{'fraction_non_alphanumeric': 0.03570187922320958, 'fraction_numerical': 0.027224643933706116, 'mean_word_length': 4.993451761308305, 'pattern_counts': {'":': 0, '<': 2, '<?xml version=': 0, '>': 2, 'https://': 0, 'lorem ipsum': 0, 'www.': 0, 'xml': 0}, 'pii_count': 0, 'substrings_counts': 0, 'word_list_counts': {'cursed_substrings.json': 3, 'profanity_word_list.json': 0, 'sexual_word_list.json': 0, 'zh_pornsignals.json': 0}}
Aquaporin Expression, Regulation and Function in the Intervertebral Disc

Intervertebral disc degeneration-associated low back pain is a debilitating condition with no current treatments directed towards halting or reversing the degenerative cascade at a cellular level. The lack of such treatments is in part due to an incomplete knowledge of the molecular mechanisms that govern IVD function in health and degeneration. Due to the unique location and role of the IVD within the spine, many factors contribute to the microenvironment that cells reside within. The survival and function of cells has been irrefutably linked to their ability to adapt to the microenvironment in which they live. However, it is not completely understood how IVD cells have been able to survive and adapt to their environment. 
The intervertebral disc is a highly hydrated tissue; the rich proteoglycan matrix imbibes water, enabling the disc to withstand compressive loads. During ageing and degeneration increased matrix degradation leads to dehydration and loss of function. Aquaporins are a family of transmembrane channel proteins that selectively allow the passage of water in and out of cells and are responsible for maintaining water homeostasis in many tissues; hence many AQPs are potentially expressed by cells within the intervertebral disc to enable their adaptation to this highly hydrated tissue. 
The aim of this thesis was to investigate the expression, regulation and function of AQP transmembrane water channels within the IVD and how they potentially contribute to the adaptation of cells to their environment. Results have highlighted NP cells express many AQP water channels in vivo, whose expression may be altered between disc development and degeneration. AQP1 and 5 were found to be upregulated by TonEBP in hyperosmotic conditions, a transcription factor controlling osmotic adaptation and matrix expression, which may implicate them in the adaptation of NP cells to their environment, which becomes unachievable when AQP1 and 5 are decreased during degeneration. AQP4 and TRPV4 function in NP cells was required for fundamental cellular processes such as cell volume regulation and water permeability, enabling adaptation to their osmotically fluxing environment. Finally, it was identified that other microenvironmental factors also contribute to AQP expression in NP cells, indicating the regulation and function of these water channels is potentially very complex. 
Together, investigations presented in this thesis have demonstrated many AQPs are expressed by the intervertebral disc and enable NP cells to respond and adapt to their environment, ultimately contributing to the overall function of the tissue. Their regulation by multiple environmental factors may signify that AQPs have diverse roles within the IVD, which remain to be elucidated. Importantly, this body of work has contributed novel findings, increased knowledge and opened new avenues of research in the field of IVD and spine biology.


Abstract
Intervertebral disc degeneration-associated low back pain is a debilitating condition with no current treatments directed towards halting or reversing the degenerative cascade at a cellular level. The lack of such treatments is in part due to an incomplete knowledge of the molecular mechanisms that govern IVD function in health and degeneration. Due to the unique location and role of the IVD within the spine, many factors contribute to the microenvironment that cells reside within. The survival and function of cells has been irrefutably linked to their ability to adapt to the microenvironment in which they live. However, it is not completely understood how IVD cells have been able to survive and adapt to their environment.
The intervertebral disc is a highly hydrated tissue; the rich proteoglycan matrix imbibes water, enabling the disc to withstand compressive loads. During ageing and degeneration increased matrix degradation leads to dehydration and loss of function. Aquaporins are a family of transmembrane channel proteins that selectively allow the passage of water in and out of cells and are responsible for maintaining water homeostasis in many tissues; hence many AQPs are potentially expressed by cells within the intervertebral disc to enable their adaptation to this highly hydrated tissue.
The aim of this thesis was to investigate the expression, regulation and function of AQP transmembrane water channels within the IVD and how they potentially contribute to the adaptation of cells to their environment. Results have highlighted NP cells express many AQP water channels in vivo, whose expression may be altered between disc development and degeneration. AQP1 and 5 were found to be upregulated by TonEBP in hyperosmotic conditions, a transcription factor controlling osmotic adaptation and matrix expression, which may implicate them in the adaptation of NP cells to their environment, which becomes unachievable when AQP1 and 5 are decreased during degeneration. AQP4 and TRPV4 function in NP cells was required for fundamental cellular processes such as cell volume regulation and water permeability, enabling adaptation to their osmotically fluxing environment. Finally, it was identified that other microenvironmental factors also contribute to AQP expression in NP cells, indicating the regulation and function of these water channels is potentially very complex.
Together, investigations presented in this thesis have demonstrated many AQPs are expressed by the intervertebral disc and enable NP cells to respond and adapt to their environment, ultimately contributing to the overall function of the tissue. Their regulation by multiple environmental factors may signify that AQPs have diverse roles within the IVD, which remain to be elucidated. Importantly, this body of work has contributed novel findings, increased knowledge and opened new avenues of research in the field of IVD and spine biology.                                      The IVD is composed of the outer annulus fibrosus with lamellar fibrocartilaginous structure and the inner gelatinous nucleus pulposus. IVDs are attached to adjacent vertebrae via cartilaginous endplates. B and C show the posterior spinous processes and the spinal cord which travels protected through the vertebral foramen. Adapted from (Medline, 2018) 25

The intervertebral disc
The IVD is a fibrocartilaginous structure positioned between each of the 24 movable vertebrae. The main function of the IVD is mechanical; it enables the movement of the head and spine and accommodates the applied loading of the body.
The IVD is comprised of 3 distinct regions; cartilaginous endplates (CEP), annulus fibrosus (AF) and nucleus pulposus (NP) (Figure 1.1). These regions within the IVD all work in unison to withstand mechanical loading and disperse energy evenly throughout the spine.

Cartilaginous endplates
Both the superior and inferior ends of the IVD are covered by a thin layer of hyaline cartilage. The cartilaginous endplate (CEP) has maximal thickness at birth which thins during aging, in adults it is approximately 0.5 -1mm in width. They act as an interface between the internal IVD structures and the vertebral bone and prevent pressure being applied directly between both. Chondrocytes reside within the CEP embedded into an aggrecan and collagen type II rich matrix, which gives the CEP a structure similar to articular cartilage. The central region of the CEP contains the highest density of cells throughout the IVD, with approximately 18 x 10 6 cells/cm 3 in the young adult, these cells do not undergo osteochondral differentiation (Johnson et al., 2006;Liebscher et al., 2011). The CEP transitions into bone through a layer of calcified cartilage. The CEP is permeable to small molecules and thus also plays a role in the viability and metabolism of cells within the IVD, as it facilitates the diffusion of 26 nutrients and oxygen between the subchondral vertebral bone and the internal structures of the IVD which are avascular, except for the outer AF. The mechanical properties of the CEP may be altered due to weakening or mineralisation of the cartilage, which can lead to fractures and cause NP tissue to be exuded into the bone (Wu, Morrison and Schweitzer, 2006). These phenomena are called Schmorl's nodes.

Annulus fibrosus
The AF can be divided into 2 regions; the outer and inner AF. The outer AF is composed of bundled collagen type I fibres that are orientated into long parallel lamellae. The collagen fibres within each lamella run in parallel at a 65° angle to the adjacent lamellae. The fibres of the outer AF are anchored to the vertebral bone ( Figure 1.2). This structural formation provides the outer AF with the ability to resist tensile forces from bending and twisting of the spine. The inner AF serves as a transition zone between the highly ordered outer AF and the disorganised NP, and characteristics of both tissues are observed. Aggrecan is found in both AF regions, whereas decorin and biglycan are mainly found within the outer AF. The inner AF contains collagens types I and II whilst the outer AF contains mainly collagen type I.
Elastin accounts for 2% of the dry weight of the entire AF and forms inter-lamellar bridges. There are also cellular differences between the AF regions; outer AF cells are fibroblast-like; elongated and fusiform, whereas inner AF cells are spherical in shape and resemble the morphology of NP cells and chondrocytes. The cell density of mature AF tissue is approximately 9 x 10 6 cells/cm 3 (Roughley, 2004). The inner AF is also attached to the CEP, whilst the OAF is attached to vertebral bone. Schematic of the organisation of concentric AF lamellae. The direction of collagen fibres alternates at a 65° angle between adjacent lamellae. Adapted from (Elliott and Setton, 2000).

Nucleus pulposus
The NP is composed of mainly water (70 -80%), randomly arranged collagen type II fibres (Figure 1.3), radially arranged elastin and small amounts of collagen types VI, IX and XI; collagen type X has also been observed within the NP during IVD degeneration and is linked to calcification (Grant et al., 2016). These components are all embedded within a matrix rich in proteoglycans, which make up to 30 -50% of the dry weight of the tissue (Figure 1.3). The main proteoglycan matrix component is aggrecan which links to hyaluronan tethering it to the tissue (Figure 1.3); other proteoglycans such as versican, perlecan, lubricin and small leucine-rich repeat proteoglycans are also present within the NP (Inkinen et al., 1998;Roughley, 2004).
Crosslinked collagen within the NP confers tensile strength, which is reduced during ageing and degeneration as crosslinks between collagen fibrils decreases (Antoniou et al., 1996). Proteoglycans and their negatively charged glycosaminoglycans (GAG) side chains, such as chondroitin sulphate and keratan sulphate, draw in water and cations via osmosis, meaning the NP exhibits high osmotic and hydrostatic pressure (Figure 1.3). This provides the NP with viscoelasticity and resistance to compression (Doege et al., 1991;Dolan and Adams, 2001). As the NP has no blood supply, the oxygen tension is very low, prompting NP cells to rely on the glycolytic pathway for energy metabolism (Agrawal et al., 2007).

30
The NP is derived from the notochord and during development the NP is highly cellular. During development and at birth the NP contains clusters of large cells with vacuolated (25 -85µm) morphology, called notochordal cells (Figure 1.4). During aging the population of notochordal cells (and the total cell number in general) decreases and is replaced by smaller, spherical, non-vacuolated, chondrocyte-like cells (NP cells) (Figure 1.4). However, it has been shown that the proteoglycan to collagen ratio in the NP of healthy young adults (15 -25 years) is much higher (27:1) when compared to articular cartilage (2:1), indicating phenotypic differences between NP cells and chondrocytes (Mwale, Roughley and Antoniou, 2004). In the mature NP, single cells encased within pericellular matrix lacunae are present, along with cell clusters that share a single lacuna (Johnson et al., 2006). The cell density is approximately 4 x 10 6 cells/cm 3 (Roughley, 2004;Chen, Yan and Setton, 2006). It has been determined that during maturation notochordal cells undergo differentiation into the smaller chondrocyte-like NP cells found in the adult IVD (Choi, Cohn and Harfe, 2008;Minogue et al., 2010bMinogue et al., , 2010aMcCann et al., 2012;Séguin et al., 2018). Yet, there is also evidence that potentially notochordal cells die and are replaced by cells migrating from adjacent tissues as the IVD matures (Kim et al., 2009;Yang et al., 2009;Séguin et al., 2018). Therefore, considerable work has been done to elucidate the NP phenotype and identify genetic markers to distinguish NP cells from AF cells and chondrocytes. Many markers have been proposed; genes involved during the development of the notochord such as sonic hedgehog (SHH) (Choi, Lee and Harfe, 2012) and brachyury (T) (Minogue et al., 2010b), proteins linked to the physiology of NP cells such as (HIF-1α) , glucose transporter 1 (GLUT-1) (Rajpurohit et al., 2002), carbonic anhydrase III and XII (CAIII and CAXII) (van den Akker et al., 2014), and other proteins like the intermediate filaments cytokeratin 18 and 19 Rodrigues-Pinto et al., 2016) and cell surface marker cluster of differentiation 24 (CD24) (Rutges et al., 2010;Rodrigues-Pinto et al., 2016). A consensus paper, authored by many groups, was published that recommended using a panel of these markers to determine the NP phenotype . However, since this publication other potential markers have been identified (Rodrigues-Pinto et al., 2016;, indicating that a true phenotypic marker differentiating NP cells remains to be discovered. Also, the panel of markers recommended by Risbud et al., 2015 was focussed on determining the young, healthy NP cell phenotype, yet Thorpe et al.,33 2016 identified different markers that distinguished mature, aging NP cells from AF cells and chondrocytes, indicating that factors such as NP maturity and the state of degeneration must also be considered when attempting to phenotypically characterise this elusive cell type.

Development of the intervertebral disc
The IVDs are derived from two embryonic structures called the notochord and the sclerotome (Peacock, 1951;Paavola, Wilson and Center, 1980). The central NP region is derived from the notochord (Peacock, 1951;Smits and Lefebvre, 2003)  In mammals, an acellular sheath containing proteoglycans, collagens and laminins surrounds the notochord (Götz, Osmers and Herken, 1995). During embryonic development the notochord acts as a primitive support system and signalling centre, directing the development and differentiation of surrounding tissues and structures, including the neural tube  and sclerotome (Fan and Tessier-Lavigne, 1994), by secreting growth factors and morphogens (Christ, Huang and Scaal, 2004;Nimmagadda et al., 2007;Christ and Scaal, 2008) (Figure 1.6).
The sclerotome is derived from the somites, transient structures adjacent to the notochord and neural tube during vertebrate development (Wilting et al., 1994).
In response to Wnt signalling from the notochord and neural tube floor plate, and SHH signalling from the ectoderm, the somites undergo compartmentalisation (Christ and Scaal, 2008;McCann and Séguin, 2016). The development of the sclerotome is characterised by the formation of the ventral, lateral and dorsal subcompartments from the somites (Monsoro-Burq et al., 1994;Peters et al., 1999). The dorsal sub-compartment differentiates and forms the dermomyotome, which generates the connective tissues of the axial skeleton (Capdevila, Tabin and Johnson, 1998). The ventral sub-compartment differentiates into the sclerotome, consisting of PAX-1-expressing cells, and produces the vertebral bodies, AF and CEP of IVDs (Peters et al., 1999). In addition, each sclerotomal segment demonstrates rostral to caudal polarity; anterior vertebral structures are formed from the caudal sclerotome, whereas posterior vertebral structures are formed from the rostral sclerotome (Christ and Scaal, 2008).
As the vertebrae, AF and CEP form, the notochord contracts from the vertebral bodies, notochordal cells are expelled from vertebral regions and are limited to the centre of the forming IVD, which then expand to produce NP regions (Alkhatib et al., 2018). The notochordal sheath contains and directs the retreating notochord and its internal hydrostatic pressure, which is essential for notochordal expansion and the subsequent formation of the NP (Adams, Keller and Koehl, 1990;Alkhatib et al., 2018).
In humans, the number of notochordal cells within the notochord and subsequent NP starts to decline during maturation, even during embryonic development (Peacock, 1951). The amount of notochordal cells continues to decline until they are virtually absent within the NP after the first decade of life (Risbud, Schaer and Shapiro, 2010). However, it has been observed in other species, such as rats and mice, that notochordal cells are still present beyond the point of skeletal maturity Duncan, 2003a, 2004a). Curiously, chondrodystrophic (CD) dogs (long bodies, short legs) lose notochordal cell populations very early in life and suffer frequent, early-onset IVD degeneration, whereas non-chondrodystrophic (NCD) dogs (bodies and limbs in proportion) retain notochordal cell populations into adulthood and suffer infrequent, later-onset IVD degeneration Smolders et al., 2013). This has led to canine studies determining a link between the decline of notochordal cells and IVD degeneration Duncan, 2003, 2004). The exact underlying mechanisms responsible for the differentiation of notochordal cells into mature NP cells has not yet been elucidated, yet progressive alterations to the mechanical loading of the IVD (and the changes to the microenvironment thereafter) during development and growth are likely to influence the phenotype of notochordal and NP cells (Navaro et al., 2015;Palacio-Mancheno et al., 2018). It has also been shown that notochordal cells require very specific culture conditions to retain their phenotype in vitro (Spillekom et al., 2014).   (Smits and Lefebvre, 2003).

The microenvironment of the intervertebral disc
Due to the unique development and positioning of the IVD within the human body, many environmental factors contribute to the physiology of the tissue and influence cellular behaviour.

Biomechanics
The function of the IVD is to enable movement and withstand the compressive forces of the spine; hence the IVD has a loaded environment subjected to a number of physical stresses. The biomechanics of the IVD is determined by 38 interactions of loading patterns between all the structures of the IVD. The IVD can withstand axial compression, torsion, flexion/extension, lateral bending and disperse mechanical energy evenly throughout the spine (Setton and Chen, 2004).
Due to the high level of hydration and gelatinous consistency the NP is considered a viscoelastic material, demonstrating both fluid and solid-like behaviours (Iatridis et al., 1999). Internal stresses and strains exist within the NP, even in the unloaded IVD, which arise from the water content resulting in a high hydrostatic pressure with a baseline of 0.1 -0.2MPa when supine, approximately 0.5MPa when standing and increasing up to 3 -4MPa during extreme physical activity (Wilke et al., 1999). Due to the variation of physical activity and posture during day and night the NP undergoes a diurnal change in intradiscal pressure . Under increased load, during load-bearing and vertical posture, the IVD deforms which causes an increase in hydrostatic pressure and fluid is slowly expelled from the NP . When in the supine position, the reduced load causes a release in intradiscal pressure and fluid is re-imbibed into the NP (McMillan, Garbutt and Adams, 1996;Malko, Hutton and Fajman, 2002). During compression the increase in NP pressure is constrained by the lamellae of the AF which produces longitudinal and circumferential AF tension (Galante, 1967). Therefore, the predominant mechanical stimuli acting on AF cells are tensile strain and shear (Galante, 1967). Whereas the predominant mechanical stimuli acting on NP cells are compressive and shear stresses, alongside hydrostatic pressure due to the water content and swelling properties of the NP .

39
As IVD cells are exposed to mechanical stimuli, it has been determined that the type, magnitude, frequency and duration of applied loading are crucial in determining cellular responses (Maclean et al., 2004;Wang, Jiang and Dai, 2007;Wuertz et al., 2009;Walter et al., 2011). IVD cells respond to mechanical stimuli by activating mechanotransduction signalling pathways. The loss of aggrecan expression by NP cells due to a decrease in hydrostatic pressure was inhibited by pre-treatment with RGD blocking peptides (Le ). However, this was only observed in NP cells derived from healthy IVD tissue, not NP cells derived from degenerate IVD tissue (Le ). This suggests not only that NP cells respond to pressure via RGD domains and integrin signalling, but also that mechanotransduction pathways are altered during IVD degeneration (Le . Healthy physiological loading, such as dynamic low pressure loads (0.1 -1.0MPa), induces matrix synthesis and is essential for matrix production and IVD homeostasis (Wuertz et al., 2007;Neidlinger-Wilke et al., 2012). In contrast, overloading (1 -4MPa) has been demonstrated to evoke degenerative changes, increasing catabolic gene expression, leading to the destruction of matrix (Iatridis et al., 1999;Korecki, MacLean and Iatridis, 2008). Thus, indicating that the IVD has adapted to its function of withstanding the compressive forces of the spine and that IVD cell biology is finely tuned by the biomechanical forces exerted upon them.

Osmolality
The extracellular matrix (ECM) of the IVD enables its mechanical function.
Proteoglycans and their negatively charged side chains draw water and ions into the 40 NP, increasing the hydrostatic pressure, so the IVD can withstand compression . These actions simultaneously increase the fixed charge density and osmotic pressure within the NP, increasing the intradiscal osmolality when compared to most other tissues (400 -500mOsm/kg H2O) (Urban and McMullin, 1985;McMillan, Garbutt and Adams, 1996). Similar to responses to mechanical loading, culturing NP cells in physiological conditions mimicking the osmolality of the healthy IVD increases the expression of matrix genes (Wuertz et al., 2007;Neidlinger-Wilke et al., 2012;O'Connell, Newman and Carapezza, 2014) via osmotically regulated signalling pathways (Tsai et al., 2006;Johnson, Shapiro and Risbud, 2014b). However, these signalling pathways may become altered when the osmolality decreases, mimicking the degenerate IVD (~300mOsm/kg H2O) when proteoglycan content decreased (Ishihara et al., 1997). Which may lead to the production of catabolic genes in preference over matrix genes (Johnson, Shapiro and Risbud, 2014b;Johnson et al., 2017), indicating that NP function is controlled by the osmotic environment bestowed upon the IVD by proteoglycans and mechanical loading. There are numerous signalling pathways, transcription factors and transmembrane channel proteins expressed in NP cells that are implicated in their adaptation to the hyperosmotic environment of the IVD, yet how NP cells sense and respond to this stimulus has not been fully elucidated (Johnson, Shapiro and Risbud, 2014b;Sadowska et al., 2018).

Nutrition, pH and oxygen tension
The IVD is the largest avascular tissue in the human body (Urban, 2002). In healthy IVDs, blood vessels only penetrate into the periphery of the outer AF and NP cells are 6 -8mm away from the nearest blood supply. Therefore, the transport of nutrients and waste products occurs via diffusion (Urban, 2002). Small amounts of diffusion take place via the blood vessels at the AF periphery, but the majority of transport occurs from blood vessels within the vertebral bodies and across the CEP (Roberts et al., 1996).
Nutrient transport through the IVD occurs almost exclusively by diffusion facilitated by load-induced water transport. This results in steep concentration gradients of glucose, oxygen and lactic acid across the IVD (Roberts et al., 1996;Urban, 2002;Bibby et al., 2005). As the concentration of glucose and oxygen progressively decreases towards the centre of the IVD, the concentration of lactic acid increases (Ohshima and Urban, 1992). Therefore, the NP has a slightly acidic pH (pH6.9 -7.2) and low oxygen microenvironment (8 -10% at the IVD-vertebral interface decreasing to 0.3 -0.5% in the NP) that cells are required to adapt to (Holm et al., 1981;Urban, 2002). Oxygen concentration has been observed at 0.7% in degenerate human IVDs (Bartels et al., 1998). Consequently, NP cells favour glycolytic metabolism (causing the increase of lactic acid) due to the lack of oxygen (Agrawal et al., 2007) and have adapted to their acidic surroundings (Urban, 2002;Silagi et al., 2018a;Silagi et al., 2018b). Such adaptation is mediated by hypoxia-inducible factor-1α (HIF-1α), which has been shown to support the survival and function of NP cells 42 by driving glycolytic metabolism and ECM production, enabling adaptation to their hypoxic niche (Pfander et al., 2003;Risbud et al., 2006;Fujita et al., 2012;. The nutrient supply into the IVD can be reduced by a number of factors such as CEP calcification (Roberts et al., 1996), decreased blood flow to segmental arteries (Kauppila, 1997) or a decrease in the water content of the IVD (Horner and Urban, 2001). The decrease in nutrient supply has been linked to IVD degeneration due to an increase in cell death, caused by decreases in glucose concentration (< 0.5mmol/L) and pH (< 6.5) (Horner and Urban, 2001), and the production of catabolic enzymes that mediate the destruction of ECM (Urban and Roberts, 2003). However, it may also be possible that the nutrient supply can be increased during IVD degeneration, due to a possible increase in vascularisation (which may also cause pain due to associated nerve ingrowth) Binch et al., 2014) and the presence of fissures, allowing rapid nutrient diffusion (Melrose et al., 2002;Stefanakis et al., 2012;Binch et al., 2015).
Recently LBP was found to be one of the 5 top causes of disability worldwide (Vos et al., 2017;James et al., 2018). LBP is multifactorial, yet 40% of cases are attributed to IVD degeneration (Luoma et al., 2000). The aetiology of IVD degeneration itself is also multifactorial; aging (Gruber and Hanley, 1998), smoking (Battié et al., 1991), genetic factors (Matsui et al., 1998), decreased nutrient uptake and waste removal (Horner and Urban, 2001;Bibby et al., 2005) and repetitive mechanical loading greater than tolerable limits (Adams et al., 2000). IVD degeneration is characterised by progressive destruction of the ECM caused by altered cell metabolism and function, matrix synthesis and degradation of matrix components (Le .

Cytokines
IVD cells produce a plethora of cytokines and chemokines (Le Maitre, Freemont and Hoyland, 2005;Hoyland, Le Maitre and Freemont, 2008;Phillips et al., 2013Phillips et al., , 2015, interleukin -1β (IL-1β) and tumour necrosis factor -α (TNF-α) are the most studied and are clearly implicated in the pathogenesis of IVD degeneration. IL-1β, IL-1 receptor I (IL-1RI) and IL-1 receptor antagonist (IL-1Ra) are all expressed in healthy IVD cells, but the expression of IL-1β and IL-1RI (not IL-1Ra) is increased with increasing severity of degeneration (Le Maitre, Freemont and Hoyland, 2005)and has been shown to induce many catabolic events (Le Maitre, Freemont and Hoyland, 2005;Hoyland, Le Maitre and Freemont, 2008). TNF-α expression is also increased during IVD degeneration, but not to as high a level as IL-1β (Le Maitre, Hoyland and Freemont, 2007a). Also, as TNF receptor I (TNF-RI) was not increased in degenerate IVDs and TNF-α inhibition has no effect on matrix degradation, whereas IL-1β inhibition completely abrogated matrix degradation, this may highlight the prominent role of IL-1β, over TNF-α, during IVD degeneration (Hoyland, Le Maitre and Freemont, 2008). The importance of IL-1β during degeneration has been strengthened by the observation of spontaneous IVD degeneration in a knockout mouse model of IL-1Ra, the natural inhibitor of IL-1β (Phillips et al., 2013), and the presence of polymorphisms in the IL-1 gene cluster linked to an increased risk of IVD degeneration (Solovieva et al., 2004(Solovieva et al., , 2006Karppinen et al., 2009). Conversely, an increase in both TNF-α receptors (TNF-RI and TNF-RII) expression has been identified in herniated IVD tissue and showed a positive correlation with pain levels assessed by the visual analogue scale (Andrade et al., 2011), indicating TNF-α also plays an important role during IVD degeneration and possibly in the manifestation of pain.
Recently the overexpression of TNF-α in a transgenic mouse model showed the early onset of IVD herniation, yet no overt signs of degeneration (Gorth, Shapiro and Risbud, 2019), which supports that TNF-α is involved in IVD degeneration, but less so than the pivotal role played by IL-1β. These changes in cellular behaviour initiate IVD degeneration by leading to the increased production of more cytokines, vascularisation and innervation causing pain, and the increased expression of matrix degrading enzymes.

Degeneration of extracellular matrix
As IVD degeneration advances, collagen type II within the NP is gradually replaced by the more fibrous collagen type I (Roughley, 2004). The overall composition of proteoglycans is reduced and altered with decreased aggrecan expression and versican, decorin and biglycan expression is increased, thus reducing the capacity of the NP to imbibe water, resulting in decreased hydration and loss of 45 IVD height (Inkinen et al., 1998;Cs-Szabo et al., 2002). Concurrently, the increased cytokine production by IVD cells upregulate a variety of catabolic mediators including matrix metalloproteinases (MMP) -1, -2, -3, -13 and -14, and a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS) -1, -4 and -5 (Séguin et al., 2006;Pockert et al., 2009;Tian et al., 2013).
MMPs and ADAMTSs are important mediators of matrix remodelling during healthy physiology, but their overexpression during degeneration (Roberts et al., 2001;Le Maitre, Freemont and Hoyland, 2004;Pockert et al., 2009) brings about the destruction of collagen and proteoglycans within the NP, which is exacerbated by the suppression of important matrix genes during these processes Tian et al., 2013;Johnson et al., 2017). The resulting compositional changes during degeneration cause alterations to the IVD microenvironment, such as decreased nutrition, pH and osmolality and excessive biomechanical loading (Section 1.4) ( Figure 1.7). This consequently leads to adverse cellular changes, including apoptosis/necrosis (Gruber and Hanley, 1998;Ding, Shao and Xiong, 2013), senescence (Roberts et al., 2006;Gruber et al., 2007; and a decrease in the viability and number of remaining cells (Le . As the IVD has developed a hostile microenvironment during degeneration, those cells still present are also more likely to develop a catabolic phenotype, continuing the degenerative cascade. Microenvironmental factors within the IVD that dictate cellular function and how these factors are altered during degeneration.

Consequences of intervertebral disc degeneration
The imbalance between normal matrix synthesis and degradation results in a loss of the structural integrity of the NP due to a reduction in hydration, osmotic swelling pressure and IVD height (Urban and McMullin, 1988). These changes result in a diminished ability of the IVD to withstand and evenly distribute the compressive loading of the spine. This may ultimately lead to IVD injuries such as the formation of tears and fissures through the AF, due to the asymmetric distribution of compressive loading, and the eventual herniation and extrusion of the NP through such tears (Adams and Roughley, 2006) (Figure 1.8). IVD herniation may also be associated with the sensation of chronic pain as extruded NP material may impinge on spinal nerves (Risbud and Shapiro, 2014) (Figure 1.8), which may also act as an access route for the vascularisation and innervation of the IVD (Binch et al., 2014). showing posterior L5-S1 IVD herniation (red arrow) in a patient with symptomatic IVD degeneration. Adapted from Risbud and Shapiro, 2014.

Current treatment
Most current practices for the treatment of LBP are directed towards alleviating and managing symptoms, including analgesic medication, physiotherapy, acupuncture and chiropractic treatment (Chou et al., 2009). However, as these practices only be able to treat symptoms such as pain, they may only be successful in the short term (Carette et al., 1997). Other options for patient treatment include surgical procedures to stabilise or remove damaged tissue. Microdiscectomy, the removal of herniated NP tissue, and spinal fusion, the removal of the degenerated IVD and vertebral fusion, can be offered to patients when conservative methods have been unsuccessful (Andrews and Lavyne, 1990). However, these treatments have been associated with alterations in spinal biomechanics and the potential degeneration of adjacent IVDs (Lund and Oxland, 2011).
It has also been reported that approximately 23% of patients receiving microdiscectomy required repeat operations, due to recurrent IVD herniation (Parker et al., 2015).
Emerging research has aimed to develop a biological approach to address the underlying pathophysiology of IVD degeneration. The use of growth factors and/or inhibitors has been investigated in attempts to promote anabolic processes, such as matrix synthesis, whilst also inhibiting catabolic processes, such as matrix degradation and inflammation (Tim Yoon et al., 2003;Christine L Le Maitre, Hoyland and Freemont, 2007;Cao et al., 2012;Gorth et al., 2012;Feng et al., 2017;Li et al., 2017;Mern et al., 2017). However, so far, the use of such modulating compounds/proteins has failed to substantially treat dysfunctional NP cells which continue to display a catabolic phenotype (Roberts et al., 2006;Gruber et al., 2007;.

50
Therefore, further research has focussed on the use of regenerative cell sources, such as mesenchymal stem cells (MSCs), together with biomaterial scaffolds. With the aim of simultaneously repopulating the NP whilst also providing a microenvironment for the preferential differentiation into cells that regenerate the correct matrix and inhibit the degenerative cascade (Le . To enable such treatments to become successful it is essential that the physiology of the IVD is understood with respect to the development and adaptation of NP cells to their multifaceted microenvironment (Thorpe et al., 2018). The elucidation of these adaptations may be essential in determining how healthy NP cells function and how dysregulation may cause or occur during IVD degeneration. Thus, identifying vital physiological traits that must be protected and restored to ensure potential future treatments not only work but are performed on patients that will benefit from them.

Aquaporins
The movement of water throughout cells and tissues is essential for the maintenance of homeostasis in all organisms. In mammals, water can pass freely in and out of cells by diffusion through the plasma membrane as individual molecules, or passively co-transported with other ions and solutes such as Na + and glucose (Loo et al., 1996). However, these modes of water transport cannot account for the rapid and highly selective water movement that has been shown to regulate many processes such as determining urine concentration in the kidneys (Marples et al., 1995), the secretion of saliva (Krane et al., 2001b) and the formation of a fluid-filled cavity alongside oocytes during folliculogenesis (McConnell et al., 2002).

51
It was almost fifty years ago when the existence of water channels was hypothesised due to the drastic decrease in water permeability observed in cells when incubated with mercurial sulfhydryl reagents (Macey and Farmer, 1970). This hypothesis was confirmed with the discovery of the first aquaporin (AQP), AQP1, in erythrocytes and renal tubules (Denker et al., 1988) and their characterisation as a selective water channel within cell membranes (Zeidel et al., 1992;Agre et al., 1993). Since their discovery, 13 members of the mammalian AQP family of membrane channel proteins have been identified (AQP0-12), which are expressed and have a variety of functions in a plethora of different tissues (Day et al., 2014).

Structure
Despite the diverse expression and roles of AQPs throughout different tissues they contain a highly conserved structure relating to their specific function to transport  Each monomer folds to form individual water-transporting pores, therefore each functional AQP homotetramer contains four pores and not one single, central pore ( Figure 1.10). This structural information was first determined through electrondiffraction studies on AQP1 in erythrocytes (Murata et al., 2000), followed by more highresolution imagery determining similar structures for other human AQPs (Horsefield et al., 2008;Ho et al., 2009). AQPs are highly specific in their roles as water channels as the structure of the pore itself determines the type of molecule that can pass through cell membranes via these channels. The majority of human AQPs (0, 1, 2, 4, 5 and 8) are selective for water molecules only, due to two selection stages within the pore. The first is due to the folding 54 of intracellular loop B and extracellular loop E into the pore, forming two half-helical regions, which both contain a conserved amino acid sequence of Asparagine-Proline-Alanine (NPA) (Sui et al., 2001). Since α-helices have a natural dipole, both half-helices focus their positive dipole towards the centre of the channel which creates an electrostatic barrier to protons and cations whilst also aligning the dipole of water molecules as they pass the NPA signature motifs, when entering the AQP pore (de Groot and Grubmüller, 2005) (Figure 1.11).
The second stage of selectivity is due to a constriction region located on the extracellular side of the AQP pore, which contains another conserved amino acid motif.
The aromatic/Arginine (ar/R) motif facilitates selectivity by re-positioning water molecules for their entry into the AQP pore and also contributes to the exclusion of protons (Wu et al., 2009) (Figure 1.11). The remaining AQPs (3, 6, 7, 9, 10, 11 and 12) have the ability to transport other molecules as well as water (Section 1.6.2).

Function
The AQP family of membrane channel proteins have been identified to be highly selective in their transport of water across cell membranes; however, some are known to have the ability of transporting other small molecules in addition to water. AQP6 has been shown to be potentially permeable to ionic molecules as well as being highly permeable to water . Unlike other AQPs that are mainly located within the plasma membrane, AQP6 is localised to intracellular membranes in acid-secreting, intercalated cells of kidney collecting ducts. This exclusive localisation of AQP6 and its potential to transport ionic molecules suggests a role in renal acid-base regulation.
AQPs 3, 7, 9 and 10 have also been shown to transport other small molecules such as glycerol, urea and ammonia (Litman, Søgaard and Zeuthen, 2009). The so-called "aquaglyceroporins" additional function may be due to structural changes in the ar/R selectivity filter lining the water-transporting pore. When the diameter of the AQP pore at the ar/R restriction site is 2.8 Å in diameter this will only allow the passage of water molecules, whereas when point mutations were introduced at both the aromatic and arginine residues (F56A and R195V, respectively) the diameter of the pore was increased to >3.4 Å. This increase led to the conductance of not only water but also other small molecules such as glycerol in AQP1 (Beitz, Wu, et al., 2006;Hub and de Groot, 2008).
Changes to the NPA motif have also been determined in other AQPs that have different functions other than as a selective water transport. The structure of this conserved motif in the "unorthodox" AQPs (11 and 12) has been demonstrated to be altered which may explain their as of yet, unknown role. Tchekneva et al., 2008 highlighted that by substituting a single amino acid within the E-loop NPA box (C227S) in mice gave rise to a phenotype that resembled AQP11 knockout mice, thus suggesting a vital role for C227 in the function of the NPA motif in AQP11. AQP structure is related 56 to their function as selective water channels as highly conserved amino acid sequences within the AQP family allow for such a high specificity for the movement of water molecules through plasma membranes. Changes in these amino acid sequences have led to a number of these channels acquiring altered functions such as the ability to transport a number of different small molecules in addition to water.

Expression and physiological roles
Due to the unique and vital molecular function of AQP channels, they contribute to many physiological processes and are expressed almost ubiquitously throughout the human body ( Figure 1.12). The expression of specific AQP isoforms within specific tissues also alludes to their potential roles. AQPs have been shown to regulate many processes such as cell volume regulation ; cell structure and adhesion (Varadaraj and Kumari, 2018); cell migration and proliferation (Papadopoulos, Saadoun and Verkman, 2008); water homeostasis in the kidney (Hasler et al., 2008); blood-brain barrier function (Clément, Rodriguez-Grande and Badaut, 2018); lens function (Schey et al., 2017); fluid secretion (Ramli et al., 2019); sperm motility (Laforenza et al., 2016;Fujii et al., 2018); and glycerol metabolism (Méndez-Giménez et al., 2018). As such, alteration to AQP expression and function has also been implicated in the pathophysiology of many disorders, including: renal water imbalance (Kortenoeven and Fenton, 2014); brain oedema (Clément, Rodriguez-Grande and Badaut, 2018); cataract formation (Biswas et al., 2014); infertility (Kasimanickam et al., 2017); and cancer metastasis (Marlar et al., 2017) are all, in-part, due to the dysregulation of AQP expression and function. This indicates that aside from passive water transport, AQPs play active roles during the adaptation of many important physiological processes. Their function, however, relies on their integration into the 57 plasma membrane of cells in which their expression is not ubiquitous; the localisation of AQPs within cells is tightly regulated by a number of mechanisms.

Aquaporin gating and channel inhibition
Gating mechanisms in AQPs have long been debated to regulate their function by changing structure of the pore to allow or halt the flow of water through cells. It is well documented that plant AQPs contain gating mechanisms that respond to a number of different stimuli including phosphorylation (Johansson et al., 1998;Nyblom et al., 2009), pH (Tournaire-Roux et al., 2003, and divalent cations and protons (Verdoucq, Grondin and Maurel, 2008). Yeast AQP gating has also been documented using x-ray crystal structures determining the involvement of AQP N-termini (Fischer et al., 2009).
Potential gating for mammalian AQPs still remains an elusive topic and is not a widely-accepted regulatory mechanism (Wang and Tajkhorshid, 2007). However, studies into mammalian AQP gating by Reichow et al., 2013 andJanosi andCeccarelli, 2013 have identified novel gating mechanisms in mammalian AQP0 and human AQP5 respectively, potentially indicating a more substantial role for gating mechanisms in mammalian AQPs (Figure 1.13). There is also evidence that phosphorylation in response to many microenvironmental factors may also contribute to AQP gating (Nesverova and Törnroth-Horsefield, 2019) and recently mechanical stimuli has proposed as a potential gating mechanism (Hill and Shachar-Hill, 2015;Ozu et al., 2018) (Figure 1.13). Structures show the pore of each AQP monomer with regulatory amino acid residues highlighted. All AQPs share the ar/R selectivity filter at the extracellular side of the pore. At the cytosolic side of the pore, residues are displayed that contribute to gating; plant and yeast mechanisms are well established and recently discovered in sheep. Adapted from Törnroth- Horsefield et al., 2010. (B) Membrane stretch as a gating mechanism of AQP function. During osmotic stress, rapid water flux causes changes in cell volume and membrane tension (horizontal arrows). It is proposed that AQP structure may become distorted, halting water transport (vertical arrows), thereby controlling the osmotic response of cells. Adapted from Ozu et al., 2018. AQPs are also described as mercury-sensitive water channels due to the ability of mercury-containing compounds, such as HgCl2, to inhibit water flow. Mercury containing compounds regulate the function of AQPs by binding to amino acid residues and occlude the pore. The specific residue is C183 which contains two mercury binding sites (Savage and Stroud, 2007). Such compounds could potentially be useful drugs to regulate AQPs in disease if it was not for the highly toxic mercury content.

G-Protein Coupled Receptors and Protein Kinases
One of the most well-established mechanisms of AQP regulation is the effect of vasopressin on the G-protein coupled receptor (GPCR) Vasopressin receptor 2 (V2R), to increase AQP2 expression and trafficking in the collecting ducts of the kidney. Upon vasopressin binding, the seven-transmembrane domain GPCR initiates adenylate cyclase through active Gs-protein (Roos et al., 2012). This results in increased levels of intracellular cyclic adenosine monophosphate (cAMP) which in turn activates protein kinase A (PKA). PKA is a serine/threonine kinase that increases expression and trafficking of AQP2.
AQP2 expression is also increased as PKA phosphorylates and activates the nuclear import and binding of cAMP responsive element-binding protein (CREB) to the cAMP responsive binding element (CRE) transcription factor (Moeller and Fenton, 2012).
CRE resides upstream and is responsible for the up-regulation of many genes, including AQP2.
PKA also has a more direct action in the regulation of AQP2 as phosphorylation of specific amino acid residues, C256, causes AQP2 trafficking to the plasma membrane; first investigated using site-directed mutagenesis of putative phosphorylation residues by Kuwahara et al., 1995. 62 PKA is not the only serine/threonine kinase to play a role in AQP trafficking. The lipid-activated and C2 calcium binding domain containing kinase, protein kinase C (PKC), also activates AQP phosphorylation and trafficking (Conner et al., 2012).
Serine/threonine kinase phosphorylation of AQPs is a required action for movement; whether this corresponds to trafficking to the plasma membrane or internalisation into the cytoplasm. A number of different phosphorylation sites for many human AQPs have been determined (Figure 1.14).
Figure 1.14. The phosphorylation sites of AQPs. NPA motifs on B and E-loops are shown that orientate and specifically allow the passage of water molecules (grey and white spheres) through the channel. The positions of serine/threonine kinase phosphorylation sites for individual AQPs are shown as symbols above. Adapted from (Conner, Bill and Conner, 2013).

Calcium
Calcium plays a role in numerous cell signalling pathways, including AQP regulation. Along with AQP phosphorylation by protein kinases, Ca 2+ interactions with AQPs is essential for their trafficking (Conner et al., 2012). Ca 2+ ions interact via the calcium binding protein calmodulin (Yáñez, Gil-Longo and Campos-Toimil, 2012). The structure of calmodulin is comprised of four identical helix-loop-helix 'EF-hand' groups.
Each EF-hand structure contains the binding site for a single Ca 2+ ion (Grabarek, 2005).
AQPs have been identified to share sequence homology with calmodulin (Fotiadis et al., 2002) and calmodulin binding sites have been identified within the structure of AQP6 (Rabaud et al., 2009). Ca 2+ ions enter cells through numerous pathways including voltage-gated ion channels, G-protein coupled receptors (GPCR) and their associated second messenger pathways and the transient receptor potential (TRP) families of voltage-insensitive Ca 2+ channels. These specialist calcium channels are sensitive to a number of stimuli such as mechanical stress, temperature and a change in osmolality, with an emphasis on the latter regulating AQP trafficking (Benfenati and Ferroni, 2010).

Cytoskeletal proteins
When not integrated into the membrane AQPs are stored within intracellular vesicles, upon calcium activation and phosphorylation, these AQP-containing vesicles are trafficked to the cell surface along microtubules using the motor proteins dynein and kinesin (Tietz et al., 2006). There is also evidence that when AQP trafficking is activated the actin cytoskeleton is remodelled and relaxed to facilitate the movement of vesicles towards the membrane (Nicchia et al., 2008).  Mechanisms that regulate AQP translocation and function. Many cellular processes, such as calcium influx, phosphorylation and cytoskeletal rearrangement all contribute to the membrane expression, and therefore function, of AQPs. Transient potential receptor channels (TRPs); G-protein coupled receptor (GPCR); calmodulin (CaM); cyclic adenosine monophosphate (cAMP); inositol triphosphate (IP3). Adapted from Conner, Bill and Conner, 2013.

Aims and objectives
The IVD is a unique tissue that is well-adapted to its role in enabling the movement of the spine and withstanding compressive forces applied by the human body. Due to its location and function, the microenvironment bestowed upon the IVD is complex. Resident cells of the IVD have adapted to their microenvironment, which enables their overall function to maintain the role of the IVD within the spine. However, it is not completely understood how IVD cells have been able to survive and adapt to their environment or how such adaptation is affected during IVD degeneration, when the microenvironment is permanently altered causing adverse cellular functions and the destruction of tissue.
The overall aim of this PhD was to test the hypothesis that because the function of NP cells is governed by many microenvironmental factors that are in-part controlled by the function of AQPs (particularly osmolality), many AQP isoforms are expressed and regulated in NP cells, contributing to their adaptation to the unique environment of the IVD, and possible dysregulation during degeneration.

2.
Investigate the hyperosmotic regulation of AQP1 and 5 in human and rat NP cells and determine if TonEBP, a transcription factor essential for osmoadaptation and 68

Introduction
The IVD, composed of the central gelatinous NP, surrounded by the AF and CEP, is a highly specialised tissue facilitating the biomechanical function of the spine (Urban et al., 1977). Resident cells within the NP produce vast amounts of ECM that contain high concentrations of proteoglycans whose hydrophilic GAG side-chains promote water and cation retention (Ishihara et al., 1997). This allows the disc to resist compressive loading and permits motion of the spine due to a high level of hydration (Maroudas, Muir and Wingham, 1969;Maroudas, 1970;Urban et al., 1977;Urban, Holm and Maroudas, 1978;Kraemer, Kolditz and Gowin, 1985;Ishihara et al., 1997). Therefore, in comparison to other tissues, the healthy IVD maintains a hyperosmolar environment (Ishihara et al., 1997;Urban, 2002;van Dijk, Potier and Ito, 2011;Johnson, Shapiro and Risbud, 2014a). However, the hydration and osmolality of the disc is not constant (Urban and McMullin, 1985). With applied loading, between 18-25% of fluid is recycled during a diurnal cycle (McMillan, Garbutt and Adams, 1996;Urban, 2002). The increased physiological osmolality within the disc has been shown to increase matrix synthesis by NP cells compared to lower osmolalities (Neidlinger- Wilke et al., 2012;O'Connell, Newman and Carapezza, 2014). This highlights that NP cells can respond to osmotic changes within the disc which may be via control of water transport ).
During human IVD development the NP contains populations of large, vacuolated notochordal (NC) cell clusters that are replaced by mature NP cells during ageing (Risbud, Schaer and Shapiro, 2010). Interestingly, there are differences in IVD development between different species and different groups within the same species.
An example of this difference is highlighted in dogs; those classified as NCD retain NC cells into adulthood, maintain a healthier IVD environment and suffer infrequent degeneration. Whereas CD dogs lose NC cells before adolescence and suffer early-onset IVD degeneration Smolders et al., 2013). These differences in canine IVD biology enable research into how cell viability, function and differentiation are affected by the IVD environment and vice versa. To preserve their phenotype during culture, NC cells require a hyperosmolar environment (Spillekom et al., 2014) and they have been shown to provide a protective and regenerative effect on NP cells when cocultured together (de Vries et al., 2016;Mehrkens et al., 2017). This indicates that the behaviour of disc cells during development and maturation is, in some part, controlled by fluctuations in extracellular osmolality, and that responses may vary between NC cells and mature NP cells. IVD degeneration can also further disrupt the osmotic flux of the disc.
Around 40% of chronic LBP cases are attributed to IVD degeneration (Luoma et al., 2000), which is characterised by altered biomechanical properties of the spine caused by an imbalance between anabolic and catabolic mechanisms leading to the destruction of the ECM Haefeli et al., 2006;Le Maitre et al., 2015). The production of proinflammatory cytokines by NP cells are known to play a role in the degenerative cascade of IVD degeneration (Takahashi et al., 1996;Burke et al., 2002;Le Maitre, Freemont and Hoyland, 2005;Weiler et al., 2005;Le Maitre, Hoyland and Freemont, 2007a;Studer et al., 2011;Andrade et al., 2013;Purmessur et al., 2013;Risbud and Shapiro, 2014).
Specifically, over-expression of cytokines leads to increased production of matrix degrading enzymes (Roberts et al., 2000;Le Maitre, Freemont and Hoyland, 2004;Patel et al., 2007;Hoyland, Le Maitre and Freemont, 2008;Pockert et al., 2009;Zhao et al., 2011), and the resulting loss of hydrophilic proteoglycans causes a decrease in IVD osmolality compared to normal physiological conditions. This could further contribute to degeneration, as under low osmolality MMP-3 expression is increased and aggrecan expression is decreased in NP cells (Wuertz et al., 2007;Neidlinger-Wilke et al., 2012).
This further highlights the importance of regulating osmotic responses in the disc.
Aquaporins (AQPs) are a family of transmembrane channel proteins that function as selective water pores driven by osmotic gradients (Agre et al., 2002).
Aquaporin 1 (AQP1) was the first member to be discovered in erythrocytes and renal tubules (Denker et al., 1988) and since then thirteen members have been discovered (AQP0-12). These channels are localised to tissues with roles relating to water homeostasis, cell volume regulation and structural function (Day et al., 2014), and therefore may contribute towards the adaptation and responses of NC and NP cells when exposed to the fluctuating osmolality of the healthy and degenerate disc.
In human and murine discs, expression of AQP1, 2, 3 and 5 have been identified (Richardson, Knowles, Marples, et al., 2008;Gajghate et al., 2009;Wang and Zhu, 2011;Taş et al., 2012;Johnson et al., 2015;Palacio-Mancheno et al., 2018). Studies have also demonstrated that expression of AQPs in NP and NCs can be regulated by physiological conditions. Hyperosmolality upregulates AQP 2 in murine NP cells , and AQP3 in murine NC cells (Palacio-Mancheno et al., 2018). Whilst the expression of AQP1 and AQP5 is maintained at basal levels by hypoxia-inducible factor 1 alpha (HIF-1α) in NP cells (Johnson et al., 2015). Johnson et al.2015 also reported that AQP1 and AQP5 expression decreased in human IVDs with increasing degeneration. To date only AQP1, 2, 3 and 5 have been shown to be expressed by NP cells (Richardson, Knowles, Marples, et al., 2008;Wang and Zhu, 2011;Taş et al., 2012;Johnson et al., 71 2015) and AQP1, 2 and 3 by NC cells Palacio-Mancheno et al., 2018). However, no studies have investigated expression of the remaining AQP family members, and few have investigated expression in NC and NP cells and determined whether expression alters during maturation or degeneration. Hence, this study aimed to identify the expression of all thirteen AQP family members in human and canine NP tissues. Discs from NCD and CD canines were used as a model to investigate changes in AQP expression during NC to NP cell transition in IVD maturation, and adult human discs were used to investigate changes in AQP expression in NP cells during IVD degeneration. 72

Experimental Design
As only a few AQP family members have been identified in the human NP, within this study firstly qRT-PCR was utilised to identify the gene expression of all 13 AQP family members. IHC was then used to determine protein expression of AQPs that were expressed at gene level. Human NP samples used were graded histologically from nondegenerate to severely degenerate which enabled this study to identify changes in AQP expression in mature human tissue as IVD degeneration progresses. This study also aimed to determine the expression of AQPs during IVD maturation which was achieved via AQP protein expression using immunohistochemistry (IHC) in NCD and CD canine NP

Canine Tissue
Canine IVD samples (n=35) were obtained from 19 NCD and CD dogs (Thompson grade 1-5) euthanized for unrelated experiments with approval from the Ethics Committee of Animal Experiments of Utrecht University (Appendix III). The gross morphology of canine discs was categorised using a modified Thompson grading scheme (TG), specifically developed for canine IVDs, and each sample given a grade (1-5) to determine the state of disc degeneration . Canine IVD samples were sent to Sheffield Hallam University after tissue processing (Section 2.2.4).

Tissue Processing
Human tissue was fixed in 10% (v/v) neutral buffered formalin (Leica Microsystems, Milton Keynes, UK) for 48h and embedded into paraffin wax. Following embedding, 4µm sections were cut and human IVDs histologically graded by performing haematoxylin and eosin staining. Paraffin was removed from sections using sub-X (Leica Microsystems) 3 x 5min washes. Sections were then rehydrated in IMS 3 x 5min before 1min staining with Mayer's Haematoxylin (Leica Microsystems). Sections were left to 'blue' in running tap water for 5min before subsequent staining with alcoholic eosin 75 (Leica Microsystems) for 1min. Sections were then dehydrated in IMS 3 x 5min and cleared in sub-X (Leica Microsystems) 3 x 5min. Sections were finally mounted with pertex (Leica Microsystems). Microscopic tissue morphology was assessed to determine the grade of degeneration for each sample based on four distinct changes to IVD morphology during degeneration: demarcation of NP tissue, presence of fissures, cell clustering and reduction of eosin staining adjacent to NP cells. Each morphological feature was scored between 0-3 (higher score is more degenerate) and added together for each sample, to give a final grade of 0-12. Samples were separated into cohorts depending on grade: non-degenerate (grade 0-4); moderately degenerate (grade 4.1-6.9) and severely degenerate (grade 7-12).
Canine lumbar spinal segment (IVD and adjacent ½ vertebral bodies) extraction was performed by Bach et al., 2018. Canine vertebral columns (T11-S1) were harvested using an electric multipurpose saw (Bosch). Vertebrae were transversely transected using a band saw (EXAKT tape saw) and spinal segments were sagitally transected using a diamond band saw (EXAKT 312). Lumbar spinal segments were fixed in 4% (v/v) neutral buffered formaldehyde (Klinipath, Duiven, the Netherlands) for 14d and decalcified in PBS with 0.5M EDTA for 2 months. Spinal segments were then paraffin embedded and 5µm sections were cut . The grade of IVD degeneration was determined using the modified TG (grade 1-5) developed by Bergknut et al., 2013.

Investigation of AQP gene expression in directly extracted human NP cells
NP tissue from surgical and PM IVD tissue was separated from surrounding AF and CEP tissue and NP cells were isolated using 2mg/mL collagenase type I (Sigma) in DMEM for 4h at 37°C. Cells were then passed through a 40µm cell strainer (Sigma) to remove tissue debris. Following extraction, NP cells were either used for direct RNA extraction or cell culture. RNA was extracted by adding trizol (Life Technologies) for 10min at RT before chloroform (Sigma) was added, samples vortexed and allowed to stand for 10min at RT before centrifugation at 12,000g for 15min at 4°C. Samples were incubated at 42°C for 1 hour, to synthesise cDNA, followed by 10 minutes at 80°C to denature the MMLV reverse transcriptase in Bioscript RT enzyme (Bioline). Expression of target genes: AQP0-12 together with housekeeping genes: glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and 18S rRNA (18S), was investigated using quantitative real-time polymerase chain reaction (qRT-PCR) employing pre-designed Taqman™ gene expression assays (Life Technologies). A volume of 2μL cDNA sample, per well, was added in duplicate into 96-well FAST PCR plates (Life Technologies), qRT-PCR mastermix was prepared (Table 2.2), and 8μL was added per well.
Plates were sealed with MicroAmp™ optical adhesive film (Life Technologies).
Expression of target genes was determined using a QuantStudio 3 real time PCR machine (Life Technologies). Fast programme was used where the temperature was first ramped up to 95°C for 10min, to activate Taq polymerase, followed by 50 cycles of denaturing at 95°C for 1sec and subsequent annealing and extension at 60°C for 20sec. qRT-PCR was conducted on 19 non-degenerate samples (grade ≤4), 13 moderately degenerate samples (grade 4.1-6.9), 38 severely degenerate samples (grade 7-12) and 32 samples with evidence of cell infiltration identified histologically (Appendix II). Results were analysed using the 2 -ΔCt method (Livak and Schmittgen, 2001) and presented as relative gene expression normalised to the average CT of GAPDH and 18S. Both housekeeping genes used in this study have been previously shown to be stably expressed, using geNorm algorithm, in human IVD samples across all grades of degeneration (Phillips et al., 2015). (v/v) P/S (PAA Laboratories) to collect NC cells. After centrifugation at 300g for 5min, NC cell pellets were resuspended in 100% FCS (PAA Laboratories); 24.4 ± 0.9 x 10 6 (mean ± SD) live NC cells were recovered per dog (Spillekom et al., 2014). Cells were encapsulated in 2.4% (w/v) sterilised alginate solution (Sigma-Aldrich) with a final cell density of 3 x 10 6 cells/mL. The alginate/cell suspension was gently pressed through a 26-gauge needle into 102mM CaCl2 (Sigma-Aldrich) and 10mM HEPES (Sigma-Aldrich) to form solid, cell-encapsulated, beads. Alginate beads were cultured in 24-well plates (~6 beads/well). Beads were cultured in 1mL α-MEM (Invitrogen) supplemented with 10%

Isolation and culture of NCs from Canine discs
(v/v) FCS (PAA Laboratories) and 1% (v/v) P/S (PAA Laboratories) for 28d at 37°C, 5% (v/v) CO2. Media osmolarity was raised to 400mOsm/L by adding 1% (w/v) 5M NaCl (Merck) and 1% (w/v) 0.4M KCl (Sigma-Aldrich), before the start of culture. Media was replaced every 3d. These culture conditions were used as they allow the retention of the NC cell phenotype in culture (Spillekom et al., 2014). After culture duration beads were fixed in 10% (v/v) neutral buffered formalin (Leica Microsystems) and embedded into paraffin wax. NC cell extraction, encapsulation into alginate beads, cell culture, fixation and paraffin embedding were all performed by Spillekom et al., 2014. Four µm sections were cut and used in immunofluorescence experiments (section 2.2.8).
Antigen retrieval methods were then performed. During heat antigen retrieval, slides were immersed in heat antigen retrieval buffer (50mM Tris, pH9.5, pre-heated to 60°C), irradiated for 5min at 40% power in an 800W microwave oven (Sanyo), left to stand at RT for 1min, before being irradiated at 20% power for 5min. Slides were then cooled at RT for 15min. If enzyme antigen retrieval was required, slides were immersed in 300mL enzyme antigen retrieval buffer (Table 2.5) for 30min at 37°C.

TBS (all other components diluted in) 1x
CaCl2 pre-heated to 37°C -  Table 2.5. Antibody and blocking information used for IHC on human IVD tissue. All antibodies diluted in 1% (w/v) BSA in 1x TBS. All antibodies and serum purchased from Abcam. ab6720 (goat anti rabbit) and ab7074 (rabbit anti mouse) secondary antibodies at 1/500 dilution were used.

Clonality
Optimal dilution  Table 2.6. Antibody and blocking information used for IHC on canine IVD tissue. All antibodies diluted in 1% (w/v) BSA in 1x TBS. All antibodies and serum purchased from Abcam. ab6720 (goat anti rabbit) and ab7074 (rabbit anti mouse) secondary antibodies at 1/500 dilution were used.
After washing in 1x TBS 3 x 5min, slides were incubated with 1:500 dilution of biotinylated secondary antibody (Abcam), diluted in 1% (w/v) BSA in 1x TBS, for 30min at RT. Slides were washed with 1x TBS 3 x 5min and secondary antibody binding was detected by adding ABC elite reagent (Vector Laboratories, UK) for 30min at RT. After washing with 1x TBS 3 x 5min, slides were incubated with 0.08% (v/v) H2O2 and 0.65mg/mL 3,3'-diaminobenzidine tetrahydrochloride (DAB) (Sigma-Aldrich) in 1x TBS for 20min at RT. Slides were washed in H2O for 5min and counterstained with Mayer's haematoxylin (Leica Microsystems) for 1min before submersion into running tap water for 5min to 'blue' slides. Following, slides were dehydrated with IMS 3 x 5min, cleared with sub-X (Leica Microsystems) 3 x 5min, and finally mounted with pertex (Leica Microsystems).

Immunofluorescence
Following extraction, human NP cells were expanded in monolayer in DMEM glutamine (all Life Technologies), 50µg/mL amphotericin B and 50µg/mL ascorbic acid (Sigma) and maintained at 37°C in a humidified atmosphere containing 5% (v/v) CO2.
Human NP cells, at passage 2, were seeded into chamber slides (Life Technologies) at a density of 1x10 4 cells/well and allowed to adhere overnight. Media was aspirated and cells washed with PBS before fixing with 10% (v/v) neutral buffered formalin (Leica Microsystems) for 5min at RT. Sectioned canine NC cells in alginate were dewaxed in Sub-X 3 x 5min and rehydrated in IMS 3 x 5min. Fixed human NP cells and canine alginate sections were washed in PBS for 3 x 5min and human NP cells permeabilised in PBS 1% (v/v) Triton-X100 (Sigma-Aldrich) for 5min at RT. For both human NP cells and canine samples, non-specific binding sites were blocked for 1h at RT in 1% (w/v) BSA (Sigma-Aldrich) in PBS with either 25% (w/v) rabbit or goat serum (Abcam) ( n/a 1/1000 n/a n/a

Identification of aquaporin gene expression in human NP cells
The native gene expression of all mammalian AQP family members in directly extracted NP cells was investigated using qRT-PCR, of which AQPs 0-7 and 9 were identified within human IVD tissue (Figure 2.2). AQP8 was only expressed in 4 out of 97 samples, with no trends seen across grades of degeneration (data not shown) and AQPs 10, 11 and 12 were not identified in any sample. Whilst the levels of AQP 0-7 and 9 expression did not alter significantly between grades of degeneration, significant

Immunodetection of aquaporin proteins in human IVD tissue
The research group has previously identified AQP1 and 5 in human IVD tissues at protein levels (Johnson et al., 2015), therefore AQPs 0, 2, 3, 4, 6, 7 and 9 were investigated here for their presence in human NP tissue in different histological grades of degeneration. Protein expression of all 7 AQP isoforms investigated was identified in the NP cells of human IVD tissues. However, the percentage immunopositivity for AQP0 and 3 was not significantly altered between grades of degeneration (Figure 2.3).
The percentage immunopositivity for AQP4 in human NP tissue significantly decreased in moderately and severely-degenerate discs compared to non-degenerate discs (p ≤ 0.05) (Figure 2.4). Conversely, immunopositivity of AQP7 in severelydegenerate human NP tissue was significantly increased when compared to moderately and non-degenerate tissue (p ≤ 0.05) (Figure 2.4). The immunopositivity of AQP6 and 9 was not significantly altered between grades of degeneration (  Immunopositivity of AQP0, 2 and 3 within human NP tissue. Three cohorts were investigated for the expression of AQP family members in human IVD; non-degenerate (grade 0-4, n=10), moderately degenerate (grade 4.1-6.9, n=10), severely degenerate (grade 7-12, n=10). Immunopositive cells were expressed as a percentage of total count; the median value is represented by the bars. Statistical significances in % immunopositivity determined using Kruskal-Wallis test * = p ≤ 0.05. Scale bar 50µm. Immunopositivity of AQP4, 6, 7 and 9 within human NP tissue. Three cohorts were investigated for the expression of AQP family members in human IVD; nondegenerate (grade 0-4, n=10), moderately degenerate (grade 4.1-6.9, n=10), severely degenerate (grade 7-12, n=10). Immunopositive cells were expressed as a percentage of total count; the median value is represented by the bars. Statistical significances in % immunopositivity determined using Kruskal-Wallis test * = p ≤ 0.05. Scale bar 50µm.     Three cohorts were investigated for the expression of AQP family members in NC cells; Thompson grade (TG) 1-3, and NP cells; TG 1-5. Immunopositive cells were expressed as a percentage of total count; the median value is represented by the bars. Statistical significances in % immunopositivity determined using Kruskal-Wallis test * = p ≤ 0.05.
NP cells reside within an osmotically challenging environment (Kraemer, Kolditz and Gowin, 1985;Urban, 2002;van Dijk, Potier and Ito, 2011;Johnson, Shapiro and Risbud, 2014a), where cells must be able to adapt to their environment to allow correct function. Previously AQPs 1, 2, 3 and 5 have been shown to be expressed by NP cells Gajghate et al., 2009;Wang and Zhu, 2011;Taş et al., 2012;Johnson et al., 2015), expression of AQP1, 2, and 5 are regulated by physiological conditions found in the disc such as hyperosmolality ) and AQP1 and 5 expression levels decreased with IVD degeneration (Johnson et al., 2015). This highlights that water transport via AQPs in the disc must be tightly controlled to allow the survival and function of NP cells, and such regulation is diminished in degeneration, possibly aggravating the catabolic cascade. However, to date, no studies have investigated the expression of all AQP family members in the IVD or determined how their expression is altered with degeneration, thus it is difficult to gain a complete understanding of the roles for AQPs in the IVD. For the first time the expression of ten AQP isoforms, AQPs 0-9, in human and canine NP tissue were shown. We determined that the expression of certain AQPs is altered during IVD degeneration of humans and canines. Utilising canine discs, we were also able to investigate expression and localisation during cellular maturation and demonstrated nine AQPs were expressed in canine NC cells, which were particularly associated with vacuolar membranes, suggesting a role in water transport into vacuoles.

AQP expression in IVD tissues
Our results agree with studies that have shown AQP1, 2, 3 and 5 expression in NP cells (Richardson, Knowles, Marples, et al., 2008;Gajghate et al., 2009;Wang and Zhu, 2011;Taş et al., 2012;Johnson et al., 2015). However, whilst  identified that AQP1 and AQP3 were expressed within human IVD in agreement with our study, they failed to demonstrate expression of AQP2. However, in the current study both gene and protein expression of AQP2 was identified in human discs and protein in canine discs. In the earlier study (Richardson, Knowles, Marples, et al., 2008)  Limited AQP family members have been identified in other musculoskeletal tissues, including AQP1, 3, 4 and 9 in articular cartilage Trujillo et al., 2004;Cai et al., 2017;Steinberg et al., 2017;Takeuchi et al., 2018), AQP1 and AQP9 in synovial tissues Nagahara et al., 2010), AQP1 and 3 in osteoblasts (Mobasheri, Wray and Marples, 2005;Barron et al., 2017) and AQP9 expression in osteoclasts (Aharon and Bar-Shavit, 2006;Liu et al., 2009). Here, for the first-time gene expression of AQPs0, 4, 6-9 was shown in human NP tissue, although AQP8 was only expressed in a small number of samples at gene level (n=4). Furthermore we demonstrate for the first time protein expression of AQPs 0, 4, 6, 7 and 9. Together this data demonstrates NP cells express many AQPs which may play important roles in control of cell volume in response to osmotic flux, there is also evidence to suggest AQP2 is also involved (Galizia et al., 2012). Therefore, AQP2 and 5 expression may indicate that vacuoles regulate NC cell responses to their hyperosmolar environment, and their decreased expression during NC to NP transition could regulate the decreased cell size seen during maturation. AQP6 expression potentially indicates that vacuoles play a role in regulating acid-base balance in NC cells. In the kidney, where AQP6 is expressed intracellularly, low pH activates the anion permeability of AQP6 (Yasui et al., 1999) and increases AQP6 expression (Promeneur et al., 2000). An increase in AQP6 expression in response to low pH also potentially explains the increase in AQP6 observed in canine NP cells during degeneration, where extracellular pH decreases.

AQP expression during degeneration of the IVD
Expression of AQP1 and 5 in the disc has been shown to be sensitive to degeneration within human IVD tissue previously (Johnson et al., 2015), with levels decreasing with degeneration. Here, AQP4 protein expression was also shown to decrease during human IVD degeneration. This potentially indicates shared functions of these AQPs which, during degeneration when expression is decreased, results in a diminished ability of NP cells to control water transport and cell volume regulation as the surrounding environment becomes increasingly osmotically challenging. This decrease in AQP4 expression is of particular importance as within the AQP family, AQP4 has the fastest rate of water permeability followed by AQP1 and 5 (Yang and Verkman, 1997;, and these AQPs have all been implicated in the control of regulatory volume decrease when cells are exposed to hypo-osmotic stimuli (Benfenati et al., 2011a;Mola et al., 2016). Thus the decreased expression of AQPs 1, 4 and 5 seen during human IVD degeneration could be a result of hypo-osmotic stimuli observed during IVD degeneration (Ishihara et al., 1997).
An important response to osmotic stress is translocation of AQPs to the plasma membrane, in primary rat cortical astrocytes, HEK293 and MDCK cells AQP4 is rapidly translocated to the plasma membrane in response to osmotic changes (Mazzaferri, Costantino and Lefrancois, 2013;. This translocation is governed by many mechanisms including vesicle transport along microtubules (Mazzaferri, Costantino and Lefrancois, 2013;. In human NP cells we observed that AQP4 co-localised with β-tubulin, a major component of microtubules, indicating AQP4 is possibly translocated between the cytosol and plasma membrane in order to respond and adapt to changes in the extracellular osmolality. In contrast to the decrease seen in AQP1, 4 and 5 with degeneration, the number of NP cells expressing AQP2 and AQP7 increased in moderate or severely degenerate human IVD tissues respectively. This may indicate that AQP2 and 7 are regulated by different underlying mechanisms and have different functions compared to AQP1, 4 and 5. The increase in the number of cells expressing these proteins may be a consequence of degeneration or a potential repair mechanism employed by NP cells to re-establish homeostasis dysregulated during degeneration. As the disc degenerates, the microenvironment in which cells reside, irreversibly changes, such as a decrease in disc pH (Urban, 2002). This potentially explains the increase in AQP2 protein expression, as AQP2 expression is increased in the rat kidney when exposed to lowered pH (Amlal, Sheriff and Soleimani, 2004). Immunopositivity of AQP3, 6 and 9 in human IVD tissue was not altered during degeneration, yet the majority of NP cells in native IVD tissue expressed all three AQPs, potentially indicating roles which may not be sensitive to IVD degeneration. Within canine discs some effects mirrored the effects seen in human discs; however, some AQPs showed differential responses between species that remain to be explored. AQP5 immunopositivity decreased (Johnson et al., 2015) and AQP7 immunopositivity increased in human and canine NP cells with IVD degeneration.

Conclusion
As the disc is an osmotically challenged tissue which is further altered during degeneration, NP cells must employ several tightly regulated mechanisms to survive and function within their unique environment. We postulate that the expression of numerous AQP family members in the disc highlights the importance of maintaining intricate control over regulation of water transport and cell volume in NP cells ( Figure   2.10). The increased proportion of NP cells expressing AQP2 and AQP7 during degeneration may reflect consequences or potential repair mechanisms employed by the disc as NP cells lose their ability to adapt to the degenerate hypo-osmolar environment, which is identified by the loss of AQP1, AQP4 and AQP5 expression during degeneration. The expression of numerous AQPs within the disc may highlight the importance of controlling water and solute transport in relation to maintaining NC and NP cell function. As yet the actual roles of AQPs within disc maturation and degeneration are unknown; further investigation is warranted to identify mechanisms of regulation and functions of AQPs to elucidate the true action of these transmembrane channel proteins on the overall behaviour and health of the disc. 110

Introduction
Many factors contribute to the correct function of the NP. The environment in which NP cells reside plays a big role in the physiology of the tissue; the correct biomechanical loading, pH, nutrition, O2 tension and adaptation to a diurnal cycle all allow NP tissues to function correctly (McMillan, Garbutt and Adams, 1996;Ishihara et al., 1997;Urban, 2002;Bibby et al., 2005;Hartman et al., 2018). As the NP has a high content of negatively charged, hydrophilic GAGs within its matrix, allowing water and cation retention, this provides a high osmotic pressure environment within the disc (Ishihara et al., 1997;van Dijk, Potier and Ito, 2011;Johnson, Shapiro and Risbud, 2014a (Burg, Ferraris and Dmitrieva, 2007). So how have NP cells been able to adapt to this constantly changing hyperosmotic environment?

TonEBP
There are many cell signalling pathways that are activated during osmotic stress including p38 MAPK (Han et al., 1994), Jnk (Galcheva-Gargova et al., 1994) and ERK1/2 (Burg, Kwon and Kültz, 1996), but one of the most well-known, classical pathways, which many tissues and cell types employ to counter osmotic stress involves the function of tonicity enhancer binding protein (TonEBP). TonEBP, is also known as nuclear factor of activated T cells 5 (NFAT5), is part of a family of transcription factors (NFAT1-5) that contains rel homology domains (DNA binding domains), but unlike NFAT1-4, TonEBP function is independent of calcineurin (a calcium-dependent serine/threonine phosphatase that activates T cells) and activator protein 1 (AP-1 ,a transcription factor controlling cellular processes such as growth and differentiation) (Macián, López-Rodríguez and Rao, 2001). Within many tissues TonEBP has the role of hyperosmotic adaptation; the exact mechanisms by which cells sense osmotic shifts is unknown, but studies suggest that integrins α6β4 (Jauliac et al., 2002) and α1β1 (Moeckel et al., 2006), a guanine nucleotide exchange factor, Brx (also known as A-kinase anchoring protein 13 (AKAP13)) (Kino et al., 2009), and biomechanical stretching (Scherer et al., 2014) may all activate TonEBP signalling. When cells are exposed to hyperosmotic stimuli, TonEBP is translocated into the nucleus. In the nucleus it forms homodimers and binds to tonicity response elements (TonE) on target genes, via rel homology domains, causing their upregulation (Miyakawa et al., 1999). These well-described target genes include Sodium myo-inositol transporter (SMIT), aldose reductase (AR), Betaine-γ-amino butyric acid transporter (BGT1) and taurine transporter (TauT), that all facilitate the exchange of charged ions within cells for small non-ionic osmolytes (Miyakawa et al., 1999;   TonEBP has an emerging role in the function of the IVD and similar tissues. It has been shown that TonEBP is a key regulator of aggrecan (Tsai et al., 2006)

Osmotic regulation of AQP expression
Many AQPs are present within NP tissue (Chapter 2). Their presence suggests that water transport is tightly controlled and very important for NP cell function. AQP1 and 5 expression in human NP tissue has previously been shown to decrease during IVD degeneration (Johnson et al., 2015). The basal expression levels of AQP1 and 5 in NP cells is regulated by HIF-1α (Johnson et al., 2015) which does not explain their decreased expression, therefore other degenerative changes may be implicated in the dysregulation of AQP1 and 5. AQP expression across many tissues and cell types has been shown to be regulated by alterations in extracellular osmolality, with many pathways implicated (Table 3.1), however only one study to date has investigated this within the IVD, which showed increased AQP2 expression .

Lung
Rat alveolar epithelial cells

AQP5
Hyperosmotic ↑ HIF-1α (Zhou et al., 2007) 115  Specifically, AQP1 expression is upregulated by hyperosmolality in mIMCD-3 kidney cells (Jenq et al., 1998;Umenishi and Schrier, 2003; and mouse cardiac endothelial cells (Rutkovskiy, Mariero, et al., 2012a). AQP5 expression has also been shown to be upregulated by hyperosmolality in 3AO ovarian cancer cells (Chen et al., 2015), rat alveolar epithelial cells (Zhou et al., 2007) and human retinal epithelial cells (Hollborn et al., 2015), in which AQP5 expression was reduced after hypo-osmotic treatment (Hollborn et al., 2015). TonEBP has also been implicated with the hyperosmotic upregulation of both AQP1 (Umenishi and Schrier, 2003; and AQP5 (Hollborn et al., 2015). This potentially indicates that both AQPs are also regulated in the same manner in NP cells, as the osmolality of NP tissue decreases during degeneration. There is also a precedence of AQP regulation in the IVD; AQP3 expression was upregulated by hyperosmolality in 3D cultured murine NC cells (Palacio-Mancheno et al., 2018) and the hyperosmotic upregulation of AQP2 in 2D cultured murine NP cells was dependent on TonEBP ). If AQP1 and 5 are regulated in this manner, it may implicate them in enabling the adaptation of NP cells to their hyperosmotic environment along with other TonEBP target genes. This adaptation may be diminished during degeneration when their expression is reduced, possibly by the lowered extracellular osmolality.
AQP expression and function has also been implicated in enabling cells to sense osmotic stress via membrane tension changes (Ozu et al., 2018), interactions with other osmotically activated membrane channels (Benfenati et al., 2011a;Mola et al., 2016) and the control of cell volume regulation , indicating that AQPs may also function upstream of TonEBP to enable cellular adaptation to osmolality changes.
As TonEBP also regulates the expression of NP matrix genes, if AQP1 and 5 expression in NP cells is linked to TonEBP, they may also be linked to matrix production and therefore the fundamental function of NP cells to maintain the integrity of the IVD.
This study aimed to investigate if AQP1 and 5 expression was regulated by hyperosmolality in 2D and 3D cultured human NP cells to determine why their expression is decreased during degeneration. Also, to identify if AQP1 and 5 are similarly regulated in rat NP cells, which may determine if this response is common across different species, highlighting that it is a fundamental process for IVD function, and how TonEBP is involved in the osmotic regulation of both AQPs. To assess the in vivo role of TonEBP in the expression of AQP1 and 5, expression in wildtype (WT) and TonEBP -/mice embryos were also investigated.

Experimental design
As the reasons for the decrease in AQP1 and 5 expression during in NP tissue during IVD degeneration are unknown, 2D cultured human NP cells were treated with hyperosmotic media (representative of physiological to healthy IVD). qRT-PCR and western blotting were utilised to determine if AQP1 and 5 gene and protein expression were sensitive to alterations in extracellular osmolality, which has been shown in a number of other tissue types. Furthermore, gene expression was also investigated in 3D cultured human NP cells exposed to hyperosmotic treatment to determine if AQP1 and  119

Human Tissue
Human IVD tissue was obtained from patients undergoing microdiscectomy surgery for the treatment of nerve root compression as a result of IVD herniation or post-mortem (PM) examination with informed consent of the patients or relatives.

Tissue Processing
Human tissue was fixed in 10% (v/v) neutral buffered formalin (Leica Microsystems) and embedded into paraffin wax. Following embedding, 4µm sections were cut and human IVDs histologically graded using methods described previously (section 2.2.4) to determine severity of IVD degeneration.

Human NP cell extraction and culture
Human NP cells were isolated from surgical and PM tissue and expanded in 2D culture as described previously (Section 2.2.5). Human NP cells at passage 2 were utilised for all experiments.

Hyperosmotic protein regulation of AQPs in human NP cells
Human NP cells were seeded into T25 culture flasks (Nunc, Paisley, UK) and treated with 425mOsm/kg media for 8, 24 or 48h. Cells in standard culture media (325mOsm/kg) at 0h were used as controls. Following treatment, cells were washed twice with ice-cold PBS before 500μL Radioimmunoprecipitation assay (RIPA) buffer was added to lyse cells. RIPA buffer was used to lyse cells as it contains both ionic and non-ionic surfactants that enable the effective lysis of whole cell extracts and membrane-bound proteins. Components of RIPA buffer are outlined in Table 3.2.

Rat NP cell extraction and culture
Wistar rats (200-250g) were euthanised with CO2 and spinal columns dissected under aseptic conditions. Lumbar IVDs were separated from the spinal column and the NP separated from the AF using a dissecting microscope. NP tissue was partially digested with 0.1% (w/v) collagenase (Sigma-Aldrich) and 10U/mL hyaluronidase (Sigma-Aldrich) for 4-6h and then maintained in DMEM supplemented with 10% (v/v) FBS and P/S. Rat NP cells migrated out of explant tissue after 1w; when confluent, cells were passaged using trypsin (0.25%) EDTA (1mM) and subsequently cultured.

Hyperosmotic gene regulation of AQPs in rat NP cells
Rat NP cells were seeded at 3x10 4 cells/well in 6-well plates and treated with altered osmolality media (425mOsm/kg or 525mOsm/kg) for 8 and 24h. NP cells cultured in standard media (325mOsm/kg) at time point 0h were used as controls.
After treatment cells were washed in ice-cold PBS before cell lysis and RNA extraction as described in section 2.2.5. RNA was finally eluted in 20μL RNase-free water.
Extracted RNA was then converted to cDNA by adding 20μL RNA to EcoDry™ premix (Takara) and incubated at 42°C for 1h, followed by 10min at 70°C,. cDNA was stored 127 at -20°C. To determine regulation of AQP1 and 5 gene expression under hyperosmotic treatment qRT-PCR was performed using Taqman methods outlined in section 2.2.5. Pre-designed primer/ probe mixes (Life Technologies) were used to determine gene expression of AQP1, 5 and GAPDH, which was used as a housekeeping control.

Hyperosmotic protein regulation of AQPs in rat NP cells
Rat NP cells were seeded at 4x10 4 cells/well in 6-well plates and treated with altered osmolality media (425mOsm/kg or 525mOsm/kg) for 8 and 24h. NP cells cultured in standard media (325mOsm/kg) at time point 0h were used as controls.
After treatment cells were washed in ice-cold PBS before adding 200μL homemade lysis buffer, (made up in mammalian protein extraction reagent MPER, Sigma-Aldrich) ( Determination of cell lysate protein concentration, separation of proteins using SDS-PAGE and transfer onto PVDF membranes is described in section 3.2.6. The only difference in the SDS-PAGE procedure was the use 5μL of pPrecision plus protein™ dual colour standards ladder (Bio-rad) to determine the molecular mass of proteins. After transfer PVDF membranes were cut horizontally at ~45kDa to separate higher molecular mass proteins (β-tubulin, 55kDa, used as housekeeping control) and lower molecular mass proteins (AQP1 and 5, 28kDa, proteins of interest). Membranes were blocked in TBS-T containing 5% (w/v) NFDM for 1h at RT, before overnight incubation with either rabbit polyclonal antibody against AQP1 (1:500, AB3272, Millipore), rabbit polyclonal antibody against AQP5 (1:500, A4979, Sigma-Aldrich) or mouse monoclonal antibody against β-tubulin (1:1000, MA5-16308, Invitrogen). Membranes were washed in TBS-T 3 x 5min before incubation with goat anti-rabbit IgG (H+L) secondary antibody, HRP conjugated (1:1000, 31460, Invitrogen, for AQP1 and 5 probed membranes) or goat anti-mouse IgG (H +L) 129 secondary antibody, HRP conjugated (1:2000, 31430, Invitrogen, for β-tubulin probed membranes) for 1h at RT. All antibodies were diluted in TBS-T containing 1% (w/v) NFDM. Immunolabelling of protein bands was detected by adding ECL reagent (Amersham) to membranes for 2min before removing excess liquid and visualised under chemiluminescence using ImageQuant™ LAS 4000 (GE Healthcare).

TonEBP knockdown in rat NP cells
To investigate the potential role of TonEBP in the hyperosmotic regulation of AQP1 and 5 in rat NP cells, the expression of this transcription factor was knocked down. HEK293T cells were seeded in 10cm 2 culture plates at 5x10 6 cells/plate in

Slides were washed (3 x 5min), mounted with ProLong™ diamond antifade mountant
with DAPI (Life Technologies) and allowed to cure at RT overnight, protected from light. Slides were stored at 4°C. Mounted slides were visualised using an Axio Imager A2 (Zeiss). All images were taken at identical exposures to enable the quantification of fluorescent intensities. 132

Statistical analysis
The regulation of AQP1 and 5 gene and protein expression, in both human and rat NP cells, was performed in triplicate on 3 patients or pooled rat cells on different days. Data was found to be non-parametric, therefore Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences between AQP1 and 5 gene expression (investigated by qRT-PCR), and protein expression (investigated by western blotting). IHC was performed on 3 WT and TonEBP-/-rats each, in at least duplicate. Fluorescent images were taken, all at the identical exposure, gain and magnification. Fluorescent intensity of NP regions was determined using ImageJ software and normalised to the number of NP cells within the same IVD. Data was found to be non-parametric, therefore Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences between fluorescent intensities of WT and TonEBP-/-mice, after AQP1 and 5 IHC experiments, to determine putative changes in expression.  Regulation of AQP1 and AQP5 gene expression, after 48h treatment with 425 or 525mOsm/kg media, in human NP cells cultured for 2w encapsulated in alginate beads prior to treatment. The osmolality of untreated controls in standard culture media was 325mOsm/kg. The osmolality of treatment media was altered using NaCl and was measured using a freezing point depression osmometer (Model 3320 osmometer, Advanced Instruments). Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

Figure 3.4. Hyperosmotic regulation of AQP1 protein expression in human NP cells. (A)
Western blot analysis of AQP1 expression in human NP cells when exposed to 8, 24 and 48h of 425mOsm/kg media treatment, when compared to 325mOsm/kg (untreated) media controls at 0h. β-actin was used as a loading control. Chameleon® duo pre-stained protein ladder (LI-COR) was used to determine the size (kDa) of protein bands on blots. (B) Densitometry measurements of AQP1 protein expression, in human NP cells, when exposed to 8, 24, and 48h 425mOsm/kg media treatment compared to 325mOsm/kg control at 0h. The relative AQP1 expression is normalised to β-actin expression. Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

TonEBP expression in native human NP tissue
TonEBP is expressed in non-degenerate (  Immunopositivity results determining the percentage of cells within NP tissue expressing TonEBP. Two-hundred cells per patient sample were counted as positive or negative. Each point represents an individual patient NP sample. The grades of degeneration are described as followed: non-degenerate (0-4), moderatelydegenerate (4.1-6.9) and severely-degenerate (7-12). Positive staining for TonEBP is indicated by brown staining and cell nuclei are counterstained blue with Mayer's haematoxylin.

AQP1 gene expression was significantly upregulated in rat NP cells after 8h
treatment with 425mOsm/kg media (p ≤ 0.05) (Figure 3.6A) and after 8h and 24h treatment with 525mOsm/kg media (p ≤ 0.05) (Figure 3.6A). After 8h of 525mOsm/kg media treatment AQP1 gene expression was significantly higher compared to when rat NP cells were exposed to the same treatment for 24h (p ≤ 0.05) (Figure 3.6A).    The effects of TonEBP knockdown on the hyperosmotic regulation of AQP1 and 5 in rat NP cells. Rat NP cells were exposed to control (325mOsm/kg) or hyperosmotic (525mOsm/kg) treatment for 24hr, after 4d of TonEBP knockdown (shTonEBP) or no knockdown control (shCTR). After knockdown and treatment, the gene and protein expression of TonEBP (A, B), AQP1 (C, D) and AQP5 (E, F) was determined respectively. Results were normalised to 325mOsm/kg shCTR controls. Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

Effect of TonEBP expression on in vivo expression of AQP1 and 5
AQP1 was expressed in the spine and tail discs of wild type TonEBP +/+ ( Figure   3.8A and B), and complete knockout TonEBP -/- (Figure 3.8C and D) embryonic mice.
AQP5 was expressed in the spine and tail discs of wild type TonEBP +/+ ( The relative fluorescence of AQP1 immunostaining in TonEBP +/+ spine discs was significantly higher when compared with TonEBP +/+ tail discs, spine and tail discs from TonEBP -/-mice (p ≤ 0.05) (Figure 9.8E) (secondary antibody controls, Appendix VII).  (E) Fluorescence intensity measurements of AQP1 immunostaining images (20x) normalised to cell number. All images were captured at the same exposure on an Axio Imager A2 using Zen software (both Zeiss). Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

Discussion
NP cells have adapted to the environment in which they reside. The physiological O2 tension (Fujita et al., 2012), pH , nutrient diffusion (Urban, 2002), mechanical loading (Neidlinger- Wilke et al., 2012) (Johnson et al., 2015), yet the cause of this, and how both AQPs are regulated, is unknown. Therefore, for the first time this study has identified that AQP1 and 5 gene and protein expression in NP cells is upregulated by hyperosmolality, which potentially explains why their expression decreases during IVD degeneration, as the osmolality of the NP decreases. This study also determined that the in vitro and in vivo expression of AQP1 and 5 expression in NP cells is reliant on TonEBP. As TonEBP is an important transcription factor involved in the overall function of the IVD, the regulation of AQPs by TonEBP may also implicate that their function is also important for the health of the IVD. 146

AQP1 and 5 in human and rat NP cells were upregulated in hyperosmotic
conditions that mimic the physiological conditions of the healthy IVD. AQP1 and 5 are also regulated in a similar manner, under similar hyperosmotic conditions, in a range of other tissues and cell types (Jenq et al., 1998;Umenishi and Schrier, 2003;Zhou et al., 2007;Rutkovskiy et al., 2012;Chen et al., 2015;Hollborn et al., 2015), possibly indicating shared mechanisms of regulation. However, hyperosmotic treatment has also led to the reduction of AQP1 and 5 expression (Muller, Sendler and Hildebrandt, 2006) and hypo-osmotic treatment has led to increased expression of AQP1 (Moon et al., 2006) in other tissues, highlighting that the osmotic regulation of AQP expression is also tissue and cell type-specific. 3D hyperosmotic treatment of human NP cells also showed upregulation of AQP1 and 5, along with 2D treatment, indicating that this response may be relevant in vivo, as NP cells are known to re-differentiate into an in vivo-like state when cultured in 3D.

AQP1 and 5 gene expression in human NP cells was upregulated by
hyperosmotic conditions with an osmolality of 425mOsm/kg, yet the highest upregulation of AQP1 and 5 gene expression in rat NP cells was observed with 525mOsm/kg treatment. This potentially indicates that there are differences in regulation across species, possibly due to the differences in NP environments between adult human and young rat discs. NC cells are preserved within rat IVDs compared to adult human IVDs where NC cells are replaced by NP cells; NC cells produce more hydrophilic matrix when compared to NP cells and therefore may raise the osmolality of the young rat NP to higher levels when compared to a mature human IVD. Consequently, rat NP cells may respond to a higher osmolality treatment (with regards to AQP1 and 5 upregulation), compared to human NP cells, as this mimics the in vivo environment more closely, the same going for the response in human NP cells, where 425mOsm/kg may represent NP tissue osmolality more closely. The differences in gene expression may also be a result of the treatment timeframe. AQP1 expression in rat NP cells is significantly upregulated after 8h treatment, yet after 24h there is no significant difference compared to baseline levels, and AQP5 is not upregulated at either time point. This may simply be due to AQP regulation by 425mOs/kg occurring before 8h or after 24h.
However, the protein expression of AQP1 and 5 in both human and rat NP cells was significantly upregulated by 425mOsm/kg treatment, highlighting that protein expression is regulated similarly across species. Yet the results for AQP1 and 5 protein expression in rat NP cells may not be entirely trustworthy as membranes were cut horizontally to separate differently sized protein bands; so AQP1/5 and βtubulin could be probed at the same time without needing to strip PVDF membranes.
This is not best practice as there is the potential to miss out staining of differently sized bands which may relate to protein dimers/trimers and post-translationally modified proteins, and therefore results may not reflect changes in the total protein concentration, only selected species. In human NP cell western blotting the whole intact membrane was scanned which may have provided more accurate results. Only 1 band, at ~28kDa, for AQP1 was observed in human experiments; this increases confidence in the observed band, also at ~28kDa, in rat experiments. Also, housekeeping proteins were used to normalise against for western blotting analyses.
This poses questions as to whether these proteins are really stably expressed under treatment and whether there is biological variation between different samples, however, published research articles have used the same housekeeping proteins, suggesting they are suitable controls under similar treatments in identical cell types Johnson et al., 2017;Choi et al., 2018).
As AQP1 and 5 are regulated by osmolality in a similar manner in human and rat NP cells, this indicates there are shared mechanisms of regulation and that AQP regulation is important for IVD function across species. AQP1 and 5 may enable NP cells to adapt to their hyperosmolar environment, and when expression decreases in degeneration (Johnson et al., 2015) this may be due to the decrease in the extracellular osmolality, causing further dysfunction of NP cells. This may mean that AQP1 and 5 expression is decreased as a consequence of initial degeneration, but also leads to a continuation of the degenerative cascade where cells can no longer adapt to their environment.

The role of TonEBP on the expression of AQP1 and 5
TonEBP is important in enabling cells to adapt to a hyperosmotic environment, which is essential for NP cells, which reside within a hyperosmotic environment during healthy IVD physiology. Hyperosmolality increases TonEBP mRNA and protein expression and causes nuclear import and upregulation of target genes (TauT, SMIT, BGT1, AR) (Burg, Ferraris and Dmitrieva, 2007) to enable cellular adaptation to the IVD environment. TonEBP also controls aggrecan and GAG synthesis under hyperosmotic conditions in NP cells (Tsai et al., 2006;, therefore TonEBP function has important implications for IVD function and the maintenance of NP osmolality. This study identified that AQP1 and 5 gene expression, along with their hyperosmotic upregulation, in rat NP cells was significantly reduced when TonEBP expression was knocked down. These results highlight that AQP1 and 5 expression in NP cells is potentially governed by the expression and function of TonEBP. TonEBP has also been shown to regulate the expression of AQP1 and 5 in other tissues (Umenishi and Schrier, 2003;Hollborn et al., 2015), providing a precedent for results observed in this study. TonEBP expression is also required for the hyperosmotic upregulation of AQP2 in NP cells , therefore AQP1, 2 and 5 may enable the adaptation of NP cells to their hyperosmolar environment along with the classical TonEBP-targeted osmotic response genes.
Expression and hyperosmotic upregulation of AQP5 protein is significantly down-regulated by TonEBP knockdown suggesting that TonEBP is essential for the correct expression of AQP5 in NP cells. However, basal levels of AQP1 protein expression were slightly reduced (not significantly) by TonEBP knock down, yet the hyperosmotic up-regulation of AQP1 remained unaffected under the same conditions. Even though AQP1 gene expression was clearly regulated by TonEBP, other mechanisms may control protein expression. A possible explanation is that under hyperosmotic conditions, AQP1 protein expression is somehow stabilised for longer periods. Evidence for such an explanation is provided by Leitch, Agre and King, 2001, who identified that in fibroblasts isolated from BALB/c mice AQP1 expression was upregulated by hyperosmotic treatment, which reduced ubiquitination and proteasomal degradation of AQP1, ultimately leading to increased protein stability under these conditions (Leitch, Agre and King, 2001). This is potentially how AQP1 protein expression is still upregulated in response to hyperosmolality, when TonEBP is knocked down in NP cells, even though gene expression is significantly reduced.
Along with the classical function in regulating the cell's response to hyperosmotic stress, TonEBP has also been shown to increase the expression of TNFα (Esensten et al., 2005), IL-6 (Ueno et al., 2013) and MCP-1 (Küper, Beck and Neuhofer, 2012;Ueno et al., 2013) under hyperosmotic conditions, all of which are upregulated in IVD degeneration (Le Maitre, Hoyland and Freemont, 2007a;Andrade et al., 2013;Phillips et al., 2015). Parallel to the role under hyperosmotic stress, TonEBP is also activated by TNF-α signalling in NP cells, which leads to the upregulation of genes involved in IVD degeneration, rather than the classical osmotic response genes (Johnson et al., 2017). This highlights that TonEBP may also play a role during IVD degeneration; as the osmolality decreases TonEBP no longer upregulates osmotic response genes, possibly including AQP1, 2 and 5, but rather is activated by TNF-α signalling, intensifying the degenerative cascade (Johnson et al., 2017). This could also explain why altered TonEBP signalling has been seen during degeneration, yet this study demonstrated no change in numbers of cells expressing TonEBP.
It is currently unknown if AQP1 and 5 could also function upstream of TonEBP, this may be possible if AQP1 and 5 at the membrane are able sense changes in membrane tension, via modulating the flow of water in response to changes in 151 osmolality (Ozu et al., 2018). They may also contribute to the activation of TonEBP along with other proposed mechanisms (Moeckel et al., 2006;Kino et al., 2009

Conclusion
AQP1 and 5 expression is decreased during human IVD degeneration (Johnson et al., 2015). This study has identified that AQP1 and 5 are upregulated by hyperosmolality, mimicking the healthy NP, and may explain why expression is decreased during degeneration, when the osmolality is decreased. This suggests AQP1 and 5 may be part of the mechanisms that allow NP cells to adapt to their hyperosmotic environment. TonEBP has already been established as an integral part of those mechanisms; as the hyperosmotic upregulation of AQP1 and 5 is dependent on TonEBP, this implies they both participate in the osmoadaptation process and matrix synthesis (Figure 3.10). During IVD degeneration TonEBP function is uncoupled from the altered osmolality and catabolic genes are upregulated instead of the classical osmotic response genes, which may also include AQP1 and 5. As expression of these genes (and AQP1 and 5) is reduced, NP cells can no longer adapt to the degenerate environment and degeneration is exacerbated (Figure 3.10).
Therefore, AQPs may play a role in adapting NP cells to their environment and maintaining the function of NP tissue. Hyperosmotic upregulation of AQP1 and 5 is controlled by TonEBP, along with the classical osmotic response genes. Therefore, possibly indicating that AQP1 and 5 also enable the adaptation of NP cells to their environment and contribute to the maintenance of matrix expression during healthy physiology. There is evidence that AQPs may also function as osmosensors when membrane tension is altered, potentially implicating AQP1 and 5 in the activation of TonEBP, however this is speculative at present (dashed arrow). During IVD degeneration osmolality is decreased and cytokines, such as TNF-α, are present. This may lead to the activation of TonEBP by non-canonical pathways which changes the genes TonEBP targets. TonEBP now favours catabolic gene expression over osmotic and matrix genes, contributing to degeneration. This may explain why AQP1 and 5 expression is decreased during IVD degeneration; decreased osmolality means TonEBP no longer increases expression of osmotic response genes (including AQP1 and 5), NP cells fail to adapt to their altered environment and start to produce catabolic factors that intensify the degenerative cascade.

154
Chapter 4: Effect of AQP4 and TRPV4 function on the water permeability of human nucleus pulposus cells 155

Introduction
The local environment of the NP is hyperosmotic, when compared to many other tissues (Ishihara et al., 1997). Yet, the osmolality never remains constant as loading applied onto the disc forces water and ions out of the tissue as proteoglycans are compressed (McMillan, Garbutt and Adams, 1996). Therefore, the IVD and its environment undergo a diurnal cycle where NP cells must adapt to daily fluctuations in tissue osmolality and mechanical loading. The diurnal cycle of loading and consequential changes to the NP environment are important for regulating the correct function of NP cells (Ishihara et al., 1997;Wuertz et al., 2007;Neidlinger-Wilke et al., 2012;O'Connell, Newman and Carapezza, 2014). During degeneration, when proteoglycan content of the IVD is reduced, the osmolality and mechanical loading of the tissue is permanently altered, interfering with the usual diurnal cycle. The altered environment contributes to degenerative processes, during which cells lose their ability to adapt to osmotic and mechanical cues and start producing catabolic factors (Le Gilbert, Hoyland and Millward-Sadler, 2010;Gilbert et al., 2011;Sowa et al., 2012). The ability of cells to perceive and respond to their environment is essential for their correct function, yet the exact mechanisms NP cells employ to adapt to the osmotic and mechanical environment, and how these are altered during degeneration, aren't completely understood.
This thesis has identified that AQPs are upregulated by the hyperosmotic environment of the healthy IVD and their expression is regulated at least in part by TonEBP, indicating roles relating to the adaptation of NP cells to their environment (Chapter 3). An adaptation that is lost during degeneration, when AQP expression is decreased (Johnson et al., 2015). However, the regulation of AQPs by osmolality in the IVD does not determine how they function or may contribute to physiological adaptations and cellular dysregulation during degeneration.

AQP function and cellular water permeability
Changes in extracellular osmolality causes flux of water, which results in cell swelling or shrinkage, depending on the osmotic gradient. The ability of cells to regulate their volume is essential for the maintenance of cellular homeostasis. AQPs allow the bidirectional movement of water and other small solutes through the cell membrane and control the water permeability of cells in response to even very small osmotic gradients (Agre et al., 2002). Thus, AQPs provide cells a means by which to alter their volume in response to changes to the osmotic environment. AQP expression and function allows the control of water permeability, enabling fundamental functions of cells (Krane et al., 2001b;Tanaka and Koyama, 2011;Day et al., 2014;Galan-Cobo, Ramirez-Lorca and Echevarr, 2016;Ozu et al., 2018). However, changes in cell size do not go unchecked; cells initiate complex mechanisms to regulate volume changes which prevent excessive swelling or shrinkage which could lead to cell damage.

TRPV4
Cells have adapted complex mechanisms to sense and respond to environmental stimuli. A group of transmembrane proteins that play a key role in adaptation to the environment are TRP channels. The TRP channel family (of which there are 28 members) is classified into six subfamilies that enable the cell to respond to a variety of environmental stimuli by altering the intracellular Ca 2+ concentration. These changes in Ca 2+ concentration induce transduction of signalling pathways, enabling cells to respond to their environment (Samanta et al., 2018). The TRPV subfamily (TRPV1-6) were 157 thought to be activated by alterations in temperature as the first member to be identified, TRPV1, is a nociceptor activated by temperature (>42°C) and capsaicin (Caterina et al., 1997). However the remaining TRPV family members have diverse functions, TRPV4 has been shown to play an important role in the cellular response to osmotic and mechanical stress (Liedtke et al., 2000;Strotmann et al., 2000;Nilius et al., 2001;Köhler et al., 2006;Toft-Bertelsen, Larsen and Macaulay, 2018) TRPV4 is expressed in a wide variety of tissues (Everaerts, Nilius and Owsianik, 2010a) and is activated by hypo-osmotic stimuli, when cell volume increases, enabling the influx of Ca 2+ . This leads to the activation of Ca 2+ -dependent K + channels (Arniges et al., 2004) and RVD mechanisms (Liedtke and Friedman, 2003;Wu et al., 2007)   Function of TRPV4 during hypo-osmotic stress and activation of regulatory volume decrease mechanisms. Cell swelling during a decrease in extracellular osmolality activates TRPV4 and causes rapid Ca 2+ influx. Intracellular Ca 2+ activates dependent K + channels and K + is transported out of the cell. AQPs transport water out of the cell along the osmotic gradient produced by K + efflux, decreasing cell volume and restoring the osmotic balance inside and outside of the cell.
However, there have been limited studies on TRPV4 expression and function in the IVD. It has been shown that exposing mouse MSCs to low magnitude compression enables their differentiation to an NP cell-like phenotype via a TRPV4-dependent pathway (Gan et al., 2018) and in bovine NP cells TRPV4 expression was upregulated by hypo-osmolality (Walter et al., 2016). The same study also showed preliminary data that suggested TRPV4 expression in human IVDs was decreased during degeneration, however results were only reported for 2 human discs (Walter et al., 2016). However, no studies to date have identified how TRPV4 function potentially affects NP cell physiology in response to osmotic stimuli.
However, TRPV4 has a more established role in cartilage physiology.

Mechanotransduction of human articular chondrocytes under dynamic loading has been
shown to be TRPV4-mediated, which regulates expression of matrix genes (O'conor et al., 2013). Furthermore, TRPV4 plays an important role in regulating RVD mechanisms in chondrocytes after hypo-osmotic stimuli (Lewis, Feetham and Barrett-Jolley, 2011), indicating that TRPV4 contributes to healthy cartilage physiology. Indeed, mutations to TRPV4 have been shown to reduce channel activity and induce osteoarthritis (Lamandé et al., 2011) and deletion of TRPV4 in a mouse model leads to a lack of osmoticallydriven Ca 2+ influx and the onset of osteoarthritis (Clark et al., 2010). As TRPV4 is clearly important for the correct function of chondrocytes and a lack of expression/function leads to osteoarthritis, which has similar pathology to IVD degeneration; TRPV4 may also be implicated in the adaptation of NP cells to their hyperosmotic environment and the overall health of the disc.
Interestingly, there is also evidence that the functions of AQPs and TRPV4 may be linked. The ability of many cell types to sense changes in extracellular osmolality and 160 trigger RVD mechanisms may rely on AQPs and TRPV4 functioning in synergy. In rabbit cortical collecting duct cells hypo-osmotic activation of TRPV4 and RVD mechanisms were reliant on AQP2 expression (Galizia et al., 2012). Similar activation of TRPV4 and RVD was reliant on AQP5 in mouse salivary gland cells  and AQP4 and TRPV4 have been shown to interact to control RVD in mouse retinal Müller cells (Jo et al., 2015). AQP4 has also been shown to co-localise with TRPV4 in mouse astrocytes and form a complex that controls the regulation of cell volume (Benfenati et al., 2011a) and Mola et al., 2016 identified that AQP1 and 4-influenced swelling was the main trigger for RVD and TRPV4 mediated calcium signalling in mouse astrocytes. This regulatory role suggests AQPs have many fundamental roles in cell biology other than their function as a passive water channel. Which have been suggested as one of many mechanisms that potentially enable NP cells to sense changes in their extracellular environment (Sadowska et al., 2018). As AQP4 is the isoform that is most permeable to water (Yang and Verkman, 1997;, understanding its role in NP cells and whether it works in unison with TRPV4, to enable adaptation to the fluctuating osmotic environment is important to determine. Especially as a decrease in AQP4 is seen in IVD degeneration (Chapter 2), which could result in a loss of adaptation, when the environment becomes hypo-osmotic.
To determine the physiological roles of AQP4 and TRPV4 in the IVD, this study investigated the expression of TRPV4 in native human NP tissue to determine if levels were altered during degeneration. Furthermore, how human NP cells adapt to changes in extracellular osmolality was determined by measuring changes in cell volume and the rate at which cell size changed within altered osmolality. In addition, the potential involvement of AQP4 and TRPV4 was investigated using specific inhibitors and the water permeability of NP cells following osmotic challenge was determined in the presence and absence of channel inhibitors. Finally, calcium influx in human NP cells, when exposed to altered osmolality, was investigated as changes in extracellular osmolality often cause calcium flux in cells which leads to cell volume regulation. To determine whether AQP4 and TRPV4 facilitated this, calcium influx was investigated in the presence and absence of channel inhibitors.

Experimental Design
To investigate TRPV4 expression, IHC was performed on non-degenerate and degenerate human NP tissue and co-localisation of AQP4 and TRPV4 in human NP cells was determined using confocal microscopy. To investigate how human NP cells responded to shifts in extracellular osmolality, human NP cells were exposed to rapid physiological alteration to osmolality to mimic healthy and degenerate conditions. A plate-reader based assay, utilising the self-quenching effects of Calcein-AM in response to relative intracellular concentration, was used to determine the rate at which NP cells respond to extracellular osmolality changes by modulating their size (Fenton et al., 2010). To determine the actual change in cell volume to rapid changes in osmolality, flow cytometry and fluorescent microscopy methods were utilised. Measurements taken from both the rate of cell volume change and the actual cell volume change were used to determine the relative water permeability from equations outlined by Fenton et al., 2010. Calcium influx in response to altered osmolality, an important cellular adaptation response, was measured using Fluo-4 direct calcium assay in a plate reader setup. To investigate the effects of AQP4 and TRPV4 function on these processes, NP 162 cells were either treated with specific inhibitors to block channel function or left as controls during all experiments. The experimental design is outlined in Figure 4.2.

Human Tissue, NP cell extraction and culture
Human IVD tissue was obtained with ethical consent, NP cells extracted and monolayer cultured according to methods outlined in section 2.2.

TRPV4 gene expression in native human NP tissue
The gene expression of TRPV4 was identified from directly extracted RNA, subsequent cDNA synthesis and qRT-PCR of non-degenerate and degenerately graded human NP tissue, using methods outlined in section 2.2. Pre-designed primer/probe mix for human TRPV4 gene (Hs01099348_m1, Life Technologies) was used and normalised to GAPDH and 18s.

Rate of human NP cell swelling and shrinkage
Human NP cells were seeded into black-walled 96-well plates (Corning) at a density of 1x10 4 cells/well for 48h prior to experiments. Calcein, AM (C1430, Invitrogen) stock at 5mM in DMSO was produced. Cells were loaded with 5μM calcein, AM (1:1000 from stock) in standard culture media for 90min at 37°C. After calcein incubation, cells were washed twice with standard culture media, a final volume of 75μL standard culture media was added per well and cells were allowed to equilibrate to new media for at least 5min at 37°C. Prior to experiments, protocols to measure the rapid rate of change in calcein fluorescence (Figure 4.3) within cells treated with altered osmolality media were set up on a CLARIOstar plate reader (BMG Labtech). Utilising the well mode to detect fluorescent intensity, protocols was set up with a preset of calcein optical settings (ex. 483-14, em. 530-30). Two kinetic windows were setup: a baseline reading of 5s (time -5s to 0s) at 50ms intervals, followed by injection (at time 0s) of 75μL osmotically altered media and a final kinetic window reading for 50s (0s -45s) at 50ms intervals (total of 1000 intervals). Protocol setup and timing is shown in Figure 4.4. The fluorescence of intracellular calcein is dependent on its relative concentration within the cell. At higher intracellular concentrations, when the cell shrinks in response to hyperosmolality, calcein undergoes self-quenching; at lower intracellular concentrations, when the cell swells in response to hypo-osmolality, calcein fluorescence increases. These rapid changes in calcein fluorescence can be recorded on a plate reader and the rate of cell volume change determined (Fenton et al., 2010). The process of calcein self-quenching is not completely understood, but it may depend on dye dimerisation, energy transfer to non-fluorescent dimers and collisional quenching interactions between dye monomers (Chen and Knutson, 1988). 168 The protocol temperature was set to 37°C and the plate reader injector and tubing was primed with an appropriate amount of treatment media prior to experiments. The production of treatment media from standard culture media is outlined in  Table 4.1 Production of the final osmotically altered treatment media, within a physiological range, to induce NP cell swelling and shrinkage when 75μL is injected into wells containing 75μL of standard culture media. All media used in experiments was phenol red-free.
Plates were loaded into the plate reader and protocols were performed on NP cells from 3 patients in triplicate to determine the rate of human NP cell swelling and shrinkage in response to physiological alterations in extracellular osmolality.
Experiments were repeated after 1h incubation of NP cells (already seeded into 96-well plates at 1x10 4 cells/well) with 300μM AQP4 inhibitor (AQP4i) (TGN 020, Tocris) or 4.8μM TRPV4 inhibitor (TRPV4i) (HC-067047, Sigma-Aldrich) to determine the function of these channels on the rate of NP cell swelling and shrinkage. During inhibition experiments, all washes and treatment injections were performed with media containing 300μM AQP4i or 4.8μM TRPV4i when appropriate. Relative fluorescence of calcein (F1/F0) over time (s) curves were plotted and the rate of the change in F1/F0 was determined by fitting plateau followed by one phase association/decay nonlinear regression analysis to curves using GraphPad Prism v7.03 software.

Cell size determination
Human NP cells (from 3 individual patients in triplicate) were trypsinised and resuspended in standard culture media at a density of 2x10 5 cells/mL. Live cells were labelled with CFSE -Cell Labelling Kit (1:1000, ab113853, Abcam) for 10min at RT protected from light and washed before being resuspended in 1mL standard culture media (without phenol red). An equivalent amount (1mL) of altered osmolality treatment media (Table 4.1) was added to cells for approx. 30s (time taken for calcein F1/F0 curves to plateau) before forward scatter (FSC) (proportional to cell size) was determined on 10,000 events using a FACSCalibur flow cytometer (BD Biosciences) and CellQuest Pro v5.2.1 software (BD Biosciences). Furthermore, to determine actual cell size after 30s treatment, cells at a density of 2x10 5 cells/mL were added to Countess™ Both cell surface area and cell volume measurements, along with the rate of change in F1/F0 over time, were used to determine the water permeability of human NP cells in response to altered extracellular osmolality (Section 4.2.8). All cell size determination experiments were also performed after 1h incubation of NP cells with either AQP4i (300μM TGN 020, Tocris) or TRPV4i (4.8μM HC-067047, Sigma-Aldrich), which were also added to each wash, resuspension and CFSE incubation steps throughout the experimental procedure.

Water permeability determination
Using values calculated from the rate of NP cell swelling and shrinkage (section 4.2.6) and NP cell size determination (section 4.2.7), the water permeability of human NP cells in response to alterations in extracellular osmolality, and the potential involvement of AQP4 and TRPV4, was determined using Equation 4.4.
.4. Water permeability. Where Pf = water permeability, K = rate constant for one phase decay/association, V0 = starting cell volume, Vmin 2 = cell volume after treatment, A = cell surface area, Δπ = applied osmotic gradient, Vw = partial molar volume of water (1.8x10 -5 m 3 /mol). As [V0 -Vmin 2 ] reflects the total cell volume change in response to the change in extracellular osmolality, K x [V0 -Vmin 2 ] describes the theoretical fraction of total volume change per one x-axis unit change (50ms) at the initial rate of volume change. Adapted from (Fenton et al., 2010).
Average Pf of human NP cells, for each patient, was calculated in response to physiological changes in osmolality (325mOsm/kg → 225, 425 or 525mOsm/kg) with or without AQP4i and TRPV4i. All Pf values were normalised to the average Pf of controls in the absence of inhibitors, to determine the effect of AQP4i and TRPV4i on the Pf of human NP cells at each extracellular osmolality change.

Fluo-4 Direct™ Calcium influx assay
Calcium influx into human NP cells in response to extracellular osmolality alterations was measured using Fluo-4 Direct™ calcium assay kit (F10471, Invitrogen).
Human NP cells were seeded into black-walled 96-well plates (Corning) at a density of 1x10 4 cells/well for 48h prior to experiments. Fluo-4 Direct™ (2x) calcium reagent loading solution (Invitrogen) was prepared following the manufacturer's guidelines: 10mL Fluo-4 Direct™ calcium assay buffer was added to one bottle of Fluo-4 Direct™ calcium reagent (component A). Media was aspirated from cells and an equal volume of Fluo-4 Direct™ (2x) calcium reagent loading solution (37.5μL) and standard culture media (37.5μL) was added to cells, for a total volume of 75μL per well. Plates were incubated for 1h at 37°C protected from light. Following incubation, plates were ready for experimental use without removing Fluo-4 Direct™ (2x) calcium reagent loading solution and standard culture media from wells. Calcium influx into human NP cells in response to physiological alterations of extracellular osmolality (325mOsm/kg →225, 325, 425 or 525mOsm/kg) was performed on a CLARIOstar plate reader (BMG Labtech) following the same protocol outlined in section 4.2.6, with optic settings changed to excitation at 494nm and emission at 516nm. Treatment media (Table 4.1) injected into wells after the first kinetic window (-5s to 0s). Experiments were also performed after 1h incubation of cells with either AQP4i (300μM TGN 020, Tocris) or TRPV4i (4.8μM HC-067047, Sigma-Aldrich), simultaneously added to cells with Fluo-4 Direct™ (2x) calcium reagent loading solution and media. During AQP4i and TRPV4i experiments, inhibitors were also added to treatment media prior to injection.

Statistical Analysis
TRPV4 protein expression in human NP tissue (assessed by IHC) was found to be non-parametric, therefore Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences between expression across grades of degeneration. The rate of cell swelling/shrinkage (determined by calcein fluorescence) was investigated in triplicate in human NP cells from 3 patients. Data was non-parametric and to determine significance between paired samples from the same patient across 3 treatment groups (CTR, AQP4i and TRPV4i), the Friedman test was used with Conover post hoc analysis. Cell volume data was non-parametric, therefore Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences in cell volume, at a fixed osmolality, when comparing 173 treatment groups (CTR, AQP4i and TRPV4i). The Friedman test was used to determine significance between the geometric mean of the FSC, and the Pf, from paired patient samples when comparing 3 treatment groups (CTR, AQP4i and TRPV4i), at a fixed osmolality. Ca 2+ influx data was non-parametric, therefore Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences in the max-min F1/F0 and time taken to reach maximum F1/F0 in NP cells treated with different osmolalities, or inhibitors (CTR, AQP4i and TRPV4i) at 225mOsm/kg.

Expression of TRPV4 within native human NP tissue
TRPV4 was expressed at both gene and protein level within native human NP tissue (Figure 4.5). The relative gene expression of TRPV4 did not change significantly during IVD degeneration (p ≤ 0.05) (Figure 4.5A). The majority of NP cells expressed TRPV4 protein in native tissue, however, immunopositivity was unaffected by IVD degeneration (Figure 4.5B) (IgG controls, Appendix VIII). TRPV4 was also still expressed by NP cells extracted from human IVD tissue at passage 2. (Figure 4.5C) Within 2D cultured human NP cells TRPV4 was found to co-localise with AQP4 ( Figure 4.5C), potentially suggesting related functions shared between these transmembrane channel proteins (IgG controls, Appendix VIII). Each point represents an individual patient. NP tissue was graded: non-degenerate (0-4, B.i), moderately-degenerate (4.1-6.9, B.ii) and severely-degenerate (7-12, B.iii). Positive staining for TRPV4 is indicated by brown staining and cell nuclei are counterstained blue with Mayer's haematoxylin. (C) Immunofluorescence highlighting the expression of TRPV4 (green) in 2D cultured human NP cells co-localises with AQP4 (red). Cell nuclei counterstained with DAPI (blue). Images taken at 63x objective magnification, using a LSM 800 confocal microscope (Zeiss). Scale bar 20μm. Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

Change in NP cell volume in response to extracellular osmolality alterations
The volume of AQP4i and TRPV4i treated human NP cells exposed to 225mOsm/kg was significantly reduced, when compared to no-inhibition control (CTR) NP cells (p ≤ 0.05) (Figure 4.8A). The volume of TRPV4i treated NP cells was significantly reduced compared to CTR cells at 325mOsm/kg (p ≤ 0.05) (Figure 4.8B).

Effect of AQP4 and TRPV4 inhibition on the water permeability of human NP cells
The normalised water permeability of human NP cells was significantly reduced with AQP4i and TRPV4i, when compared with no-inhibition control cells, at both 225mOsm/kg (p ≤ 0.05) (Figure 4.12A) and 425mOsm/kg (p ≤ 0.05) ( Figure   4.12B). At 525mOsm/kg treatment, the normalised water permeability of TRPV4i treated NP cells was also significantly reduced when compared to no-inhibition control cells (p ≤ 0.05) (Figure 4.12C). AQP4i did not significantly change the water permeability of human NP cells, when compared to no-inhibition controls, at 525mOsm/kg treatment (Figure 4.12C). and averaged per patient. The normalised Pf of AQP4i (1h, 300μM, TGN 020) and TRPV4i (1h, 4.8μM, HC-067047) was compared to CTR at (A) 225, (B) 425 and (C) 525mOsm/kg treatments. Prior to treatment, media osmolality was 325mOsm/kg. Statistical significance determined using Freidman test * = p ≤ 0.05.

Human NP cell Ca 2+ influx in response to altered extracellular osmolality
The F1/F0 of intracellular Fluo-4 direct, relative to Ca 2+ , was altered in realtime, when NP cells were exposed to altered extracellular osmolality (225,325,425 and 525mOsm/kg), after 5s baseline readings in standard culture media (325mOsm/kg) (Figure 4.13A). The max-min F1/F0 of intracellular Fluo-4 direct was significantly increased in 225mOsm/kg treated NP cells, compared to all other osmotic treatments (p ≤ 0.05) (Figure 4.13B).  Ca 2+ influx in human NP cells when exposed to altered extracellular osmolality. (A) Fluo-4 direct assay (Invitrogen) was used to measure the rate of Ca 2+ influx into human NP cells. Baseline fluorescence of calcein-loaded NP cells was recorded for 5s (-5s -0s), before injection of altered osmolality media and change in relative Fluo-4 fluorescence (F1/F0), dependent on the intracellular Ca 2+ concentration, was recorded for a further 45s (0s -45

Effect of AQP4 and TRPV4 inhibition on hypo-osmotic Ca 2+ influx in human NP cells
The F1/F0 of intracellular Fluo-4 direct, relative to Ca 2+ , was altered in realtime, when NP cells were exposed to 225mOsm/kg extracellular osmolality after AQP4i, TRPV4i or CTR treatment, and 5s baseline readings in standard culture media  Effect of AQP4 and TRPV4 inhibition on the hypo-osmotic induction of Ca 2+ influx. (A) Fluo-4 direct assay (Invitrogen) was used to measure the rate of Ca 2+ influx into human NP cells when exposed to hypo-osmotic treatment (225mOsm/kg). Baseline fluorescence of calcein-loaded NP cells was recorded for 5s (-5s -0s), before injection of 225mOsm/kg media and change in relative Fluo-4 fluorescence (F1/F0), dependent on the intracellular Ca 2+ concentration, was recorded for a further 45s (0s -45

Discussion
The hyperosmotic environment of the IVD is constantly changing due to diurnal loading, where water is imbibed and then dissipated as the mechanical loading of the spine changes. In physiologically-matched osmolality, matrix expression is increased in NP cells (Wuertz et al., 2007;Neidlinger-Wilke et al., 2012;O'Connell, Newman and Carapezza, 2014), indicating NP cells have adapted their function to the hyperosmotically fluxing environment within the IVD. Therefore, mechanisms must be in place to protect NP cells, and enable their adaptation to hypo-and hypertonic shifts in extracellular osmolality. It is well documented that TonEBP contributes to NP cells' long-term adaptation to hyperosmolality (Chapter 3) and the expression of matrix molecules (Tsai et al., 2006;Johnson, Shapiro and Risbud, 2014b). However, the mechanisms that control the initial response to altered extracellular osmolality, such as the rapid flux of water and ions and changes in cell volume, in NP cells are not completely understood. Especially how hypo-osmotic conditions, observed during IVD degeneration, may impacts these fundamental cellular processes and the overall function of the IVD.
This study identified that NP cells rapidly modulate their cell volume and the rate which volume is changed in response to extracellular osmolality alterations that mimicked the environment of the healthy and degenerate IVD.

NP cell volume change in response to extracellular osmolality
Upon exposure to alterations in extracellular osmolality, cells adapt by altering their volume accordingly via osmotically driven water transport and the activation of regulatory volume changes (Corasanti, Gleeson and Boyer, 1990;Lang et al., 1998). This study has shown that human NP cells are no different; under hypoosmotic treatment cell volume was increased, and under hyperosmotic treatment, cell volume was reduced. These results may indicate that during IVD degeneration, when osmolality is decreased compared to healthy conditions, human NP cell size increases. Bovine NP cells show a similar trend when exposed to low osmolality ) and rabbit NP cell size was also increased after an AF puncture model of degeneration (He et al., 2013), indicating that NP cell size could possibly be used to determine the degenerative state of the IVD.
Fluorescent microscopy was used to quantify the volume of CFSE-labelled NP cells when exposed to altered osmolality. Flow cytometry, specifically shifts in FSC, was used alongside microscopy to determine how NP cell size was altered. Shifts in FSC have been previously used to determine changes in cell size after osmotic alteration (MacDonald and Zaech, 1982;Sloot, Hoekstra and Figdor, 1988;Adan et al., 2017) and NP cell size has been characterised using the same method (Chen, Yan and Setton, 2006). As results from independently analysed microscopy and flow cytometry experiments on NP cell size both indicate a similar response to osmotic stimuli, this strengthens the observations presented in this study.
However, FSC measurements may not be directly related to the changes in cell size, as it may alter depending on the light scattering properties of different cells (McGann, Walterson and Hogg, 1988;Adan et al., 2017;Yurinskaya et al., 2017), therefore flow cytometry results presented here may only reflect the relative change in NP cell size, rather than the actual change in size. Furthermore, the actual size of human NP cells could not be determined from flow cytometry measurements because the FSC of NP cells could not be integrated with the FSC of size calibration beads, indicating again that FSC is not always directly proportional to cell size. Many properties may effect FSC, such as cell hydration state (Yurinskaya et al., 2017). This may explain why the FSC of NP cells and calibration beads did not align; as the pericellular matrix surrounding NP cells in culture (Sato et al., 2001, Sato et al., 2004Horner et al., 2002)  . This would be a more accurate measure of cell volume in response to altered osmolality (Fenton et al., 2010), enabling measurements of precise volume change in NP cells.

Role of AQP4 and TRPV4 on NP cell size
When exposed to decreased extracellular osmolality (225mOsm/kg), NP cell volume was increased compared to osmotic controls (325mOsm/kg). In the presence of either AQP4i or TRPV4i, the hypo-osmotic increase in NP cell size was significantly reduced. These results indicate that the function of both AQP4 and TRPV4 is required for NP cells to respond in the correct manner to decreased osmolality, by increasing volume. Potentially indicating that both channels work synergistically, enabling NP cells to correctly regulate their volume and trigger downstream mechanisms (such as RVD), in order to adapt to their environment. This synergistic function of AQP4 and TRPV4 has been observed in astrocytes and the regulation of the blood-brain-barrier previously (Solenov et al., 2004;Jo et al., 2015;Mola et al., 2016).
Cell volume responses to increased osmolality (425 and 525mOsm/kg) showed no significant change, possibly meaning there is no effect of channel inhibition or the methods used to measure cell volume were not sensitive enough to detect the decrease in cell size that normally follows a decrease in extracellular osmolality.

The rate of NP cell volume change in response to altered osmolality
This is the first time the rate of NP cell volume change in response to altered osmolality, and the effects of AQP4 and TRPV4 inhibition, has been investigated. The rate of NP cell swelling and shrinkage was determined using calcein-AM loaded cells in a plate reader-based method, used previously to investigate volume change rates and AQP function within vesicles, proteoliposomes, MDCK and CHO cell lines, astrocytes, erythrocytes and fibroblasts (Mola et al., 2009;Fenton et al., 2010;Kitchen et al., 2016).
Calcein-AM has been used in experimental systems such as vesicles (Chen and Knutson, 1988), proteoliposomes (Zeidel et al., 1992), and mammalian cells (Solenov et al., 2004) to determine rapid changes in volume and AQP function. These measurements are achieved by the self-quenching properties of calcein at higher concentrations (Chen and Knutson, 1988) and insensitivity to pH, Ca 2+ or NaCl concentration (Wehner et al., 1995;Solenov et al., 2004). Hamann et al., 2002 were the first to discover that the fluorescent intensity of calcein-loaded mammalian cells was rapidly altered according to the osmotic challenge applied (increase during hypoosmotic, decrease during hyperosmotic), demonstrating the usefulness of calcein to measure the rapid changes and rate of cell volume. The first methods utilising this technique relied on fluorescent microscopy (Wehner et al., 1995;Hamann et al., 2002;Solenov et al., 2004), more recently plate reader methods have been developed which enable rapid and reproducible investigations into the rapid realtime rate of cell volume changes and screening of membrane protein modulators (Mola et al., 2009;Fenton et al., 2010).
This study followed protocols of Fenton et al., 2010, who determined how calcein fluorescence intensity was related to cell volume, and speculated that a variety of cell types could be tested with their protocols (Fenton et al., 2010). This is also the first time that this method has been used to determine the rate of cell volume change in response to extracellular osmolality in human NP cells. Calcein fluorescence depends linearly on extracellular osmolality in MDKC cells (Fenton et al., 2010), which has also been shown to be the case for human NP cells (Appendix X

Control of NP cell water permeability
197 The water permeability of NP cells was determined by combining cell volume changes and the rates of change in volume (Fenton et al., 2010). AQP4i decreased the water permeability of NP cells when exposed to healthy (425mOsm/kg) and degenerate (225mOsm/kg) osmolalities, but not when NP cells were exposed to higher osmolality (525mOsm/kg). This may indicate that AQP4 only controls NP cell responses under specific conditions within the microenvironment of the IVD. It has previously, been shown that AQP4 driven water permeability may depend on cell membrane compression and tension (Tong, Briggs and McIntosh, 2012;Ozu et al., 2018), which changes as cells respond to altered osmolality. Thus, AQP4 may exert a greater effect on NP cells within a certain osmotic range, and multiple other AQPs expressed by NP cell may contribute outside this range.
TRPV4 function on the other hand, significantly impacted on the water permeability of NP cells across all osmolalities and was expressed by the vast majority of NP cells regardless of degenerative state. This indicates a potential overarching role during NP cell physiology and adaptation to the hyperosmotic environment of the IVD. TRPV4 is known to enable chondrocyte mechanotransduction and matrix synthesis (O'conor et al., 2013) and lack of TRPV4 expression can induce osteoarthritis in animal models (Clark et al., 2010;Lamandé et al., 2011), thus may contribute to similar functions in the IVD. TRPV4 contributes to NP cell water permeability, yet it is not a water channel itself. Therefore, TRPV4 function must somehow be linked to the function of AQPs and cell volume regulation. TRPV4 has been shown to have physical and functional links with AQPs to enable cells to sense extracellular osmolality changes and initiate cell volume responses Benfenati et al., 2011a;Galizia et al., 2012;Jo et al., 2015;Mola et al., 2016). TRPV4 was also found to co-localise with AQP4 in human NP cells suggesting related functions and interaction between both channels.
Increased osmolality treatments (425 and 525mOsm/kg) used in these experiments were produced using sucrose as the key osmolyte. This ensured that the actual water permeability of NP cells was able to be determined. NP cells contain transmembrane channel proteins that readily transport osmolytes such as Na + and Cl + , urea and mannitol. Therefore, if any of these were used to alter osmotic treatments, cell volume may have reached equilibrium at a different rate as NP cells could utilise the movement of osmolytes used to regulate volume changes, rather than employing the actual mechanisms used to regulate cell volume. However, as IVD matrix is negatively charged, one of the main ions imbibed into the tissue will be Na + (Urban, 2002). So repeating experiments using NaCl will enable the comparison between methods of changing medium osmolality (determining if results are an osmotic or substrate effect) and may be more physiological to the in vivo environment of the IVD.

Ca 2+ influx in response to extracellular osmotic shifts
Cellular response to osmotic shifts in their environment is to modulate calcium flux to trigger downstream pathways to enable adaptation to altering environment. TRPV4 in particular has a well-established role in regulating the cellular response to hypo-osmotic shifts in extracellular osmolality, where calcium influx is increased. This enables the triggering of RVD mechanisms and shrinkage to adapt (Caterina et al., 1997;Liedtke and Friedman, 2003;Arniges et al., 2004;Everaerts, Nilius and Owsianik, 2010;Toft-Bertelsen, Larsen and Macaulay, 2018). NP cells showed they also increase Ca 2+ influx under hypo-osmotic conditions, compared to hyper-osmotic conditions, suggesting they may employ similar pathways to adapt to their diurnal environment. Previous research has also shown that NP cells do not increase calcium influx under similar hyperosmotic conditions. (Pritchard, Erickson and Guilak, 2002). Pritchard, Erickson and Guilak, 2002 also identified that Ca 2+ influx and cell volume may be controlled by an actin cytoskeletal-dependent mechanism.
Which in turn could potentially be regulated by AQP4; as in astrocytes AQP4 knockdown lead to drastic rearrangement of the actin cytoskeleton network and reduced water permeability (Nicchia et al., 2005). Cytoskeletal rearrangement under alterations in osmolality have also been observed in bovine NP cells . However, when NP cells were injected with control media (325mOsm/kg, same as pre-injection media) a Ca 2+ influx response was still observed. This highlights that Ca 2+ influx may also be due to mechanical pressure applied onto NP cells from the injection system of the plate reader, as TRPV4 and other Ca 2+ channels are also known to be mechanotransducive (Samanta et al., 2018). This may indicate Ca 2+ influx observed was not solely attributable to the extracellular osmolality, however hypo-osmotic increased Ca 2+ influx to a greater extent than control injection.
TRPV4i significantly reduced hypo-osmotic Ca 2+ influx, suggesting important NP cell responses to hypo-osmotic stimuli and the potential activation of RVD.
However, Ca 2+ influx was not totally nullified, therefore other channels may also be involved in these processes. AQP4i significantly reduced the time taken for maximum 200 Ca 2+ influx and the maximum Ca 2+ influx, despite its sole function as a water channel.
AQP4 and TRPV4 function is linked to osmosensing and the adaptation to extracellular osmolality of astrocytes (Benfenati et al., 2011a;Jo et al., 2015;Mola et al., 2016;Chmelova et al., 2019), and results described here also indicate there may be similar mechanisms employed by NP cells.

Conclusion
AQP4 and TRPV4 function may be linked in NP cells in order to control the rate of cell volume regulation, water permeability and Ca 2+ influx, triggering downstream mechanisms enabling the adaptation of NP cells to their osmotically challenging environment (Figure 4.15). During IVD degeneration AQP4 expression is decreased (Chapter 2); as a result, NP cells will not be able to respond to the hypoosmotic environment due to impaired cell volume regulation and Ca 2+ influx (

Introduction
This thesis has identified that certain AQPs are regulated by the physiological osmolality of the IVD, which potentially indicates how they are regulated in vivo and alludes to the specific functions of these AQPs surrounding the osmotic adaptation of NP cells. However, the native environment in which NP cells reside is also hypoxic, acidic and has low nutrient supply, when compared to other tissues, and also has to withstand biomechanical loading (Urban, 2002). During degeneration these conditions can be severely altered and the degenerative cascade is exacerbated by the increased production of cytokines, which trigger degenerative processes (Takahashi et al., 1996;Le Maitre, Freemont and Hoyland, 2005;Le Maitre, Hoyland and Freemont, 2007a;Risbud and Shapiro, 2014). Therefore, the functions of NP cells are, at least in part, regulated by their environment and dysregulated during degeneration when the environment is altered. This indicates that AQPs may also be regulated by many of the other environmental cues within the IVD.
AQP expression in many tissues has been shown to be regulated by physiological conditions which change in the IVD during degeneration, such as O2 concentration, cytokines and pH (Tables 5.1-5.3 respectively). However, there have been few studies which have investigated the regulation of AQP expression by these conditions in NP cells. It was determined that AQP1 and 5 expression in rat NP cells was not regulated by changes in O2 concentration, but basal expression levels were maintained under hypoxia by HIF-1α (Johnson et al., 2015) (Table 5.1), indicating that within the native environment of the IVD, where O2 concentration is always low and HIF-1α is stably expressed (Fujita et al., 2012), the maintenance of AQP expression in 204 NP cells is controlled by the environment. In bovine NP cells it was determined that TNF-α decreased expression of AQP1   (Table 5.2), signifying that during IVD degeneration, when TNF-α expression is increased (Le Maitre, Hoyland and Freemont, 2007a), AQP expression may be differentially regulated when compared to healthy IVD physiology. These studies show that potentially more than one environmental factor may contribute to the regulation of AQP expression and function within NP cells. No studies, to date have investigated pH regulation of AQP expression in NP cells, with only a few identifying such regulation in the kidney (Promeneur et al., 2000;Amlal, Sheriff and Soleimani, 2004), where AQP2 and 6 are potentially involved in acid-base regulation (    (Promeneur et al., 2000) Rat kidney AQP2 NH4induced metabolic acidosis ↑ AVP (Amlal, Sheriff and Soleimani, 2004)  The regulation of AQP expression by these conditions occurs across many cell and tissue types and regulation of expression is potentially specific to each type.
Many of the AQPs expressed in NP tissue (Chapter 2) are regulated by conditions in many tissues that are also present during healthy and degenerate IVD physiology, highlighting AQPs may also be regulated in a similar manner in NP cells. The potential regulation of AQP expression by physiological conditions may be an indication of the potential roles specific AQPs contribute towards with the disc.
This study aimed to investigate if AQPs expressed in human NP tissue were expressed in extracted NP cells and whether expression was regulated by culture conditions, including 2D and 3D culture and stimulation by cytokines, hypoxia and pH. Furthermore, these conditions were then combined to mimic the conditions representative of the IVD environment and effects on AQP expression determined. 208

Experimental design
To determine that all AQPs expressed by native human NP tissue (AQP0-7 and 9, Chapter 2) were expressed by human NP cells during culture; IHC was performed on cytospins of human NP cells taken from passage 0 -2 and after 2w alginate bead culture which is utilised to re-differentiate cells to an NP like phenotype. How AQP expression in NP cells is regulated has not been previously established. Therefore, human NP cells, encapsulated in alginate beads to re-differentiate into an in vivo-like phenotype, were treated with cytokines (IL-1β, IL-6 and TNF-α), altered pH (pH7.4 -6.5) under 5% and 21% O2. Furthermore, combined treatments mimicking healthy and degenerate IVD conditions were investigated to determine if AQP gene expression within NP cells is regulated by conditions observed during IVD physiology.
The experimental design is outlined in Figure 5.1.

209
Figure 5.1. Experimental design of methods used in this study.

Human Tissue, NP cell extraction and culture
Human IVD tissue was obtained with ethical consent, NP cells extracted and monolayer cultured according to methods outlined in section 2.2.

Cytospins
Human NP cells were taken at passage 0 (straight after isolation from NP tissue), at passage 1, passage 2 during splitting and after 2 week culture and isolation from alginate beads (section 3.2.6), and centrifuged at 300g for 5min. Supernatant was discarded and cell pellets resuspended in 10% neutral buffered formalin for 5min. Fixed cells were centrifuged at 300g for 5min and pellets resuspended in PBS at 1x10 6 /mL. Cytospins, filter cards (Thermo Fisher, UK) and slides (Leica Microsystems) were assembled and 200μL of cell suspension was added to each cytospin and centrifuged at 300g for 5min to deposit a single layer of cells onto slides.
Slides were stored at 4°C. IHC was performed (Section 2.2.7) to determine if the expression of AQPs in native human NP tissue (AQP0-7, 9) was retained in isolated human NP cells during culture duration and conditions used in experiments.

Regulation of AQP gene expression under physiological IVD conditions
At passage 2, human NP cells were encapsulated into alginate beads as previously described (section 3.2.6). To assess catabolic cytokine regulation of AQP gene expression in human NP cells, alginate beads were treated with 0 (control), 1, 10 and 100ng/mL recombinant human IL-1β (200-01B, Peprotech), recombinant human IL-6 (200-06, Peprotech) or recombinant human TNF-α (300-01A, Peprotech) for 48h. To determine the effect of physiological pH on the gene expression of AQPs, alginate beads were treated with standard culture media with altered pH: pH 7.4 (media control), pH 7.1 (healthy IVD), pH 6.8 (moderate IVD degeneration) or pH 6.5 (severe IVD degeneration) for 48h. The pH of standard culture media was adjusted using HCl and replaced every 12h to ensure treatment pH was constant and media remained unbuffered. All treatments were performed at 21% (v/v) O2 (to represent standard culture conditions) and 5% (v/v) O2 (to represent physioxia of the native IVD). All treatment conditions were combined to produce a set of treatments to represent the healthy and degenerate IVD niche (Table 5.4), which alginate beads were also treated with for 48h. Following 48h treatment, NP cells were released from alginate beads and RNA extracted (section 3.2.6), cDNA synthesised and the gene expression of AQPs investigated using previously described methods (section 2.2.5).

Statistical analysis
All experiments investigating AQP gene expression by physiological IVD conditions were performed on at least 3 patient-derived human NP cells in triplicate.
Data was shown to be non-parametric, therefore, Kruskall-Wallis with Dwass-Steel-Critchlow-Fligner post hoc analysis test was used to identify significant differences in in AQP gene expression after 48h treatment (p ≤ 0.05).

AQP expression during 2D and 3D culture of human NP cells
Throughout all culture conditions (passage 0 -2 and 2w 3D culture in alginate beads) prior to experimental procedures, immunopositivity was seen for all AQPs which were expressed in native human NP tissue (Chapter 2) ( Figure 5.2).

Regulation of AQPs by cytokines in human NP cells
The gene regulation of all AQPs expressed by human NP tissue (AQP0-7 and 9) was investigated during all experiments, however, results only show AQPs that were detected by qRT-PCR, all other AQP family members were undetected.

AQP1
The relative gene expression of AQP1 was significantly down-regulated by

Regulation of AQPs by pH in human NP cells
The relative gene expression of AQP5 in alginate encapsulated human NP cells was not significantly regulated by physiological changes in extracellular pH (pH7.4 -6.5) for 48h at 21% O2 ( Figure 5.7A) or 5% O2 (Figure 5.7B). Gene expression of AQP5 was also unchanged in 5% O2 when compared to expression in 21% O2 after 48h treatment in standard culture media (pH7.4) ( Figure 5.7C). However, the relative gene expression of AQP5 was significantly up-regulated by 48h culture at 5% O2, when compared to culture at 21% O2, when extracellular pH was altered to pH7.1 (p ≤ 0.05)

Regulation of AQPs by physiological IVD conditions
The relative gene expression of AQP1 and 2 was unaltered when 3D alginatecultured human NP cells were treated for 48h in healthy or degenerate conditions compared to standard culture controls, although an increase was seen in 5% O2 in degenerate conditions this did not reach significance ( (Table 5.4), at 21% (grey) and 5% (magenta) O2 concentration, for 48h. The gene regulation of (A) AQP1, (B) AQP2 and (C) AQP5 was compared to standard media controls (21% O2). Statistical significance determined using Kruskal-Wallis test * = p ≤ 0.05.

Discussion
NP cells have adapted to a hypoxic, hyperosmotic, acidic and mechanically stimulated environment that also has low nutrient diffusion. This environment contributes to the overall function of NP cells to maintain the integrity of the tissue (Bibby et al., 2005;Tsai et al., 2006;Wuertz et al., 2007;Neidlinger-Wilke et al., 2012

AQP gene regulation by cytokines
Cytokines, such as IL-1β, IL-6 and TNF-α, are upregulated and trigger IVD degeneration by altering the expression of many genes (Le Maitre, Freemont and Hoyland, 2005;Le Maitre, Hoyland and Freemont, 2007a;Hoyland, Le Maitre and Freemont, 2008;Phillips et al., 2015). Potential regulation of to investigate changes in AQP 9 protein expression both in vivo and in vitro.

AQP gene regulation by pH
As the IVD is an avascular and hypoxic tissue, NP cells favour anaerobic glycolysis as their main form of metabolism, producing an acidic environment (Bibby et al., 2005). NP cells have adapted several mechanisms to adapt to this environment (Silagi et al., 2018a;Silagi et al., 2018b), yet during degeneration the pH decreases further, as removal of waste products (lactic acid) is reduced. There is limited evidence for pH regulation of AQP expression (Table 5.3), highlighting such regulation is only observed in tissues where pH balance is essential (or because no one has investigated this in other tissues), such as the kidney (Promeneur et al., 2000;Amlal, Sheriff and Soleimani, 2004) and possibly the IVD.
AQP5 gene expression was unchanged by altered pH in human NP cells cultured in 21% or 5% O2, indicating that AQP5 expression may not be sensitive to oxygen tension or pH. However, when NP cells were cultured in physiological pH (pH7.1 -6.5) and physioxia (5% O2) AQP5 expression was up-regulated compared to 21% O2 controls. This indicates that both physiological pH and oxygen tension may contribute to the correct regulation of AQP5 expression in NP cells, whereas individual treatment with either does not. This provides evidence highlighting that multiple environmental cues all provide NP cells with the correct (and incorrect, in the case of degeneration) stimuli to regulate expression and function accordingly, which will be the case in vivo. Hence NP cell treatment with combined treatments mimicking the healthy and degenerate IVD niche was investigated.

AQP gene regulation by combined healthy and degenerate IVD conditions
The regulation of AQP1, 2 and 5 gene expression by healthy and degenerate IVD conditions was unclear. The gene expression of these AQPs was largely unaffected by these conditions at 21% and 5% O2 when compared to standard culture conditions at 21% O2, indicating that AQPs may not be regulated within NP cells during degenerative processes and their functions may be independent of degeneration. However, only changes in oxygen tension, osmolality, pH and cytokines were used to 'mimic' the healthy and degenerate IVD niche, compared to standard culture media. This is a simplified environment and thus, full recapitulation of the degenerate and normal niche is needed to investigate this more accurately.
The pH of treatments was altered using HCl; this may be an ineffective way of altering media pH as desired alterations may be buffered out, an alternative which may be more physiologically relevant could be to use lactic acid. Furthermore, the addition of the appropriate amount of NaHCO3 to media (without NaHCO3), calculated using the Henderson-Hasselbalch equation, will enable the correct maintenance of the desired media pH (Esser, Thermo Scientific, 2010;Gilbert et al., 2016).
The native environment of the NP is far more complicated than the treatments devised in these experiments. Treatments failed to address the mechanical forces (Neidlinger- Wilke et al., 2012), low nutrient diffusion (Urban et al., 1977) and diurnal patterns (McMillan, Garbutt and Adams, 1996) that NP cells are exposed to in vivo. Also, an increased level of calcium deposition within the endplates and IVD has been observed during degeneration (Grant et al., 2016), indicating the 229 inclusion of increased calcium concentration may be important to mimic degenerate conditions as well. Therefore, without including all environmental factors that contribute to IVD health and degeneration, treatments aiming to mimic these complex systems will not be physiologically relevant and should be expanded on, to gain a more complete view of AQP regulation. Furthermore, these physiological conditions may affect AQPs at protein levels rather than gene expression levels, as many post-translational modifications (channel gating, phosphorylation, calcium binding cytoskeletal remodelling and protein interactions) (Yasui et al., 1999;Nicchia et al., 2008;Benfenati et al., 2011b;Conner, Bill and Conner, 2013;Jo et al., 2015;Kitchen, P. et al., 2015) all potentially play a significant role on the expression, regulation, trafficking and function of AQPs during IVD physiology, which should be explored further, particularly due to the issues experienced with gene expression analysis of AQPs within NP cells.

Conclusion
AQPs within NP cells may be regulated by many environmental cues within NP cells, such as cytokines and pH ( Figure 5.9), along with (the well documented) alterations in osmolality. Yet, to date the true regulation and function of AQPs within the IVD is relatively unknown (Figure 5.9). The in vivo conditions of the IVD must be replicated very specifically to gather the information required to unfold the potential roles AQPs, and all other genes and proteins, may play during IVD physiology. Regulation of AQP gene expression in 3D alginate cultured human NP cells. Cytokines present during IVD degeneration upregulate and downregulate AQP gene expression, depending on the individual cytokine and AQP. AQP5 is upregulated by simultaneous physiological pH and O2 tension treatment, indicating many microenvironmental factors may influence AQP expression in human NP cells, in unison. However, there are more factors that contribute to NP cell function in vivo than have been investigated in this study, such as nutrition and mechanical loading. To determine how AQP expression is truly regulated in the IVD, the in vivo conditions must be recapitulated very precisely.

231
Chapter 6: General discussion and future directions 232 IVD degeneration-associated LBP remains a debilitating condition with no treatments currently available that are directed towards halting and reversing the degenerative cascade at a cellular level. The lack of such treatments is in part due to incomplete knowledge of the molecular mechanisms that govern IVD function in health and degeneration. Due to the unique location and role of the IVD within the spine, many factors contribute to the microenvironment that cells reside within. The survival and function of cells has been irrefutably linked to their ability to adapt to the microenvironment in which they live. During IVD degeneration the microenvironment is altered which has a detrimental effect on cell behaviour and, in part, plays a role in the dysfunction that causes cells to display a catabolic phenotype, intensifying IVD degeneration. As such, increasing the understanding of normal cellular functions, and how the IVD microenvironment contributes to this, may help elucidate mechanisms that are altered during degenerative processes. Potentially revealing novel strategies and treatments to target the cellular causes of IVD degeneration.
The overall aim of this thesis was to investigate the expression, regulation and function of AQP transmembrane water channels within the IVD and how they potentially contribute to the adaptation of cells to their environment. Chapter 2 investigated the expression of AQPs in the IVD, specifically how expression may be altered during developmental and degenerative processes. Chapter 3 presented the regulation of AQP expression by hyperosmolality, physiological to the IVD and well known to regulate AQPs in other tissues, and the control of regulation by a transcription factor (TonEBP) implicated in NP cell function. Chapter 4 demonstrated 233 that AQPs contribute to fundamental cellular processes such as the control of cell volume in response to altered extracellular osmolality in NP cells, and that other transmembrane channels work in unison with AQPs to facilitate this. Chapter 5 set out to further characterise that AQP expression in NP cells was also regulated by other microenvironmental factors that contribute to IVD health and degeneration. A summary of key findings from this body of work is provided below.

The expression of AQPs within the intervertebral disc
The IVD is an osmotically challenged tissue which is altered from development to maturation and further changes during degeneration. Therefore, NC and NP cells must employ several tightly regulated mechanisms to survive and function within their unique osmotic environment. Their function as transmembrane water channels, highlighted that many AQP isoforms may be expressed by these cells to enable their adaptation to the surrounding environment. In fact, results here did identify the expression of 9 different AQP isoforms (AQP0 -7 and 9) within the developing and mature IVD, of which 5 had not previously been shown to be expressed (AQP0, 4, 6, 7 and 9). Results presented here imply that the expression of numerous AQP family members in the disc highlights the importance of maintaining intricate control over regulation of water and solute transport and cell volume in NP cells. The increased proportion of NP cells expressing AQP2 and AQP7 during degeneration may reflect consequences or potential repair mechanisms employed by the disc as NP cells lose their ability to adapt to the degenerate hypo-osmolar environment, which is identified by the loss of AQP1, AQP4 and AQP5 expression during degeneration. The expression of numerous AQPs within the disc may highlight the importance of controlling water and solute transport in relation to maintaining NC and NP cell function. As yet the actual roles of AQPs within disc maturation and degeneration are unknown; warranting further investigation into the identification of mechanisms of regulation and functions of AQPs (determined by later chapters), to elucidate the true action of these transmembrane channel proteins on the overall behaviour and health of the disc.

The osmotic regulation of AQP1 and 5 in nucleus pulposus cells
AQP1 and 5 expression is decreased during human IVD degeneration (Johnson et al., 2015). This study has identified that AQP1 and 5 are upregulated by hyperosmolality, mimicking the healthy NP, and may explain why expression is decreased during degeneration, when the osmolality is decreased. This suggests AQP1 and 5 may be part of the mechanisms that allow NP cells to adapt to their hyperosmotic environment. TonEBP has already been established as an integral part of those mechanisms; as the hyperosmotic upregulation of AQP1 and 5 is dependent on TonEBP, this implies they both participate in the osmoadaptation process and matrix synthesis. During IVD degeneration TonEBP function is uncoupled from the altered osmolality and catabolic genes are upregulated instead of the classical osmotic response genes, which may also include AQP1 and 5. As expression of these genes (and AQP1 and 5) is reduced, NP cells can no longer adapt to the altered environment and degeneration is exacerbated Therefore, AQPs may play a role in adapting NP cells to their environment and maintaining the function of NP tissue.

The regulation of cell volume and water permeability in nucleus pulposus cells
Identifying how NP cells adapt to their osmotically challenging environment is key to understanding their fundamental functions, which in turn enables their adaptation. The activity of AQP4 and TRPV4, an osmotic and mechano-sensitive Ca 2+ channel, were identified by this thesis to contribute to the fundamental functions of NP cells when physiological extracellular osmolality levels were changed. AQP4 and TRPV4 function may be linked in NP cells in order to control the rate of cell volume regulation, water permeability and Ca 2+ influx, which may trigger downstream mechanisms enabling the adaptation of NP cells to their osmotic environment.
During IVD degeneration, this thesis identified that AQP4 expression was decreased; as a result, NP cells will not be able to respond to the hypo-osmotic environment due to impaired cell volume regulation and Ca 2+ influx. Due to the lack of osmotic response, downstream mechanisms may not be employed by NP cells to ensure their survival and function in the increasingly degenerate environment. Further study is warranted to determine what impact AQP4 and TRPV4 expression and function have on potential downstream mechanisms such as RVD, cell survival and matrix synthesis.

Regulation of AQP expression by physiological conditions within the intervertebral disc
AQPs within NP cells may be regulated by many environmental cues within NP cells, such as cytokines and pH, along with alterations in osmolality. Yet, to date the true regulation and function of AQPs within the IVD is relatively unknown.
Cytokines along with combined physiological pH and O2 concentration regulated AQP expression in NP cells indicating that multiple conditions control AQP expression in the IVD, and that multiple conditions may work in tandem to finely tune AQP expression. However, AQP regulation by healthy and degenerate conditions was unclear, and other factors that contribute to the IVD microenvironment, such as nutrition and mechanical loading, were not investigated. The in vivo conditions of the IVD must be replicated very specifically to gather the information required to unfold the potential roles AQPs, and all other genes/proteins, may play during IVD physiology.

AQP membrane trafficking
The regulation of AQP expression at gene and protein level in NP cells has been investigated and linked to the potential roles AQPs play within the IVD, in this thesis. However, the function of AQPs as channels relies on their insertion into the cell membrane; which has been shown to be regulated by cellular trafficking mechanisms, such as protein kinase phosphorylation, calcium signalling and cytoskeletal rearrangement (Section 1.6).
In order to investigate if AQP (particularly 1, 4, and 5) membrane expression is regulated by physiological hyperosmolality in NP cells, the use of cell surface biotinylation can be used. This ELISA-based method biotinylates cell surface proteins after short (minutes) or longer-term (hours) osmotic treatments, and then immobilises them to avidin-coated wells. Proteins of interest can then be probed using antibodies and HRP-substrate absorbance changes detected to determine AQP 237 abundance at the cell membrane (Kitchen, P. et al., 2015). The mechanisms that control AQP trafficking can also be determined by introducing protein kinase inhibitors or transfection of AQP phosphorylation mutants (Kuwahara et al., 1995;Conner et al., 2010Conner et al., , 2012Kitchen, P. et al., 2015), calcium inhibitors such as BAPTA-AM (Kitchen, P. et al., 2015), and microtubule and actin inhibitors (Nicchia et al., 2008;Conner et al., 2010). If the trafficking of AQPs was determined, this would identify another level of functional regulation and control of AQPs within NP cells that investigations which this thesis has not elucidated.

AQP contribution to NP cell function
This thesis determined that AQP expression is altered during IVD development and degeneration, and that physiological conditions regulated their expression and function. However, it was not determined whether changes in expression were causes or consequences of the changing physiology of the IVD and what changes in AQP expression actually meant for the function of NP cells.
Performing AQP knockdown on NP cells by using siRNA or lentiviral vectors and transfection of AQP over-expression plasmids and determining the effects of AQP deletion and over-expression on matrix synthesis and cytokine, ADAMTS and MMP expression, will give an indication on how AQP expression (and changes during degeneration) may be linked to the functions of NP cells.
The effects of AQP3 expression on NP cell function has previously been investigated (Xie et al., 2016). It was determined that siRNA knockdown of AQP3 in NP cells was detrimental for matrix synthesis and proliferation and ADAMTS 238 expression was upregulated (Xie et al., 2016). This investigation shows promise that AQPs do contribute to the overall functions of NP cells and the integrity of the IVD, but as AQP3 expression is not altered during human IVD degeneration (Chapter 3), it remains to be determined what changes in AQP1, 2, 4, 5 and 7 (which showed altered expression during degeneration (Chapter 2)) actually correspond to with regards to would be beneficial to use NP cells extracted from tissue and used at a low passage, such as the NP cells utilised in this thesis.

AQP regulation by mechanical loading
Many environmental factors govern the function of NP cells. This thesis identified that physiological conditions regulate AQP expression and function in NP cells, however, not all physiological conditions were considered. Mechanical loading of the IVD, and how NP cells have adapted to this, is an essential part of how NP cells function. Yet, to date no studies have investigated how AQPs may contribute to this adaptation.
The regulation of AQP expression by mechanical loading in healthy and degenerate NP tissue explants could be investigated by using mechanical testing equipment such as the ElectroForce BioDynamic 5200 (TA Instruments) mechanical stimulation bioreactor. This system enables axial compression of tissue to simulate the physiological loading of the IVD on ex vivo NP tissue explants, whilst media is perfused through a bioreactor chamber (also allowing the nutrient diffusion and osmolality to be controlled at the same time), to determine how this effects AQP expression. The potential underlying mechanisms that control AQP expression under load, could be investigated using Flexcell (Burlington, NC, USA) culture plates, for example limited studies have previously shown that AQP function may be linked to integrin-linked kinase and focal adhesion kinase (Cano-Peñalver et al., 2014;Meng et al., 2014;Mamuya et al., 2016;Hatem-Vaquero et al., 2017). This technology allows the application of loading, such as tension and compression, to cells in monolayer which will enable the use of function-modulating compounds to determine mechanisms that control AQP expression. Such experiments may be able to determine how AQPs contribute to NP cell adaptation to mechanical loading and how they may be involved in altered mechanotransduction during IVD degeneration (Le ).

Concluding remarks
The anatomy of the IVD and age-related changes has been well documented over the past few centuries. However, LBP still costs the UK £12 billion per year in the 21 st century, with 40% of cases attributed to IVD degeneration. Much of the physiology contributing to the disc during health and degeneration has been elucidated in recent years. However, despite this, current treatment options only ameliorate symptoms; there are no treatments that target the underlying cellular causes to halt or reverse IVD degeneration. There is current research interest to develop therapies to combat IVD degeneration, but whether research findings will 240 translate into clinical use remains to be seen. It is clear however, that an increased understanding of the fundamental processes that control the physiology of the IVD is of great importance to enable the development of new therapies that take into consideration how IVD cells determine the fate of the tissue.
Together, the data presented in this thesis have contributed novel findings and increased knowledge in the field of IVD biology and spine-related research.
Results have highlighted NP cells express many AQP water channels in vivo, whose expression may be altered between disc development and degeneration. AQP1 and 5 were found to be upregulated by TonEBP in hyperosmotic conditions, mimicking the healthy IVD, which may implicate them in the adaptation of NP cells to their environment, which is no longer achievable when AQP1 and 5 are decreased during degeneration. The importance of AQP4 and TRPV4 function in the way NP cells adapt to their osmotically fluxing environment has also been determined. Additionally, this PhD project has raised new avenues of research for future study, which have the potential to unravel the roles of AQPs in the fundamental cellular processes of IVD biology.
Appendix X. Example of non-linear regression curve fitting to determine rate of cell volume change in response to extracellular osmolality changes. The change in intracellular calcein F1/F0 is plotted when NP cells are exposed to altered osmolality. To determine the rate of change in calcein F1/F0, plateau followed by one-phase association (increase in F1/F0) or decay (decrease in F1/F0) curves were fitted to data using GraphPad Prism v7.03 software. The average baseline value of Y (Y0) before experimental intervention (X0) was constrained to 1; X0 was constrained to 0.05s (the first fluorescence reading after injection).