak turbinate concentra-tions of the radioactive tracer were achieved at differenttimes in the older animals, an additional series of experi-ments was performed in the 12- to 19-month-old rats. Inthis series, the tracer concentrations were monitored 10,20, 40 and 60 min after 125I-HSA installation in the ven-tricles (Figure 4). For comparison, the time-course datafrom approximately 3-month-old animals obtained in ourprevious report [13] have been replotted in Figure 4(inset). In the younger animals, movement of the CSFtracer across the cribriform plate into the olfactory turbi-nates has a distinct peak at 30 min after injection. In con-trast, the pooled data from the 12- to 19-month-old rats

0 3 6 9 12 15 18 21

0

2

4

6

8

0 3 6 9 12 15 18 21

0

2

4

6

8

10T

race

r E

nric

hmen

t in

Olf

acto

ry T

urbi

nate

s

Time (months)

A

B

0 3 6 9 12 15 18 210

2

4

6

8

* *† †

Figure 3. Relationship between the age of the rats and the tracerenrichment in the middle turbinates. In (A), all of the data havebeen plotted with the regression line. B illustrates the averaged datafor each age group. One-way anova revealed a significant effect ofage on the movement of the CSF tracer into the olfactoryturbinates. Pair-wise comparisons indicated that the tracerenrichment in the turbinates of 3- and 6-month-old animals wassignificantly greater than those measured in the 12- and19-month-old groups. In B, *P < 0.05 compared with the3-month-old rats; †P < 0.05 compared with the 6-month-oldanimals. No significant differences were observed between the3- and 6-month-old rats or between the 12- and 19-month-oldanimals. The inset in B illustrates the average concentration of theCSF tracer in blood in all age groups. CSF, cerebrospinal fluid.

Time (minutes)

0 10 20 30 40 50 60Tra

cer

Enrichm

ent in

Mid

dle

Turb

inate

s

0

2

4

6

8

10

0 10 20 30 40 50 60

0

2

4

6

8

10

Figure 4. Relationship between the enrichment of the CSF tracerin the middle turbinates and the time of assessment. For thisanalysis, the data from the 12- and 19-month-old animals werepooled (10 min, n = 4; 20 min, n = 4; 30 min, n = 18, 40 min,n = 5, 60 min, n = 4). The inset illustrates the time course of theappearance of the tracer in the olfactory turbinates for3-month-old rats. These data have been replotted from an earlierpublication [13]. In the young animals (inset), the concentration ofthe tracer peaks at 30 min after introduction into a lateralventricle. In contrast, in the 12- to 19-month-old rats, tracerappearance in the middle olfactory turbinates increases slowly withtime, suggesting an age-related impediment to CSF transport acrossthe cribriform plate. CSF, cerebrospinal fluid.



150

indicated a slower transport into the turbinate tissues witha steady increase in the enrichment of the tracer up to60 min.

Discussion

The concept that CSF absorptive capacity declines withage has been based on estimates of global CSF outflowresistance [2] or studies that have indicated that radio-iodinated serum albumin is cleared from the brain veryslowly in older individuals compared with younger agegroups [15]. Consequently, the location of the transportdeficit (if it truly exists) has never been established. In thetextbook view, essentially all bulk CSF absorption isbelieved to occur through the microscopic arachnoid villiand macroscopic granulations into the lumen of cranialvenous sinuses. It would seem reasonable to suggest,therefore, that some impediment to CSF absorption mayoccur at these sites. However, while there are some in vitrodata using isolated portions of dura that suggest a func-tion for arachnoid projections [16,17], quantitativeexperiments in cats, rabbits, monkeys [18–20] and sheep[5,6] indicate that little, if any, CSF transports into thecranial venous system at normal CSF pressures. The avail-able data suggest that CSF transport can occur into thecranial venous system but only at pathologically highintracranial pressures [5,6]. This suggests that if animpediment to CSF outflow occurs in the elderly, it mightbe prudent to look at other potential absorption sites.

In this regard, there is abundant quantitative and quali-tative evidence to suggest that extracranial lymphaticvessels have an important role in CSF transport [21–24].The most important connections between the subarach-noid compartment and extracranial lymphatics appear toexist at the level of the cribriform plate. Lymphatic vesselswithin the olfactory submucosa form a distinctive asso-ciation with the olfactory nerves, and this anatomicalrelationship serves to facilitate CSF removal from thesubarachnoid space [7,10].

Once instilled in ventricular CSF, a radioactive proteinenters the olfactory turbinates rapidly [13]. Taking advan-tage of this observation, in this report we compared theconcentration of 125I-HSA administered into the CSF spacein the turbinates in rats of varying ages. We provide thefirst data to support the view that ‘normal’ ageing isassociated with a decline in CSF transport across thecribriform plate into the ethmoidal lymphatic system. Forexample, the tracer enrichment levels in the middle olfac-

tory turbinates at 12 and 19 months were 37% and 36%of those measured in the 3-month-old animals. It shouldbe noted that the CSF volume in rats is known to increasewith age [1]. As we injected the same amount of 125I-HSAinto the ventricles in all animals, this tracer would tend tobe diluted more in the CSF of older animals than in theyounger rats. Consequently, every unit volume of CSF thattransported across the cribriform plate would containless radioactivity in the older rats compared with theiryounger counterparts. Nonetheless, we believe that thesedifferences would be small. Based on published data [1],the increase in CSF volume from 3 to 19 months isapproximately 25%. Taking this factor into consideration,we might assume that the turbinate recoveries in the19-month-old rats would be underestimated by thisamount. If we were to add 25% to the tracer enrichment ofthe middle turbinates in the 19-month-old animals, thiswould give a value of 3.03, which is still considerablybelow the 6.68 measured in the youngest animals.

Whether this impaired transport was due to reducedmovement of CSF to the subarachnoid space surroundingthe olfactory bulbs, decreased transport through the crib-riform foramina or resulted from pathological changesin the olfactory turbinates that limited lymphatic CSFuptake, will have to be determined with additional experi-ments. It is of interest to note in this regard, that othershave observed a 47% reduction in human cribriformplate foramina area in males greater than 50 years oldcompared with younger individuals (28% reduction infemales) [25]. This may be due to increased calcification ofthe cribriform plate in the elderly [26]. The narrowing ofthe cribriform foramina would likely reduce CSF transportinto the nasal lymphatics. In addition, in a TGFb1 mousemodel of hydrocephalus, the time taken for ink adminis-tered into the lateral ventricles to stain the cervical lymphnodes was considerably lengthened compared with non-hydocephalic animals [27,28]. This suggested that thecribriform-lymphatic connection was disrupted in theTGFb1-infused hydrocephalic group. Taken together, theseobservations indicate the potential for disruption of CSFtransport across the cribriform plate.

In addition to a reduction in CSF absorption, there areother important changes in the CSF system that occurwith normal ageing. CSF volumes increase in the elderlydue at least in part to brain atrophy [29,30]. Additionally,in humans there is some indication that the permeabil-ity of the blood–CSF barrier increases with age suchthat albumin permeability indexes in individuals aged



151

51–70 years and in patients over 71 years of age was sig-nificantly greater than those measured in patients aged20–50 years [31]. Finally, CSF production is less in theelderly [32–34]. In the ageing rat, for example, CSF secre-tion declines 46% between 3 and 30 months of age [1].However, this reduction does not achieve significanceuntil the animals are 30 months old. It would seem then,that in the rat there is a period during which CSF produc-tion remains relatively normal while CSF absorptionthrough the cribriform plate has declined.

In this regard, it seems likely that the host can afford tolose a certain (unknown) number of CSF absorption sites,as presumably, other locations or mechanisms can com-pensate up to a certain extent. The recruitment of newabsorption pathways might initially work to maintain CSFpressure in the normal range. However, as more and moreCSF absorption sites become compromised, the diminishedCSF absorptive capacity would likely be reflected by areduced CSF clearance to plasma with attendant elevationof CSF outflow resistance and increases in intracranialpressure. Other potential absorption pathways couldinclude drainage through the perineurial spaces of othercranial/spinal nerves and/or clearance across the arach-noid villi and granulations into the cranial veins. Theabsorption of CSF from the spinal subarachnoid compart-ment may be particularly important [35]. In support ofthis, we noted previously that CSF outflow resistanceincreased when CSF transport through the cribriformplate was obstructed [36], but this increase was muchmore evident when CSF absorption from the spinal cordwas prevented [4,37].

At face value, the data in the inset to Figure 3 wouldsuggest that CSF transport to plasma was not affected sig-nificantly by a decline in CSF movement across the cribri-form plate. However, the methods used in this report donot permit an adequate exploration of this issue. What isrequired is a quantitative estimate of the mass transportrates of the tracer to plasma with correction of the plasmarecoveries for filtration using methods we employed in pre-vious studies [21,24]. This is important as there is evi-dence that the permeability of the vasculature is affectedby age. Indeed, a decrease in protein permeability with agehas been observed in rats [38]. This suggests that lesstracer would refilter back into the tissues in the olderanimals and thus, the plasma concentrations of the CSFtracer would be relatively higher than those in a youngeranimal, in which filtration out of the vascular compart-ment would progress at a faster rate.

Relevance to humans

The issue of lymphatic CSF transport in humans has notbeen resolved adequately. For obvious ethical reasons, ithas been very difficult to address this issue experimentally,but this situation may change with the development ofappropriate imaging modalities in the future. Nonetheless,recent studies have demonstrated a direct anatomicallinkage between CSF and extracranial lymph in humanand non-human primates [11,12]. Based on these data,we have no reason to believe that humans have evolvedsignificantly different CSF outflow mechanisms (at leastqualitatively) compared with those of other mammalianspecies. Indeed, indirect evidence from other groupswould also seem to support this view [39–42].

It is not uncommon to hear the view that sealing thecribriform plate clinically does not lead to hydrocephalus.However, the physiological and surgical realities are verycomplex and several factors must be considered. First, 8%of patients that undergo cranial base surgery for tumoursdo develop hydrocephalus [43], but the mechanism hasnever been determined. Second, during the repair of CSFleaks (after traumatic injury, for example), it would be dif-ficult to determine whether all or only a portion of the‘olfactory pathways’ that link the CSF and lymph com-partments have been obliterated. Third, as noted earlier,sealing the cribriform plate leads to the recruitment ofother lymphatic absorption sites, especially those associ-ated with the spinal subarachnoid compartment [37]. Infact, CSF appears to convect along most of the cranial andspinal nerves [44]. However, it is very difficult to quantifythis transport. The movement of CSF through theforamina of the cribriform plate into olfactory lymphaticvessels represents the paradigm for lymphatic CSF trans-port and has the practical advantage that absorption atthis location can be measured several ways. Until we findevidence to the contrary, it also seems likely that anyimpediments that occur at this location will be mirroredby similar deficits at other sites at which CSF and lymphcome in contact.

Significance of these findings

The reduction in CSF formation and the decline in CSFabsorption that accompanies ageing suggest that CSFturnover is ultimately compromised in the elderly. Theinability to clear potentially toxic metabolic productscould lead to their enhanced levels in brain interstitial



152

fluid, creating a hostile environment for neuronal func-tion and survival. Some investigators have postulated thatthis mechanism may contribute to the pathogenesis ofnormal pressure hydrocephalus and Alzheimer’s disease[3]. Of course, there are other important factors to con-sider. Cerebrovascular disease is believed to compromisethe periarterial interstitial fluid drainage pathways of thebrain, leading to the accumulation of A-beta in the brainparenchyma of Alzheimer’s patients [45,46]. In anyevent, the combination of a CSF absorption deficit, the lossof CSF secretory capacity and the presence of cerebrovas-cular disease may contribute to a reduction in CSF turn-over, and could provide the setting in which metabolicproducts toxic to the brain increase in concentration andalter neurological function negatively in several diseasesassociated with the elderly.

Acknowledgements

This research was funded by the Canadian Institutes ofHealth Research (CIHR). We also wish to thank M. Katic(Department of Research Design and Biostatistics, Sunny-brook Health Sciences Centre) for assistance in the com-putational analyses of the data.

References

1 Preston JE. Ageing choroid plexus-cerebrospinal fluidsystem. Microsc Res Tech 2001; 52: 31–7

2 Albeck MJ, Skak C, Nielsen PR, Olsen KS, Borgesen SE,Gjerris F. Age dependency of resistance to cerebrospinalfluid outflow. J Neurosurg 1998; 89: 275–8

3 Silverberg GD, Mayo M, Saul T, Rubenstein E, McGuire D.Alzheimer’s disease, normal-pressure hydrocephalus,and senescent changes in CSF circulatory physiology: ahypothesis. Lancet Neurol 2003; 2: 506–11

4 Papaiconomou C, Bozanovic-Sosic R, Zakharov A,Johnston M. Does neonatal cerebrospinal fluid absorptionoccur via arachnoid projections or extracranial lymphat-ics? Am J Physiol Regul Integr Comp Physiol 2002; 283:R869–76

5 Papaiconomou C, Zakharov A, Azizi N, Djenic J, JohnstonM. Reassessment of the pathways responsible for cere-brospinal fluid absorption in the neonate. Childs Nerv Syst2004; 20: 29–36

6 Zakharov A, Papaiconomou C, Koh L, Djenic J,Bozanovic-Sosic R, Johnston M. Integrating the roles ofextracranial lymphatics and intracranial veins in cere-brospinal fluid absorption in sheep. Microvasc Res 2004;67: 96–104

7 Kida S, Pantazis A, Weller RO. CSF drains directly from thesubarachnoid space into nasal lymphatics in the rat.

Anatomy, histology and immunological significance.Neuropathol Appl Neurobiol 1993; 19: 480–8

8 Koh L, Zakharov A, Johnston M. Integration of the sub-arachnoid space and lymphatics: is it time to embrace anew concept of cerebrospinal fluid absorption? Cerebrospi-nal Fluid Res 2005; 2: 6

9 Zakharov A, Papaiconomou C, Djenic J, Midha R,Johnston M. Lymphatic cerebrospinal fluid absorptionpathways in neonatal sheep revealed by subarachnoidinjection of Microfil. Neuropathol Appl Neurobiol 2003; 29:563–73

10 Zakharov A, Papaiconomou C, Johnston M. Lymphaticvessels gain access to cerebrospinal fluid through uniqueassociation with olfactory nerves. Lymphat Res Biol 2004;2: 139–46

11 Johnston M, Zakharov A, Papaiconomou C, Salmasi G,Armstrong D. Evidence of connections between cere-brospinal fluid and nasal lymphatic vessels in humans,non-human primates and other mammalian species.Cerebrospinal Fluid Res 2004; 1: 2

12 Johnston M, Zakharov A, Koh L, Armstrong D. Subarach-noid injection of Microfil reveals connections betweencerebrospinal fluid and nasal lymphatics in the non-human primate. Neuropathol Appl Neurobiol 2005; 31:632–40

13 Nagra G, Koh L, Zakharov A, Armstrong D, Johnston M.Quantification of cerebrospinal fluid transport across thecribriform plate into lymphatics in rats. Am J Physiol RegulIntegr Comp Physiol 2006; 291: R1383–9

14 Paxinos G, Watson C. The Rat Brain in Stereotaxic Coordi-nates, 3rd edn. San Diego, CA: Academic Press, 1997

15 Henriksson L, Voigt K. Age-dependent differences of dis-tribution and clearance patterns in normal RIHSA cister-nograms. Neuroradiology 1976; 12: 103–7

16 Welch K, Friedman V. The cerebrospinal fluid valves.Brain 1960; 83: 454–69

17 Welch K, Pollay M. Perfusion of particles through arach-noid villi of the monkey. Am J Physiol 1961; 201: 651–4

18 McComb JG, Davson H, Hyman S, Weiss MH. Cerebrospi-nal fluid drainage as influenced by ventricular pressure inthe rabbit. J Neurosurg 1982; 56: 790–7

19 McComb GJ, Hyman S. Lymphatic drainage of cere-brospinal fluid in the cat. In: Hydrocephalus. Eds KShapiro, A Marmarou, H Portnoy. New York: RavenPress, 1984; 83–97

20 McComb GJ, Hyman S. Lymphatic drainage of cere-brospinal fluid in the primate. In: Pathophysiology of theBlood-Brain Barrier. Eds BB Johansson, H Widner. NewYork: Elsevier Science Publishers, 1990; 421–37

21 Boulton M, Flessner M, Armstrong D, Hay J, Johnston M.Lymphatic drainage of the CNS: effects of lymphaticdiversion/ligation on CSF protein transport to plasma.Am J Physiol 1997; 272: R1613–19

22 Boulton M, Young A, Hay J, Armstrong D, Flessner M,Schwartz M, Johnston M. Drainage of CSF throughlymphatic pathways and arachnoid villi in sheep:



153

measurement of 125I-albumin clearance. NeuropatholAppl Neurobiol 1996; 22: 325–33

23 Boulton M, Flessner M, Armstrong D, Mohamed R, Hay J,Johnston M. Contribution of extracranial lymphatics andarachnoid villi to the clearance of a CSF tracer in the rat.Am J Physiol 1999; 276: R818–23

24 Boulton M, Flessner M, Armstrong D, Hay J, Johnston M.Determination of volumetric cerebrospinal fluid absorp-tion into extracranial lymphatics in sheep. Am J Physiol1998; 274: R88–96

25 Kalmey JK, Thewissen JG, Dluzen DE. Age-related sizereduction of foramina in the cribriform plate. Anat Rec1998; 251: 326–9

26 Doty RL. Influence of age and age-related diseases onolfactory function. Ann N Y Acad Sci 1989; 561: 76–86

27 Moinuddin SM, Tada T. Study of cerebrospinal fluid flowdynamics in TGF-beta 1 induced chronic hydrocephalicmice. Neurol Res 2000; 22: 215–22

28 Tada T, Zhan H, Tanaka Y, Hongo K, Matsumoto K,Nakamura T. Intraventricular administration of hepato-cyte growth factor treats mouse communicating hydro-cephalus induced by transforming growth factor beta1.Neurobiol Dis 2006; 21: 576–86

29 Lemaitre H, Crivello F, Grassiot B, Alperovitch A, TzourioC, Mazoyer B. Age- and sex-related effects on the neu-roanatomy of healthy elderly. NeuroImage 2005; 26:900–11

30 Matsumae M, Kikinis R, Morocz IA, Lorenzo AV, SandorT, Albert MS, Black PM, Jolesz FA. Age-related changes inintracranial compartment volumes in normal adultsassessed by magnetic resonance imaging. J Neurosurg1996; 84: 982–91

31 Pakulski C, Drobnik L, Millo B. Age and sex as factorsmodifying the function of the blood–cerebrospinal fluidbarrier. Med Sci Monit 2000; 6: 314–18

32 May C, Kaye JA, Atack JR, Schapiro MB, Friedland RP,Rapoport SI. Cerebrospinal fluid production is reduced inhealthy aging. Neurology 1990; 40: 500–3

33 Redzic ZB, Preston JE, Duncan JA, Chodobski A,Szmydynger-Chodobska J. The choroid plexus-cerebrospinal fluid system: from development to aging.Curr Top Dev Biol 2005; 71: 1–52

34 Serot JM, Bene MC, Faure GC. Choroid plexus, aging ofthe brain, and Alzheimer’s disease. Front Biosci 2003; 8:s515–21

35 Bozanovic-Sosic R, Mollanji R, Johnston MG. Spinal andcranial contributions to total cerebrospinal fluid trans-port. Am J Physiol Regul Integr Comp Physiol 2001; 281:R909–16

36 Silver I, Kim C, Mollanji R, Johnston M. Cerebrospinalfluid outflow resistance in sheep: impact of blocking cere-brospinal fluid transport through the cribriform plate.Neuropathol Appl Neurobiol 2002; 28: 67–74

37 Mollanji R, Bozanovic-Sosic R, Silver I, Li B, Kim C, MidhaR, Johnston M. Intracranial pressure accommodation isimpaired by blocking pathways leading to extracraniallymphatics. Am J Physiol Regul Integr Comp Physiol 2001;280: R1573–81

38 Barber BJ, Dutta S, Parameswaran S, Babbitt RA. Age-related changes in perimicrovascular protein distribu-tion. Am J Physiol 1995; 269: H1213–20

39 Weller RO, Kida S, Zhang ET. Pathways of fluid drainagefrom the brain – morphological aspects and immunologi-cal significance in rat and man. Brain Pathol 1992; 2:277–84

40 Caversaccio M, Peschel O, Arnold W. The drainage ofcerebrospinal fluid into the lymphatic system of the neckin humans. ORL J Otorhinolaryngol Relat Spec 1996; 58:164–6

41 Csanda E, Obal F, Obal F Jr. Central nervous system andlymphatic system. In: Lymphangiology. Eds M Foldi, JCasley-Smith. New York: F.K. Schattauer Verlag, 1983;475–508

42 Lowhagen P, Johansson BB, Nordborg C. The nasalroute of cerebrospinal fluid drainage in man. A light-microscope study. Neuropathol Appl Neurobiol 1994; 20:543–50

43 Duong DH, O’malley S, Sekhar LN, Wright DG. Postop-erative hydrocephalus in cranial base surgery. Skull BaseSurg 2000; 10: 197–200

44 Bradbury MW, Cserr HF. Drainage of cerebral interstitialfluid and of cerebrospinal fluid into lymphatics. In:Experimental Biology of the Lymphatic Circulation. Ed. MJohnston. New York: Elsevier Science, 1985; 355–94

45 Weller RO, Massey A, Newman TA, Hutchings M, KuoYM, Roher AE. Cerebral amyloid angiopathy: amyloidbeta accumulates in putative interstitial fluid drainagepathways in Alzheimer’s disease. Am J Pathol 1998; 153:725–33

46 Weller RO, Yow HY, Preston SD, Mazanti I, Nicoll JA.Cerebrovascular disease is a major factor in the failure ofelimination of Abeta from the aging human brain: impli-cations for therapy of Alzheimer’s disease. Ann N Y AcadSci 2002; 977: 162–8

Received 13 December 2006Accepted after revision 16 March 2007



154

Extracellular fluid systems in the brain and the ... · Extracellular fluid systems in the brain...

Documents

Transcript of Extracellular fluid systems in the brain and the ... · Extracellular fluid systems in the brain...