Abstract
The cerebrospinal fluid (CSF) performs key functions for the developing central nervous system and for the adult brain. It is, indeed, a complex molecular private milieu of the brain clearing a series of compounds and conveying a wealth of signal molecules. The flow of the CSF throughout the ventricular system involves two different mechanisms: the bulk flow, driven by arterio-venous pressure gradients and arterial pulsations, and the laminar flow, driven by cilia beating of ependymal cells. Disruption of normal CSF circulation and turnover contributes to the development of many diseases. This review is aimed to bring into discussion early and new evidence concerning the brain development, ependymogenesis, and the probable mechanisms by which abnormalities in the ependymogenesis program may lead to both foetal onset hydrocephalus and abnormal neurogenesis. Evidence strongly suggests that several genetic mutations and certain foreign signals all convey into a final common pathway leading to a cell junction pathology of cells lining the ventricular walls (ventricular zone, VZ). The early disruption of the VZ of the embryonic telencephalon implies the loss of neural stem cells (NSC) and abnormal neurogenesis, while the disruption of the VZ of the Sylvius aqueduct during the perinatal period results in the loss of multiciliated ependyma, aqueduct stenosis/obliteration, alteration of the laminar, and bulk flow of CSF and hydrocephalus. These findings establish the bases for the transplantation of NSC into the ventricles of foetuses developing hydrocephalus to diminish/repair the outcomes of VZ disruption.
Access provided by Autonomous University of Puebla. Download chapter PDF
Keywords
- Cerebrospinal fluid
- Congenital hydrocephalus
- Ependymogenesis
- Neurogenesis
- Neural stem cells
- Junction pathology
- Ventricular zone disruption
- Stem cells therapy
Balanced View of CSF Physiology
Aiming to a Balanced View of CSF Physiology
Several functions have been ascribed to the cerebrospinal fluid (CSF), including protection to the brain, excretion of metabolites, homeostasis of the brain chemical environment, and as a transport pathway between different brain areas [18, 80, 98, 108]. These various functions, coupled with its rapid turnover, perpetual formation, and continuous circulation and absorption have led to consider the CSF as the “third circulation,” as first referred to by Cushing [16].
For decades, the CSF was regarded as a water solution of ions and few other components, such as glucose and vitamins. The concept of waste drainage was also associated to the “physiology” of CSF. Furthermore, the functional significance of the complex structure of the ventricular and subarachnoid compartments and the multiple populations of cell types lining discrete areas of the ventricular walls were, and still are, overlooked or neglected.
When the cerebrospinal fluid-contacting neurons were discovered, the concept that the CSF could be a pathway for signal molecules started to develop. This idea was strongly substantiated by the demonstration that the choroid plexus is a true gland that, in addition to transport water and ions, it also has the capacity to transport peptides and proteins from blood to CSF and to synthesize and secrete into the CSF a series of biologically active molecules [14, 78, 98]. Although the series of peptides, proteins, and neurotransmitters detected in the CSF using different methods increased throughout three decades, it was the analysis by mass spectrometry that suddenly revealed the enormous complexity of the molecular composition of the CSF.
In recent years, the discovery of aquaporins and other water transporters, all highly selective for water molecules, has again moved the balance to the oversimplified view that CSF physiology refers almost exclusively to water exchange between brain compartments. The glymphatic concept emphasizes the transport of water and waste molecules from the brain parenchyma into subarachnoid space along perivascular pathways of the Virchow Robin spaces, overlooking the fact that this “brain parenchyma” refers to the most superficial region of the brain cortex. It is really disturbing when the physiology of CSF is only associated with the movement of water through the different brain compartment, what leads several authors to talk about CSF secretion when in actuality they are only referring to water transport, completely disregarding the rich heterogeneity of the ventricular walls (circumventricular organs included) and the wealth of signal molecules that use the CSF as a pathway.
The CSF as a Pathway for a Cross Talk Between Different Periventricular Regions
CSF proteomics is showing a wealth of over 200 proteins [113]. A long series of peptides and neurotransmitters are also present in the CSF. Some of these compounds move by bulk flow from the interstitial fluid of brain parenchyma, many are secreted by neurons, glia, and ependyma into the CSF, others are transported by specific transport systems from blood to ventricular CSF (choroid plexus) while a few of them originate from cells present in the CSF. For many of these compounds, CSF levels bear hardly any relationship to peripheral levels in the blood [98].
The long series of biologically active proteins, peptides, and neurotransmitters present in the CSF reach this fluid through different mechanisms: (1) Neurotransmitters and their metabolites reach the CSF via the bulk flow of parenchymal fluid. (2) Regulated secretion into the CSF of biologically active compounds by the circumventricular organs (subcommissural organ, pineal gland, choroid plexuses, and median eminence), such as SCO-spondin, basic fibroblast growth factor, melatonin, transthyretin, transthyretin-T4 complex, transthyretin-T3 complex, nerve growth factor (NGF), transforming growth factor-β (TGFβ), vascular endothelial growth factor (VEGF), transferrin and vasopressin [28, 42, 43, 81]. (3) Selective and circadian regulated secretion by CSF-contacting neurons of serotonin and neuropeptides such vasopressin, oxytocin, and somatostatin [80, 99, 100]. (4) Transport of peripheral hormones through the choroid plexus. Most of the transported hormones, such as leptin, prolactin, and thyroxin, have specific targets, mostly the hypothalamus [14, 81]. The concentrations of these neuroactive compounds vary between locations, suggesting they are important for the changes in brain activity that underlie different brain states [98].
Furthermore, a series of findings indicate that cells forming the ventricular walls release into the CSF microvesicles containing signalling and intracellular proteins [13, 22, 32, 55, 96]. Harrington et al. [32] suggested that this bulk flow of nanostructures generates a dispersed signal delivery, of longer duration.
Thus, the early view that the CSF is a medium carrying brain-borne and blood-borne signals to distant targets within the brain [80] has largely been supported by numerous investigations [42, 45, 81, 108]. Worth mentioning here is the much neglected system of CSF-contacting neurons most likely playing receptive functions sensing CSF composition. Most of these neurons are bipolar with the dendritic process reaching the CSF and endowed with a 9 + 0 single cilium [100].
In brief, a good body of evidence is revealing that the dynamic and molecular composition of the CSF and, consequently, the CSF physiology is much more complex and fascinating than the simplistic view held for decades. Signal molecules, either specifically transported from blood to CSF or secreted into the CSF by a series of periventricular structures, use the CSF to reach their targets in the brain. This allows a cross talk between brain regions located beyond the blood-brain-barrier, thus keeping the brain milieu private [29, 98].
Changing of CSF Composition as It Moves Through the Ventricular System
The ventricular CSF changes its molecular composition as it unidirectionally moves through the various ventricular and subarachnoid compartments (Fig. 1.1a). The choroid plexus of the lateral ventricles, the interstitial fluid of the parenchyma surrounding these ventricles , and axon endings secreting into these cavities are the source of molecules forming this “first” fluid. At the third ventricle, new compounds are added to the CSF by hypothalamic neurons, the pineal gland, and the local choroid plexus [42, 69, 80]. When entering the Sylvius aqueduct (SA), the CSF is enriched by the secretion of the subcommissural organ [101] (Fig. 1.1a). Consequently, the CSF of the fourth ventricle is different as compared to that of the lateral ventricles [113]. This partially explains the different protein composition between the CSF collected from the lateral ventricles and that obtained from a subarachnoid compartment [101].
Furthermore, at the interphase brain cortex/subarachnoid space there is a bidirectional flow of CSF and interstitial fluid along the large paravascular spaces that surround the penetrating arteries and the draining veins (Fig. 1.1a). Since water movement along this pathway is mediated by astroglial aquaporin-4 (AQP4) water channels, this paravascular pathway has been termed “glymphatic system” [35, 36]. This pathway facilitates efficient clearance of interstitial solutes, and its failure may lead to neurodegeneration [37].
Multiciliated Ependyma and CSF Flow
The mechanisms responsible for the CSF circulation are not fully understood. The following factors do play a role: (1) the hydrostatic difference between the production and drainage sites; (2) the pulsations of the cerebral arterial tree; (3) the directional beating of ependymal cilia [18, 106, 107]. The relative contribution of each of these forces is still controversial.
The flow of the CSF throughout the ventricular system involves two different mechanisms, the bulk flow and the laminar flow. Bulk flow is driven by arterio-venous pressure gradients and arterial pulsations. The laminar flow occurs in a thin layer along the walls in a variety of directions [98]. It has been shown that cilia beating is responsible for the laminar flow of CSF, whereas its role in the bulk of CSF taking place in the core of the ventricular cavities is probably insignificant [15, 59, 66, 106]. The cilia beating of the ependyma of the lateral ventricles generate currents as far as 200 μm away from the surface [66] (Fig. 1.1b–d). In the frog brain, about 75% of the CSF within the ventricles is mixed as a result of ciliary activity [66]. Ciliary currents adjacent to the ependyma have been observed in rats, dogs, and humans [109].
The ciliary beating is supported by a sialic acid-induced hydration mantle on the ependymal surface [98]. The frequency of cilia beating is stimulated by the activation of serotonin receptors [68]. Serotonin is released by axon terminals of the suprapendymal serotonergic plexus originated in raphe nuclei [12, 68, 102].
Using computational fluid dynamics, the relative impact of macroscale (choroid plexus pulsation and ventricular wall motion) and microscale (beating of cilia) effects on near-wall CSF dynamics has been investigated [95]. This study revealed a marked effect of the cilia on the near-wall dynamics and directionality but not on the bulk flow. Conversely, the bulk flow alone does not produce any notable directionality of the flow near or on the surface of the lateral ventricles. The authors concluded that in the lateral ventricles, near-wall CSF dynamics is dominated by ependymal cilia action [95]. This confirms early observations that the ciliary movement plays a key role in the maintenance of an adequate CSF flow [31, 109, 111]. The role of the multiciliated ependyma in CSF dynamics is strongly supported by the demonstration that primary cilia dyskinesia, a syndrome that impairs ciliary activity, leads to the development of hydrocephalus [2, 17, 27, 34, 50, 51, 97]. Shimizu and Koto [93] have suggested that immotility of cilia is of particular importance in narrow canals, such as the Sylvius aqueduct, for the development of hydrocephalus.
Human Ependymogenesis
The Wall of the Lateral Ventricles of the Human Developing Brain Is a Complex and Dynamic Mosaic
Very little, if any, attention has been paid to the complexity of the cell organization of different domains of the ventricular walls. To make it even more complex, such a mosaic undergoes changes during foetal development. This dynamic complexity of the ventricular walls partially resembles that of the ventricular walls of the rodent developing brain. During the last few day of foetal life, the medial wall of the lateral ventricles of the rat is already lined by multiciliated ependyma while the lateral and dorsal walls continue formed by neural stem cells (NSC) [29]. In human embryos, there is also an early division of labour between the medial and the latero-dorsal walls of the lateral ventricles [57]. Such a partition of labour seems an efficient design; while the latter is permanently involved in neurogenesis, the medial wall is progressively engaged in the flow of CSF. Indeed, coupled multiciliated ependymal cells generate the laminar flow of CSF [59, 66] that is essential for the CSF flow along the ventricular system (see above).
In the human developing telencephalon, ependymogenesis starts about the 18th gestational week (GW) (Fig. 1.2a) in the medial wall of the lateral ventricles and progressively continues through the lateral and then to the dorsal walls [57, 83]. Thus, in young foetuses, the lateral wall of the ventricle is fully involved in neurogenesis while the medial wall starts to change its role, from neurogenesis to ependymogenesis. In pre-term foetuses, while the medial wall is fully lined by multiciliated mature ependyma, the lateral wall has mixed populations of cells suggestive of neurogenesis and ependymogenesis.
Cell Types of the VZ. Evidence for an Ependymogenesis Program
According to Rakic [76], the human telencephalic proliferative zone contains considerably complex progenitor cell groups that change during the course of development. In young embryos (5–6 GW) the VZ contains a mixed population of cells; most of them express neural stem markers only (GFAP, GLAST) while others, in addition, also express neuronal markers (βIII-tubulin, MAP-2) indicating their multipotential capacity [114]. Later in development (10–22 GW), vimentin +, GFAP+ cells displaying a long basal process persisted. Proliferation of GFAP+ cells of the ventricular zone (VZ) occurs until the 23rd GW, coinciding with the formation of the ependyma [114]. According to Gould et al. [26], in early pregnancy, the VZ is formed by GFAP+ radial glia/neural stem cells, whereas in late pregnancy, the VZ is formed by GFAP+ ependymal cells with a short basal process,
By use of several markers, Sarnat [85, 86, 88] has followed in human foetuses the temporal and spatial differentiation of cells lining the ventricular walls. In these studies, Sarnat has regarded as ependyma all cells lining the foetal ventricular system. Using this criterion, he concluded that the expression of GFAP and vimentin in foetal ependymal cells follows a regional and temporal distribution [85, 86], with the ependyma of the roof and floor plates being the first to differentiate. During several gestational weeks, GFAP is co-expressed with vimentin in most foetal ependymal cells. At birth, only scattered ependymal cells of the lateral ventricles still express GFAP, and it disappears entirely within the first few weeks of postnatal life [88].
The true process of ependymogenesis in the human remains largely unknown due, to a great extent, to limitations to obtain samples of the ventricular walls from systematically selected regions and selected gestational ages, and to process these samples for different methods. A recent investigation has provided some new evidence on ependymogenesis [57]. Based on the immunoreactivity to GFAP, AQP4, βIV-tubulin, and βIII-tubulin, their morphology (basal process, one or multiple cilia) and their spatial and temporal distribution, we have distinguished seven cell types in the VZ of the lateral ventricles of human foetuses [57, 83]. Type 1 cells with a long radial process, expressing AQP4 in the plasma membrane domain and GFAP throughout the cytoplasm, displaying a single cilium, is the main cell type in the VZ of young foetuses and most likely correspond to NSC (Fig. 1.2b). Type 2 cells are identical to type 1 cells but also express βIV-tubulin, a well-known marker of multiciliated ependyma, suggesting they correspond to NSC that have started to differentiate into ependymal cells (Fig. 1.2c). Cells type 3 through 6 would reflect progressive stages of ependymal differentiation ending in the differentiated multiciliated, βIV-tubulin+, ependyma (type 7) present during the last trimester of foetal life and throughout adulthood (Fig. 1.2d).
Hydrocephalus
A Concept
Foetal-onset hydrocephalus is a heterogeneous disease. Genetic and environmental factors, such as vitamin B or folic acid deficiency [40], viral infection of ependyma [44], and prematurity-related germinal matrix and intraventricular haemorrhage [7], contribute to its occurrence.
Numerous investigations in humans and mutant animals have substantiated the view that hydrocephalus is not only a disorder of CSF dynamics but also a brain disorder, and that derivative surgery does not resolve most aspects of the disease [46]. Actually, 80–90% of the neurological impairment of neonates with foetal onset hydrocephalus is not reversed by derivative surgery. How can we explain the inborn and, so far, irreparable neurological impairment of children born with hydrocephalus? In 2001, Miyan and his co-workers asked a key question [60]: “Humanity lost: the cost of cortical maldevelopment in hydrocephalus. Is there light ahead?” Although this appealing question has not been responded, there is some light in the horizon. A strong body of evidence indicates that the common past of hydrocephalus and brain maldevelopment starts early in the embryonic life with the disruption of the ventricular (VZ) and subventricular (SVZ) zone. However, the nature, mechanisms, and extent of the brain impairment linked to hydrocephalus are far from been fully unfolded. Certainly, a better treatment of hydrocephalus and the associated neurological impairment will come from a better understanding of the biological basis of the brain abnormalities in hydrocephalus [19, 105]. This view may represent one of the “lost highways” in hydrocephalus research, as described by Jones and Klinge [46].
To have clarity of the timetable of neurogenesis and ependymogenesis in normal rodents and humans seems essential for a better understanding of the early events occurring in foetal onset hydrocephalus.
Prenatal Neurogenesis. Timetable of Neural Proliferation and Migration, Gliogenesis, and Ependymogenesis
Virtually, all cells of the developing mammalian brain are produced in two germinal zones that form the ventricular walls, the VZ and the SVZ [6, 8, 25, 58, 39, 53]. The VZ is a pseudostratified neuroepithelium that contains multipotent radial glia/stem cells, hereafter called neural stem cells (NSC). NSC line the ventricular lumen and through a long basal process reach the pial surface. A landmark of NSC is their primary cilia that project to the ventricle and are bathed by the foetal CSF [47, 64]. During a fixed period of brain development, NSC divide asymmetrically, with one daughter cell remaining as a NSC and the other becoming a neural progenitor cell (NPC). NPC proliferate and cluster underneath the VZ, forming the so-called SVZ. NPC differentiate into neuroblasts that start migration using the basal process of NSC as scaffold. In the human, the bulk of neural proliferation and neuroblast migration occurs at a rather short period, between GW 12 and 18 (Fig. 1.2a).
Gliogenesis starts at about the 15th GW and continues for several months after birth. Ependymal cell differentiation starts at about the 18th GW and is completed after birth (Fig. 1.2a) [83, 85, 86].
Over the years, based on our own and other investigators’ evidence, we have progressively come to the view that a disruption of the VZ and SVZ, in most cases due to genetic defects, triggers onset of congenital hydrocephalus and abnormal neurogenesis (Fig. 1.2a, c). We will discuss this evidence below.
Brain Damage Versus Brain Defects
A distinction must be made between (1) brain maldevelopment due to a primary pathology of the VZ that precedes or accompanies onset of hydrocephalus and (2) brain damage caused by hydrocephalus. The former occurs during development, and consequently neonates are born with a neurological deficit. Brain damage is mainly a postnatal acquired defect, essentially caused by ventricular hypertension and abnormal CSF flow and composition.
Brain damage may be associated to regional ischemia, disruption of white matter pathways, and alteration of microenvironment of neural cells [19, 20]. Derivative surgery, the almost exclusive treatment of hydrocephalus today, is aimed to prevent or diminish brain damage. It is clear that hydrocephalic patients improve clinically after surgery due to correction of intracranial pressure, improvement in white matter blood flow [19], and probably to resumption of the clearance role of CSF. However, derivative surgery does not reverse the inborn brain defects. This has led a study group on hydrocephalus to conclude that “Fifty years after the introduction of shunts for the treatment of hydrocephalus, we must acknowledge that the shunt is not a cure for hydrocephalus” [5].
Ventricular Zone Disruption
A Concept and Definitions
For clarity purposes, we shall define the terms used in the present chapter to refer to the ventricular zone. At stages of development when the VZ is mostly formed by neural stem cells (NSC), the acronym VZ will be used. When the VZ is mostly or exclusively formed by multiciliated ependymal cells, the term “ependyma” will be used. The terms “denudation,” “disruption,” or “loss” will be alternatively used to refer to the disassembling, disorganization, or loss of the VZ cells [81]. A solid body of evidence indicates that radial glial cells are neural stem cells. Throughout the present text, we shall use the term neural stem cells, and its acronym NSC, to refer to the cells forming the embryonic ventricular zone, characterized by a long basal process, a single 9 + 0 cilium projecting to the ventricle and by expressing certain markers such a nestin [57].
In mutant animals, the disruption of the VZ follows a program that has temporal and spatial patterns, progressing as a “tsunami” wave running from caudal to rostral regions of the developing ventricular system, leaving behind a severe damage (Fig. 1.2e) [29, 41, 74, 81, 103]. A similar process of VZ disruption occurs in human hydrocephalic foetuses [21, 29, 82, 83, 94]. Since the VZ disruption is a continuous process, starting during the embryonic life and continuing during the first postnatal week, the pathology first affects NSC, then the NSC differentiating into ependymal cells and finally the differentiated multiciliated ependyma. These three cell types have distinct phenotypes and certainly play quite different roles. What do they have in common so that the denudation wave will hit them all? Junction proteins appear to be the key to understanding this devastating phenomenon [29].
A Stormy Intracellular Traffic of Junction Proteins in NSC and Ependymal Cells Leads to Ventricular Zone Disruption
What is the molecular mechanism underlying the VZ disruption occurring in human hydrocephalic foetuses, the HTx rat and in various mutant mice developing hydrocephalus? Overall, a series of findings indicates that disruption of VZ arises from a final common pathway involving alterations of vesicle trafficking, abnormal cell junctions, and loss of VZ integrity [23, 38, 48, 52, 77]. The abnormal localization of N-cadherin and connexin 43 in NSC and ependymal cells and the formation of subependymal rosettes suggest that VZ disruption results from a defect in cell polarity and in cell–cell adhesion of VZ cells. The accumulation of N-cadherin and connexin 43 in the soon-to-detach VZ cells and their virtual absence from the plasma membrane indicate that they are synthesized by the disrupting cells but are not properly transported to the plasma membrane (Fig. 1.3a–d) [29]. The mechanism actually involved in this abnormal expression and translocation of N-cadherin is unknown. The specific disruption of N-cadherin-based junctions is enough to induce ependymal disruption. Indeed, antibodies against chicken N-cadherin injected into the CSF of chick embryos disrupt the VZ, lead to denudation of the SVZ and formation of periventricular rosettes [24]. Similarly, the use of N-cadherin antibodies or synthetic peptides harbouring a cadherin-recognition sequence triggers the detachment of ependymal cells from explants of the dorsal wall of the bovine Sylvius aqueduct [71]. The abnormal localization of connexin 43 in the NSC and ependymal cells could be associated to the faulty localization of N-cadherin. Indeed, it has been reported that gap junction proteins are delivered to the plasma membrane at adherent junction sites [90].
In mutant mice, several gene mutations leading to abnormal trafficking of junction proteins and resulting in VZ disruption have been reported [11, 38, 41, 48, 52, 54, 91]. The nature of the genetic defect in hydrocephalic patients [21, 72, 94] is unknown. It may be postulated that they all carry a defect at one or another point of the pathways assembling adherent and gap junctions.
Nongenetic mechanisms leading to VZ disruption have to be considered also [92, 112]. In fact, lysophosphatidic acid, a blood-borne factor found in intraventricular haemorrhages, binds to receptors expressed by the VZ cells resulting in abnormal N-cadherin trafficking, VZ disruption, and hydrocephalus [112]. The vascular endothelial growth factor is elevated in the CSF of patients with hydrocephalus, and when administered into the CSF of normal rats, it causes alterations of adherent junctions, ependyma disruption, and hydrocephalus [92]. Thus, the possibility that signals from the hydrocephalic CSF may contribute, or even trigger VZ disruption, has to be kept in mind. Furthermore, it should be kept in mind that foetal CSF is the internal milieu of NSC [42]. Interestingly, the CSF of hydrocephalic HTx rats has an abnormal protein composition that contribute to the abnormal neurogenesis occurring in this mutant [56, 61, 62, 101].
Temporal and Spatial Programs of VZ Disruption
The process of VZ disruption has temporal and spatial patterns. The temporal program implies that disruption starts when the VZ is formed by NSC and finishes when the VZ is formed by multiciliated ependyma. In the mean time, a progressive transition from NSC to multiciliated ependyma occurs. The spatial program discloses that disruption begins in caudal regions of the ventricular system and progresses rostrally to reach the lateral ventricles [41, 74, 103]. Each of the two programs has its own outcomes.
In the temporal program, the early VZ disruption implies the loss of NSC and abnormal neurogenesis, while the late VZ disruption results in the loss of multiciliated ependyma and alterations in the laminar flow of CSF and hydrocephalus [29, 94]. In the hyh mutant mouse, the program is turned on at E12 and turned off by the end of the second postnatal week [41, 74, 103]. In the HTx mutant rat, disruption in the telencephalon starts at E19 and finishes at the first postnatal week [29]. In hydrocephalic foetuses, disruption of the VZ in the telencephalon has been shown as early as 16 GW [21, 29].
In the spatial program, the disruption of the VZ of the SA implies aqueduct stenosis/obliteration, alteration of the laminar, and bulk flow of CSF and hydrocephalus. At variance, the disruption of the VZ of the telencephalon leads to abnormal neurogenesis [29, 83].
With the years and based on solid evidence, we have progressively come to the conclusion that foetal onset hydrocephalus and abnormal neurogenesis are two inseparable phenomena, because they are linked at the etiological level.
In the pathophysiologic programs of VZ disruption, the loss of NSC and ependyma occurs in specific regions of the SA and ventricular walls, and also at specific stages of brain development. This explains why only certain brain structures have an abnormal development, which in turn results in a specific neurological impairment.
Pathophysiology of Foetal Onset Hydrocephalus
The Complex Cell Organization of the Walls of the Sylvius Aqueduct
The walls of the Sylvius aqueduct of wild-type hyh mice are formed by several populations of ependymal cells [103]. Interestingly, in mutant hydrocephalic hyh mice, some of these ependymal populations undergo proliferation, others are resistant to denudation whereas others denude [4, 74, 103]. In full-term human foetuses, the dorsal, lateral, and ventral walls of the SA three populations of ependymal cells have been described [94]. The functional significance of three ependymal populations is unclear. However, in spina bifida aperta foetuses, there seems to be an association between ependymal lineages of SA and the observed SA pathology. The ependymal cells lining the ventral wall display a normal subcellular distribution of N-cadherin and connexin 43; these cells do not detach. At variance, the ependymal cells of the lateral SA walls display an abnormal intracellular location of junction proteins and are likely to undergo denudation. The formation of large rosettes is mostly associated to this ependyma [94].
Ventricular Zone Disruption in the Sylvius Aqueduct, Aqueduct Stenosis/Obliteration, and Noncommunicating Hydrocephalus
In the hyh mouse, a programmed disruption of the VZ of the ventral wall of the SA starts early in foetal life (E12.5) and precedes the onset of a moderate communicating hydrocephalus. The loss of the ependyma of the dorsal wall of the SA occurring shortly after birth leads to fusion of the denuded ventral and dorsal walls of SA, resulting in aqueduct obliteration and severe hydrocephalus (Fig. 1.3e–e′′′) [41, 74, 103]. The phenomenon of VZ denudation associated with the onset of hydrocephalus has also been found in other mutant mice [38, 48, 52, 65, 77].
In human hydrocephalic foetuses, ependymal denudation of SA precedes and probably triggers the onset of hydrocephalus [21, 72, 94]. It can be postulated, on solid grounds, that a primary alteration of the VZ of the aqueduct due to various genetic defects triggers the onset of congenital hydrocephalus.
Ventricular Zone Disruption in the Sylvius Aqueduct, Loss of Multiciliated Ependyma and Communicating Hydrocephalus
In full-term human foetuses and in the perinatal period of mice the SA is mostly lined by multiciliated ependymal cells [94, 103]. The disruption occurring in this period in hydrocephalic humans and mutant mice implies the loss of multiciliated ependyma. Prior to denudation, the abnormal ependymal cells display abnormalities in the amount and subcellular distribution of N-cadherin and connexin 43 (Fig. 1.3) [94]. Since connexin 43 and N-cadherin co-assemble during their traffic to the plasma membrane [104], the abnormal formation of adherent junctions would also result in abnormal gap junctions. Thus, defects of adherent junctions between ependymal cells in hydrocephalic foetuses could alter gap junction-dependent ependymal physiology prior to, or in the absence of, ependymal disruption. An alteration of the CSF laminar flow through the SA of human hydrocephalic foetuses could be envisaged, even if denudation is confined to small areas of the aqueduct wall and hydrocephalus courses with a patent aqueduct (Fig. 1.3a, b). This could be part of the mechanism resulting in a communicating hydrocephalus.
At Late Gestational Stages, the Disruption in the Ventricular Zone of the Telencephalon Leads to the Loss of Multiciliated Cells and Likely Alterations in the Laminar CSF Flow
The disruption wave starting in the fourth ventricle, after a few days, reaches the telencephalon; then it continues along the walls of the lateral ventricles following a fixed route but avoiding certain discrete regions that are disruption resistant. This phenomenon occurs in certain mutant animals [41, 103] and part or most of it also occurs in human hydrocephalic foetuses [21, 29] and in premature hydrocephalic foetuses with intraventricular haemorrhage [57].
Ciliary beating of ependymal cells is responsible, at least in part, for the laminar flow of CSF occurring on the ventricular surface (see above). Long ago, Worthington and Cathcart [109] concluded that in humans, small areas of ependymal injury and ciliary destruction may affect CSF flow far beyond the region of local damage. During the third trimester of gestation, VZ disruption occurring in hydrocephalic foetuses and in cases with posthaemorrhagic hydrocephalus leaves large areas of the ventricular walls denuded [21, 29, 57]. It seems likely that these local disturbances may impair laminar CSF flow and contribute to the development of hydrocephalus.
Abnormal Neurogenesis Linked to Foetal Onset Hydrocephalus
Disruption of the Ventricular Zone of the Telencephalon Is Associated to Abnormal Neurogenesis
In human hydrocephalic foetuses [21, 29], premature infants with posthaemorrhagic hydrocephalus [57], the HTx rat [29] and the hyh mouse [23], the VZ disruption results in two neuropathological events: formation of periventricular heterotopias and translocation of NSC/NPC to the CSF (Fig. 1.4a–d).
At regions of disruption where NSC have been lost, the neuroblasts generated in the SVZ no longer have the structural scaffold to migrate and consequently accumulate in periventricular areas forming periventricular heterotopias. In human hydrocephalic foetuses, periventricular heterotopias have been found in young (21 GW) and full-term (40 GW) foetuses, indicating that they were formed early in development and had remained in situ until the end of foetal life and, probably, after birth (Fig. 1.4a, b). Interestingly, a 2-month-old child with a disrupted VZ carried periventricular heterotopias [23]. Humans with disruption in the VZ of the telencephalon carry periventricular heterotopias primarily composed of later-born neurons [23]. Periventricular heterotopias behave as epileptogenic foci [30]. This may explain why 6–30% of hydrocephalic children, including the present case, develop epilepsy that is not solved by CSF drainage surgery [72, 89].
The Cerebrospinal Fluid Is the Main Fate of the Disrupting NSC/NPC
In hydrocephalic human foetuses [21, 29] and premature infants with posthaemorrhagic hydrocephalus [49], NSC/NPC reach the ventricle at sites of VZ disruption and can be collected from the CSF. Furthermore, cells collected from CSF of two SBA foetuses develop into neurospheres [83].
In the hydrocephalic HTx rat, proliferative NPC from the SVZ reach the ventricle through the sites of VZ disruption and can be collected from the CSF. Nestin+ NSC from the VZ also appear to reach the CSF (Fig. 1.1c, d). When processed for the neurosphere assay, the cells collected from CSF proliferate and become assembled again through adherent junctions to form neurospheres. After 2 days in culture, the neurospheres start to express an adherent junction pathology (Fig. 1.4e, f) and become disrupted, mirroring the pathology of NSC in the VZ of the living hyHTx. This finding strongly indicates that a genetic defect and not epigenetic factors, such as increased CSF pressure or changes of CSF composition, underlies the disruption phenomenon.
The findings discussed indicate that NSC and NPC collected from the CSF of hydrocephalic patients can be used to investigate cell and molecular alterations underlying the disease. Thus, the inability to obtain human brain biopsies for diagnostic and research reasons may be overcome.
In brief, the evidence discussed in the present chapter identifies a new mechanism underlying the abnormal neurogenesis associated to foetal-onset hydrocephalus (Fig. 1.4g). A cell junction pathology of NSC is associated to the disruption of the VZ, the formation of periventricular heterotopias, and the abnormal translocation of NSC and NPC to the foetal CSF. The outcomes of these abnormalities continue to the end of foetal life and most likely during postnatal life. These abnormalities could explain the neurological impairments, such as epilepsy, of children born with hydrocephalus. Furthermore, the new evidence also provides the basis for the use of the neurosphere assay for diagnosis and cell therapy [29, 83]. We agree with Del Bigio [19] and Williams et al. [105] that “better treatment of hydrocephalus and the associated neurological impairment will come from a better understanding of the biological basis of the brain abnormalities in hydrocephalus.”
Repair Mechanisms of the Disrupted Ventricular Zone
In the hyh mouse, the pathophysiologic program leading to hydrocephalus includes a repairing stage in which the missing VZ is replaced by a layer of astrocytes forming a new interface between the CSF and the brain parenchyma (Fig. 1.5a–c) [74, 79, 103]. This unique astrocyte layer prevents the NPC still present in the SVZ from being displaced into the ventricle. This response occurs shortly after VZ disruption and takes weeks to complete [74, 79]. This phenomenon has also been described in the hydrocephalic HTx rat [29] and the human hydrocephalic foetuses [21, 29, 57, 87, 94].
The astrocytes re-populating the denuded areas are different from astrocytes of the normal brain parenchyma and from reactive astrocytes found after brain injury. They share several cytological features with multiciliated ependyma and similar para-cellular and intra-cellular routes of transport of cargo molecules moving between CSF, the subependymal neuropile and the pericapillary space (Fig. 1.5a–c) [79]. How do astrocytes arriving at the denuded ventricular surface become arranged into a compact cell layer? In the hyh mice, the numerous interdigitations between the cell bodies of astrocytes and the dense network formed by their processes might explain the stability of this newly formed cell layer (Fig. 1.5b, c) [79]. What are the signals mediating this response? In hyh mice and human hydrocephalic foetuses, VZ disruption takes place at prenatal stages previous to a detectable hydrocephalus. Therefore, intraventricular pressure or expanding ventricles cannot be considered responsible for the denudation of the VZ or its repairing by astrocytes [79].
In hyh mice, the periventricular astrocyte reaction appears at stages when ventriculomegaly is starting to develop. The most robust astrocyte layer occurs in the denuded floor of the fourth ventricle, a cavity displaying a minor dilatation [79]. What are the physiopathological consequences for the brain of the assembly of a compact layer of astrocytes replacing the lost ependyma? This is an important question open to investigation. Still, there are already some clues. Astrocytes replacing the denuded ependyma have a high expression of AQP4 and a high endocytosis and transcytosis activity, suggesting they function as a new CSF–brain interphase involved in water and solute transport, contributing to re-establish some of the functions of the lost ependyma [79].
Interestingly, the disruption of the VZ that occurs in foetal life of the hydrocephalic HTx rat and the repairing astrocyte mechanisms occurring postnatally is followed by a second disruption process, this time affecting the astroglial layer. The outcome of this new disruption is the massive translocation of neurons into the ventricle [73]. This second and devastating disruption process observed in 1-month-old rats could be part of the mechanism leading to death.
Cell Therapy in the Horizon
Once establishing that foetal-onset hydrocephalus and abnormal neurogenesis are two inseparable phenomena turned on by a cell junction pathology first affecting NSC/NPC and later the multiciliated ependyma; the grafting of stem cells into hydrocephalic foetuses appears as a valid therapeutic task to repair the VZ disruption and its outcomes.
Growing evidence has shown that stem cell transplantation represents a great opportunity for the treatment of many neurological diseases. Stem cells used for transplantation into the central nervous system (CNS) include mesenchymal stem cells (MSC) [84], NSC [3, 9], and NPC [70, 110]. In most of the early investigations, the stem cells were grafted in the vicinity of the injured or altered neural tissue. However, delivery of stem cells into the CSF is emerging as an alternative, particularly for those diseases with a broad distribution in the central nervous system [3, 67, 75, 110]. A key question whether the hydrocephalic CSF would be a friendly medium to host grafted NSC has been recently solved. When neurospheres obtained from non-affected HTx rats are further cultured in the presence of CSF from hydrocephalic HTx rats, neural stem cells differentiate into neurons, astrocytes, and ependyma [33].
On-going experiments in our laboratory grafting normal neurospheres into the lateral ventricle of hydrocephalic HTx rats has shown that 48 hs after transplantation, the grafted NSC moves selectively to the area devoid of VZ, proliferate, and differentiate into patches of multiciliated ependyma; a second subpopulation move into the cerebral cortex. According to our current investigations, it seems likely that the new multiciliated ependyma formed after NSC grafting would help the laminar flow of CSF and, consequently, attenuate the hydrocephalus condition. If NSC grafting results in a functional recovery of the neurological deficit of the rats born with hydrocephalus is under research.
Toward the frontier of the bed side
The isolation and expansion of NSC of human origin are crucial for the successful development of cell therapy approaches in human brain diseases. A relevant step forward has been achieved by scientists of the Neuroscience Center of Lund (Sweden) who developed an immortal neural stem cell line and have standardized a protocol to obtain neurospheres from foetal striatum-derived neural stem [10, 63]. An additional key point to consider is the time and opportunity when NSC should be transplanted. It seems reasonable to suggest that NSC grafting should be performed shortly after the disruption process of the VZ had been turned on. In human hydrocephalic foetuses, VZ disruption starts at about the 16th GW and continues throughout the second and third trimester of pregnancy (see above). The opportunity for transplantation may be the foetal surgery performed to repair neural tube defects, such as spina bifida aperta, that is performed within a well-defined gestational period (19th–25th GW) [1]. It may be hoped that grafting of stem cells into the hydrocephalic brain would result in the repopulation of the disrupted areas of the VZ and/or the generation of a protective microenvironment to diminish/prevent the outcomes of VZ disruption. (Fig. 1.6).
Abbreviations
- AQP4:
-
Aquaporin 4
- CSF:
-
Cerebrospinal fluid
- GW:
-
Gestational week
- NPC:
-
Neural progenitor cell
- NSC:
-
Neural stem cells
- SA:
-
Sylvius aqueduct
- SVZ:
-
Subventricular zone
- VZ:
-
Ventricular zone
References
Adzick NS, Thom EA, Spong CY, Brock JW, Burrows PK, Johnson MP, Howell LJ, Farrell JA, Dabrowiak ME, Sutton LN, Gupta N, Tulipan NB, D’Alton ME, Farmer DL, Investigators MOMS. A randomized trial of prenatal versus postnatal repair of myelomeningocele. N Engl J Med. 2011;364:993–1000.
Afzelius BA. The immotile-cilia syndrome: a microtubule-associated defet. CRC Crit Rev Biochem. 1985;19:63–87.
Bai H, Suzuki Y, Noda T, Wu S, Kataoka K, Kitada K, Ohta M, Chou H, Ide C. Dissemination and proliferation of neural stem cells on injection into the fourth ventricle of the rat: a transplantation. J Neurosci Methods. 2003;124:181–7.
Bátiz LF, Páez P, Jiménez AJ, Rodríguez S, Wagner C, Pérez-Fígares JM, Rodríguez EM. Heterogeneous expression of hydrocephalic phenotype in the hyh mice carrying a point mutation in alpha-SNAP. Neurobiol Dis. 2006;23:152–68.
Bergsneider M, Egnor MR, Johnston M, Kranz D, Madsen JR, JP MA 2nd, Stewart C, Walker ML, Williams MA. What we don’t (but should) know about hydrocephalus. J Neurosurg. 2006;104:157–9.
Bonfanti L, Peretto P. Radial glial origin of the adult neural stem cells in the subventricular zone. Prog Neurobiol. 2007;83:24–36.
Boop FA. Posthemorrhagic hydrocephalus of prematurity. In: Cinalli C, Maixner WJ, Sainte-Rose C, editors. Pediatric hydrocephalus. Milan: Springer-Verlag; 2004.
Brazel CY, Romanko MJ, Rothstein RP, Levison SW. Roles of the mammalian subventricular zone in brain development. Prog Neurobiol. 2003;69:49–69.
Buddensiek J, Dressel A, Kowalski M, Runge U, Schroeder H, Hermann A, Kirsch M, Storch A, Sabolek M. Cerebrospinal fluid promotes survival and astroglial differentiation of adult human neural progenitor cells but inhibits proliferation and neuronal differentiation. BMC Neurosci. 2010;11:48.
Cacci E, Villa A, Parmar M, Cavallaro M, Mandahl N, Lindvall O, Martinez-Serrano A, Kokaia Z. Generation of human cortical neurons from a new immortal fetal neural stem cell line. Exp Cell Res. 2007;313:588–601.
Chae TH, Kim S, Marz KE, Hanson PI, Walsh CA. The hyh mutation uncovers roles for Snap in apical protein localization and control of neural cell fate. Nat Genet. 2004;36:264–70.
Chan-Paly V. Serotonin axons in the supra- and subependymal plexuses and in the leptomeninges; their roles in local alterations of cerebrospinal fluid and vasomotor activity. Brain Res. 1976;102:103–30.
Chiasserini D, van Weering JR, Piersma SR, Pham TV, Malekzadeh A, Teunissen CE, de Wit H, Jiménez CR. Proteomic analysis of cerebrospinal fluid extracellular vesicles: a comprehensive dataset. J Proteome. 2014;106:191–204.
Chodobski A, Szmydynger-Chodobska J. Choroid plexus: target for polypeptides and site of their synthesis. Microsc Res Tech. 2001;52:865–82.
Cifuentes M, Rodríguez S, Pérez J, Grondona JM, Rodríguez EM, Fernández-Llebrez P. Decreased cerebrospinal fluid flow through the central canal of the spinal cord of rats immunologically deprived of Reissner’s fibre. Exp Brain Res. 1994;98:431–40.
Cushing H. Studies in intracranial physiology and surgery: the third circulation, the hypophysis, the gliomas. Serie: Cameron-prize lecture. London: H. Milford, Oxford University Press; 1926.
Davis RE, Swiderski RE, Rahmouni K, Nishimura DY, Mullins RF, Agassandian K, Philp AR, Searby CC, Andrews MP, Thompson S, Berry CJ, Thedens DR, Yang B, Weiss RM, Cassell MD, Stone EM, Sheffield VC. A knockin mouse model of the Bardet-Biedl syndrome 1 M390R mutation has cilia defects, ventriculomegaly, retinopathy, and obesity. Proc Natl Acad Sci U S A. 2007;104:19422–7.
Davson H, Segal MB. Physiology of the CSF and blood–brain barriers. Boca Raton: CRC Press; 1995.
Del Bigio MR. Pathophysiologic consequences of hydrocephalus. Neurosurg Clin N Am. 2001;12:639–49.
Del Bigio MR. Neuropathology and structural changes in hydrocephalus. Dev Disabil Res Rev. 2010;16:16–22.
Domínguez-Pinos MD, Páez P, Jiménez AJ, Weil B, Arráez MA, Pérez-Fígares JM, Rodríguez EM. Ependymal denudation and alterations of the subventricular zone occur in human fetuses with a moderate communicating hydrocephalus. J Neuropathol Exp Neurol. 2005;64:595–604.
Feliciano DM, Zhang S, Nasrallah CM, Lisgo SN, Bordey A. Embryonic cerebrospinal fluid nanovesicles carry evolutionarily conserved molecules and promote neural stem cell amplification. PLoS One. 2014;9(2):e88810.
Ferland RJ, Bátiz LF, Neal J, Lian G, Bundock E, Lu J, Hsiao YC, Diamond R, Mei D, Banham AH, Brown PJ, Vanderburg CR, Joseph J, Hecht JL, Folkerth R, Guerrini R, Walsh CA, Rodríguez EM, Sheen VL. Disruption of neural progenitors along the ventricular and subventricular zones in periventricular heterotopia. Hum Mol Genet. 2009;18:497–516.
Ganzler-Odenthal SI, Redies C. Blocking N-cadherin function disrupts the epithelial structure of differentiating neural tissue in the embryonic chicken brain. J Neurosci. 1998;18:5415–25.
Götz M, Huttner WB. The cell biology of neurogenesis. Nat Rev Mol Cell Biol. 2005;6:777–88.
Gould SJ, Howard S, Papadaki L. The development of ependyma in the human fetal brain: an immunohistological and electron microscopic study. Dev Brain Res. 1990;55:255–67.
Greenstone MA, Jones RWA, Dewar A, Neville BGR, Cole PJ. Hydrocephalus and primary ciliary dyskinesia. Arch Dis Child. 1984;59:481–2.
Gross PM. Circumventricular organs and body fluids, Vol. I, II, and III. Boca Raton: CRC Press; 1987.
Guerra M, Henzi R, Ortloff A, Lichtin N, Vío K, Jimémez A, Dominguez-Pinos MD, González C, Jara MC, Hinostroza F, Rodríguez S, Jara M, Ortega E, Guerra F, Sival DA, den Dunnen WFA, Pérez-Figares JM, McAllister JP, Johanson CE, Rodríguez EM. Cell junction pathology of neural stem cells is associated with ventricular zone disruption, hydrocephalus, and abnormal neurogenesis. J Neuropathol Exp Neurol. 2015;74:653–71.
Guerrini R, Barba C. Malformations of cortical development and aberrant cortical networks: epileptogenesis and functional organization. J Clin Neurophysiol. 2010;27:372–9.
Hagenlocher C, Walentek P, M Ller C, Thumberger T, Feistel K. Ciliogenesis and cerebrospinal fluid flow in the developing Xenopus brain are regulated by foxj1. Cilia. 2013;2:12.
Harrington MG, Fonteh AN, Oborina E, Liao P, Cowan RP, McComb G, Chavez JN, Rush J, Biringer RG, Huhmer AF. The morphology and biochemistry of nanostructures provide evidence for synthesis and signaling functions in human cerebrospinal fluid. Cerebrospinal Fluid Res. 2009;6:10.
Henzi R, Guerra M, Vío K, González C, Herrera C, McAllister JP, Johanson C, Rodríguez EM. Neurospheres from neural stem/neural progenitor cells (NSPC) of non-hydrocephalic HTx rats produce neurons, astrocytes and multiciliated ependyma. The cerebrospinal fluid of normal and hydrocephalic rats supports such a differentiation. Cell Tissue Res. 2018;373:421–38.
Ibañez-Tallon I, Pagenstecher A, Fliegauf M, Olbrich H, Kispert A, Ketelsen UP, North A, Heintz N, Omran H. Dysfunction of axonemal dynein heavy chain Mdnah5 inhibits ependymal flow and reveals a novel mechanism for hydrocephalus formation. Hum Mol Genet. 2004;13:2133–41.
Iliff JJ, Wang M, Liao Y, Plogg BA, Peng W, Gundersen GA, Benveniste H, Vates GE, Deane R, Goldman SA, Nagelhus EA, Nedergaard M. A paravascular pathway facilitates CSF flow through the brain parenchyma and the clearance of interstitial solutes, including amyloid β. Sci Transl Med. 2012;4(147):147ra111.
Iliff JJ, Wang M, Zeppenfeld DM, Venkataraman A, Plog BA, Liao Y, Deane R, Nedergaard M. Cerebral arterial pulsation drives paravascular CSF-interstitial fluid exchange in the murine brain. J Neurosci. 2013;33:18190–9.
Iliff JJ, Chen MJ, Plog BA, Zeppenfeld DM, Soltero M, Yang L, Singh I, Deane R, Nedergaard M. Impairment of glymphatic pathway function promotes tau pathology after traumatic brain injury. J Neurosci. 2014;34:16180–93.
Imai F, Akimoto K, Koyama H, Miyata T, Ogawa M, Noguchi S, Sasaoka T, Noda T, Ohno S. Inactivation of aPKClambda results in the loss of adherens junctions in neuroepithelial cells without affecting neurogenesis in mouse neocortex. Development. 2006;133:1735–44.
Jacobsen M. Developmental neurobiology. New York: Plenum; 1991.
Jellinger G. Anatomopathology of nontumoral aqueductal stenosis. J Neurosurg Sci. 1986;30:1Y16.
Jiménez AJ, Tomé M, Páez P, Wagner C, Rodríguez S, Fernández-Llebrez P, Rodríguez EM, Pérez-Fígares JM. A programmed ependymal denudation precedes congenital hydrocephalus in the hyh mutant mouse. J Neuropathol Exp Neurol. 2001;60:1105–19.
Johanson CE, Duncan JA 3rd, Klinge PM, Brinker T, Stopa EG, Silverberg GD. Multiplicity of cerebrospinal fluid functions: new challenges in health and disease. Cerebrospinal Fluid Res. 2008;5:10.
Johansson PA. The choroid plexuses and their impact on developmental neurogenesis. Front Neurosci. 2014;8:340.
Johnson RT, Johnson KP, Edmonds CJ. Virus-induced hydrocephalus: development of aqueductal stenosis in hamsters after mumps infection. Science. 1967;157:1066Y67.
Johnson AK, Gross PM. Sensory circumventricular organs and brain homeostatic pathways. FASEB J. 1993;7:678–86.
Jones HC, Klinge PM. Hydrocephalus, 17–20th September, Hannover Germany: a conference report. Cerebrospinal Fluid Res. 2008;5:19.
Kazanis I, Lathia J, Moss L, ffrench-Constant C. The neural stem cell microenvironment. StemBook [Internet]. Cambridge, MA: Harvard Stem Cell Institute; 2008.
Klezovitch O, Fernandez TE, Tapscott SJ, Vasioukhin V. Loss of cell polarity causes severe brain dysplasia in Lgl1 knockout mice. Genes Dev. 2004;18:559–71.
Krueger RC, Wu H, Zandian M, Daniel-PouRM KP, Yu JS, Sun YE. Neural progenitors populate the cerebrospinal fluid of pre-term patients with hydrocephalus. J Pediatr. 2006;148:337–40.
Lechtreck KF, Delmotte P, Robinson ML, Sanderson MJ, Witman GB. Mutations in Hydin impair ciliary motility in mice. J Cell Biol. 2008;180:633–43.
Lee L. Riding the wave of ependymal cilia: genetic susceptibility to hydrocephalus in primary ciliary dyskinesia. J Neurosci Res. 2013;91:1117–32.
Ma X, Bao J, Adelstein RS. Loss of cell adhesion causes hydrocephalus in nonmuscle myosin II-B-ablated and mutated mice. Mol Biol Cell. 2007;18:2305–12.
Malatesta P, Appolloni I, Calzolari F. Radial glia and neural stem cells. Cell Tissue Res. 2008;331:165–78.
Markham NO, Doll CA, Dohn MR, Miller RK, Yu H, Coffey RJ, McCrea PD, Gamse JT, Reynolds AB. DIPA-family coiled-coils bind conserved isoform-specific head domain of p120-catenin family: potential roles in hydrocephalus and heterotopia. Mol Biol Cell. 2014;25:2592–603.
Marzesco AM, Janich P, Wilsch-Bräuninger M, Dubreuil V, Langenfeld K, Corbeil D, Huttner WB. Release of extracellular membrane particles carrying the stem cell marker prominin-1 (CD133) from neural progenitors and other epithelial cells. J Cell Sci. 2005;118:2849–58.
Mashayekhi F, Draper CE, Bannister CM, Pourghasem M, Owen-Lynch PJ, Miyan JA. Deficient cortical development in the hydrocephalic Texas (H-Tx) rat: a role for CSF. Brain. 2002;125:1859–74.
McAllister P, Guerra M, Lc R, Jimenez AJ, Dominguez-Pinos D, Sival D, den Dunnen W, Morales DM, Schmidt RE, Rodríguez EM, Limbrick DD. Ventricular zone disruption in human neonates with intraventricular hemorrhage. J Neuropathol Exp Neurol. 2017;76(5):358–75.
Merkle FT, Alvarez-Buylla A. Neural stem cells in mammalian development. Curr Opin Cell Biol. 2006;18:704–9.
Milhorat TH. The third circulation revisited. J Neurosurg. 1975;42:628–45.
Miyan J, Sobkowiak C, Draper C. Humanity lost: the cost of cortical maldevelopment. Is there light ahead? Eur J Pediatr Surg. 2001;11(Suppl 1):S4–9.
Miyan JA, Nabiyouni M, Zendah M. Development of the brain: a vital role for cerebrospinal fluid. Can J Physiol Pharmacol. 2003;81:317–28.
Miyan JA, Zendah M, Mashayekhi F, Owen-Lynch PJ. Cerebrospinal fluid supports viability and proliferation of cortical cells in vitro, mirroring in vivo development. Cerebrospinal Fluid Res. 2006;3:2.
Monni E, Cusulin C, Cavallaro M, Lindvall O, Kokaia Z. Human fetal striatum-derived neural stem (NS) cells differentiate to mature neurons in vitro and in vivo. Curr Stem Cell Res Ther. 2014;9:338–46.
Mori T, Buffo A, Gotz M. The novel roles of glial cells revisited: the contribution of radial glia and astrocytes to neurogenesis. Curr Top Dev Biol. 2005;69:67–99.
Nechiporuk T, Fernández TE, Vasioukhin V. Failure of epithelial tube maintenance causes hydrocephalus and renal cysts in Dlg5-/- mice. Dev Cell. 2007;13:338–50.
Nelson DJ, Wright EM. The distribution, activity, and function of the cilia in the frog brain. J Physiol Lond. 1974;243:63–78.
Neuhuber B, Barshinger AL, Paul C, Shumsky JS, Mitsui T, Fischer I. Stem cell delivery by lumbar puncture as a therapeutic alternative to direct injection into injured spinal cord. J Neurosurg Spine. 2008;9:390–9.
Nguyen T, Chin WC, O'Brien JA, Verdugo P, Berger AJ. Intracellular pathways regulating ciliary beating of rat brain ependymal cells. J Physiol. 2001;531.(Pt 1:131–40.
Nicholson C. Signals that go with the flow. Trends Neurosci. 1999;22:143–5.
Ohta M, Suzuki Y, Noda T, Kataoka K, Chou H, Ishikawa N, Kitada M, Matsumoto N, Dezawa M, Suzuki S, Ide C. Implantation of neural stem cells via cerebrospinal fluid into the injured root. Neuroreport. 2004;15:1249–53.
Oliver C, González C, Alvial G, Flores CA, Rodríguez EM, Batiz LF. Disruption of CDH2/N-cadherin-based adherens junctions leads to apoptosis of ependymal cells and denudation of brain ventricular walls. J Neuropathol Exp Neurol. 2013;72:846–60.
Ortega E, Muñoz RI, Luza N, Guerra F, Guerra M, Vio K, Henzi R, Jaque J, Rodriguez S, McAllister JP, Rodriguez EM. The value of early and comprehensive diagnoses in a human fetus with hydrocephalus and progressive obliteration of the aqueduct of Sylvius: case report. BMC Neurol. 2016;16:45.
Ortloff A, Lichtin N, Guerra M, Vío K, Rodríguez EM. The disruption of the ventricular zone that occurs in foetal life of the hydrocephalic HTx rat is followed by a second disruption in the postnatal life. 57th annual meeting of Society of Research into Hydrocephalus and Spina Bifida, Cologne, Germany, 2013.
Páez P, Bátiz LF, Roales-Buján R, Rodríguez-Pérez LM, Rodríguez S, Jiménez AJ, Rodríguez EM, Pérez-Fígares JM. Patterned neuropathologic events occurring in hyh congenital hydrocephalic mutant mice. J Neuropathol Exp Neurol. 2007;66:1082–92.
Pluchino S, Quattrini A, Brambilla E, Gritti A, Salani G, Dina G, Galli R, Del Carro U, Amadio S, Bergami A, Furlan R, Comi G, Vescovi AL, Martino G. Injection of adult neurospheres induces recovery in a chronic model of multiple sclerosis. Nature. 2003;422:688–94.
Rakic P. Elusive radial glial cells: historical and evolutionary perspective. Glia. 2003;43:19–32.
Rasin M, Gazula V, Breunig J, Kwan KY, Johnson MB, Liu-Chen S, Li HS, Jan LY, Jan YN, Rakic P, Sestan N. Numb and Numbl are required for maintenance of cadherin-based adhesion and polarity of neural progenitors. Nat Neurosci. 2007;10:819–27.
Redzic ZB, Segal MB. The structure of the choroid plexus and the physiology of the choroid plexus epithelium. Adv Drug Deliv Rev. 2004;56:1695–716.
Roales-Buján R, Páez P, Guerra M, Rodríguez S, Vío K, Ho-Plagaro A, García-Bonilla M, Rodríguez-Pérez LM, Domínguez-Pinos MD, Rodríguez EM, Pérez-Fígares JM, Jiménez AJ. Astrocytes acquire morphological and functional characteristics of ependymal cells following disruption of ependyma in hydrocephalus. Acta Neuropathol. 2012;124:531–46.
Rodríguez EM. The cerebrospinal fluid as a pathway in neuroendocrine integration. J Endocrinol. 1976;71:407–43.
Rodríguez EM, Blázquez JL, Guerra M. The design of barriers in the hypothalamus allows the median eminence and the arcuate nucleus to enjoy private milieus: the former opens to the portal blood and the latter to the cerebrospinal fluid. Peptides. 2010;31:757–76.
Rodríguez EM, Guerra MM, Vío K, González C, Ortloff A, Bátiz LF, Rodríguez S, Jara MC, Muñoz RI, Ortega E, Jaque J, Guerra F, Sival DA, den Dunnen WF, Jiménez AJ, Domínguez-Pinos MD, Pérez-Fígares JM, McAllister JP, Johanson C. A cell junction pathology of neural stem cells leads to abnormal neurogenesis and hydrocephalus. Biol Res. 2012;45:231–42.
Rodríguez EM, Guerra M. Neural stem cells and fetal onset hydrocephalus. Pediatr Neurosurg. 2017; https://doi.org/10.1159/000453074.
Satake K, Lou J, Lenke LG. Migration of mesenchymal stem cells through cerebrospinal fl uid into injured spinal cord tissue. Spine. 2004;29:1971–9.
Sarnat HB. Role of human fetal ependyma. Pediatr Neurol. 1992a;8:163–78.
Sarnat HB. Regional differentiation of the human fetal ependyma: immunocytochemical markers. J Neuropathol Exp Neurol. 1992b;51:58–75.
Sarnat HB. Ependymal reactions to injury. A review. J Neuropathol Exp Neurol. 1995;54:1–15.
Sarnat HB. Histochemistry and immunocytochemistry of the developing ependyma and choroid plexus. Microsc Res Tech. 1998;41:14–28.
Sato O, Yamguchi T, Kittaka M, Toyama H. Hydrocephalus and epilepsy. Childs Nerv Syst. 2001;17(1–2):76–86.
Shaw RF, Fay AJ, Puthenveedu M, et al. Microtubule plus-end-tracking proteins target gap junctions directly from the cell interior to adherens junctions. Cell. 2007;128:547–60.
Shibasaki T, Tokunaga A, Sakamoto R, Sagara H, Noguchi S, Sasaoka T, Yoshida N. PTB deficiency causes the loss of adherens junctions in the dorsal telencephalon and leads to lethal hydrocephalus. Cereb Cortex. 2013;23:1824–35.
Shim JW, Sandlund J, Han CH, Hameed MQ, Connors S, Klagsbrun M, Madsen JR, Irwin N. VEGF, which is elevated in the CSF of patients with hydrocephalus, causes ventriculomegaly and ependymal changes in rats. Exp Neurol. 2013;247:703–9.
Shimizu A, Koto M. Ultrastructure and movement of the ependymal and tracheal cilia in congenitally hydrocephalic WIC-Hyd rats. Childs Nerv Syst. 1992;8:25–32.
Sival DA, Guerra M, den Dunnen WFA, Bátiz LF, Alvial G, Rodríguez EM. Neuroependymal denudation is in progress in full-term human foetal spina bifida aperta. Brain Pathol. 2011;21:163–79.
Siyahhan B, Knobloch V, de Zélicourt D, Asgari M, Schmid Daners M, Poulikakos D, Kurtcuoglu V. Flow induced by ependymal cilia dominates near-wall cerebrospinal fluid dynamics in the lateral ventricles. J R Soc Interface. 2014;11:20131189.
Street JM, Barran PE, Mackay CL, Weidt S, Balmforth C, Walsh TS, Chalmers RT, Webb DJ, Dear JW. Identification and proteomic profiling of exosomes in human cerebrospinal fluid. J Transl Med. 2012;5:10–5.
Tissir F, Qu Y, Montcouquiol M, et al. Lack of cadherins Celsr2 and Celsr3 impairs ependymal ciliogenesis, leading to fatal hydrocephalus. Nat Neurosci. 2010;13:700–7.
Veening JG, Barendregt HP. The regulation of brain states by neuroactive substances distributed via the cerebrospinal fluid; a review. Cerebrospinal Fluid Res. 2010;7:1.
Vigh-Teichmann I, Vigh B. The cerebrospinal fluid-contacting neuron: a peculiar cell type of the central nervous system. Immunocytochemical aspects. Arch Histol Cytol. 1989;52:195–207.
Vigh B, Manzano e Silva MJ, Frank CL, Vincze C, Czirok SJ, Szabó A, Lukáts A, Szél A. The system of cerebrospinal fluid-contacting neurons. Its supposed role in the nonsynaptic signal transmission of the brain. Histol Histopathol. 2004;19:607–28.
Vío K, Rodríguez S, Yulis CR, Oliver C, Rodríguez EM. The subcommissural organ of the rat secretes Reissner’s fiber glycoproteins and CSF-soluble proteins reaching the internal and external CSF compartments. Cerebrospinal Fluid Res. 2008;5:3.
Voutsinos B, Chouaf L, Mertens P, Ruiz-Flandes P, Joubert Y, Belin MF, Didier-Bazes M. Tropism of serotonergic neurons towards glial targets in the rat ependyma. Neuroscience. 1994;59:663–72.
Wagner C, Bátiz LF, Rodríguez S, Jiménez AJ, Páez P, Tomé M, Pérez-Fígares JM, Rodríguez EM. Cellular mechanisms involved in the stenosis and obliteration of the cerebral aqueduct of hyh mutant mice developing congenital hydrocephalus. J Neuropathol Exp Neurol. 2003;62:1019–40.
Wei CJ, Francis R, Xu X, Lo CW. Connexin43 associated with an N-cadherin-containing multiprotein complex is required for gap junction formation in NIH3T3 cells. J Biol Chem. 2005;280:19925–36.
Williams MA, McAllister JP, Walker ML, Kranz DA, Bergsneider M, Del Bigio MR, Fleming L, Frim DM, Gwinn K, Kestle JR, Luciano MG, Madsen JR, Oster-Granite ML, Spinella G. Priorities for hydrocephalus research: report from a National Institutes of Health-sponsored workshop. J Neurosurg. 2007;107:345–57.
Wrigh EM. Transport processes in the formation of the cerebrospinal fluid. Rev Physiol Biochem Pharmacol. 1978;83:1–34.
Wright EM. Secretion and circulation of the cerebrospinal fluid. In: Rodriguez EM, van Wimersma Greidanus TB, editors. Front Horm Res. Basel: Karger; 1981.
Wood JH. Neurobiology of cerebrospinal fluid. New York: Plenum; 1983.
Worthington WC Jr, Cathcart RS 3rd. Ciliary currents on ependymal surfaces. Ann N Y Acad Sci. 1966;130:944–50.
Wu S, Suzuki Y, Noda Y, Bai H, Kitada M, Kataoka K, Nishimura Y, Ide C. Immunohistochemical and electron microscopic study of invasion and differentiation in spinal cord lesion of neural stem cells grafted through cerebrospinal fluid in rat. J Neurosci Res. 2002;69:940–5.
Yamadori T, Nara K. The directions of ciliary beat on the wall of the lateral ventricle and the currents of the cerebrospinal fluid in the brain ventricles. Scan Electron Microsc. 1979;3:335–40.
Yung YC, Mutoh T, Lin ME, Noguchi K, Rivera RR, Choi JW, Kingsbury MA, Chun J. Lysophosphatidic acid signaling may initiate fetal hydrocephalus. Sci Transl Med. 2011;3:99ra87.
Zappaterra MD, Lisgo SN, Lindsay S, Gygi SP, Walsh CA, Ballif BA. A comparative proteomic analysis of human and rat embryonic cerebrospinal fluid. J Proteome Res. 2007;6:3537–48.
Zecevic N. Specific characteristic of radial glia in the human fetal telencephalon. Glia. 2004;48:27–35.
Author information
Authors and Affiliations
Corresponding author
Editor information
Editors and Affiliations
Rights and permissions
Copyright information
© 2019 Springer Nature Switzerland AG
About this chapter
Cite this chapter
Rodríguez, E.M., Guerra, M.M., Ortega, E. (2019). Physiopathology of Foetal Onset Hydrocephalus. In: Limbrick Jr., D., Leonard, J. (eds) Cerebrospinal Fluid Disorders . Springer, Cham. https://doi.org/10.1007/978-3-319-97928-1_1
Download citation
DOI: https://doi.org/10.1007/978-3-319-97928-1_1
Published:
Publisher Name: Springer, Cham
Print ISBN: 978-3-319-97927-4
Online ISBN: 978-3-319-97928-1
eBook Packages: MedicineMedicine (R0)