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Review

In Vitro Models of the Blood–Cerebrospinal Fluid Barrier and Their Applications in the Development and Research of (Neuro)Pharmaceuticals

Department of Pediatrics, Pediatric Infectious Diseases, Medical Faculty Mannheim, Heidelberg University, 68167 Mannheim, Germany
*
Author to whom correspondence should be addressed.
Pharmaceutics 2022, 14(8), 1729; https://doi.org/10.3390/pharmaceutics14081729
Submission received: 8 July 2022 / Revised: 11 August 2022 / Accepted: 16 August 2022 / Published: 18 August 2022

Abstract

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The pharmaceutical research sector has been facing the challenge of neurotherapeutics development and its inherited high-risk and high-failure-rate nature for decades. This hurdle is partly attributable to the presence of brain barriers, considered both as obstacles and opportunities for the entry of drug substances. The blood–cerebrospinal fluid (CSF) barrier (BCSFB), an under-studied brain barrier site compared to the blood–brain barrier (BBB), can be considered a potential therapeutic target to improve the delivery of CNS therapeutics and provide brain protection measures. Therefore, leveraging robust and authentic in vitro models of the BCSFB can diminish the time and effort spent on unproductive or redundant development activities by a preliminary assessment of the desired physiochemical behavior of an agent toward this barrier. To this end, the current review summarizes the efforts and progresses made to this research area with a notable focus on the attribution of these models and applied techniques to the pharmaceutical sector and the development of neuropharmacological therapeutics and diagnostics. A survey of available in vitro models, with their advantages and limitations and cell lines in hand will be provided, followed by highlighting the potential applications of such models in the (neuro)therapeutics discovery and development pipelines.

Graphical Abstract

1. Background

The research and development platforms of therapeutics targeted to the brain or aimed to be excluded from the central nervous system (CNS) depends mainly on their accomplishment in assessing the reciprocal behavior of brain barriers and the pharmacologically active agent. This effort is made to conquer the economic and technical burden on the pharmaceutical industry attributable to high risk and poor approval rates of neurotherapeutics. During the long journey of drug development, based on the nature of the product under study and in the cases of pregnant women, neonates, and any conditions with the possibility that drugs might have short- or long-term damaging effects, substantial data are required to prove that a substance is excluded effectively from the brain. On the other hand, when the aim is efficient drug delivery to the CNS, the development of neurotherapeutics faces a number of major challenges, with difficulty in delivering them to their desired site of action in the brain being on top. The presence of highly selective barriers (the blood–brain barrier (BBB) and the blood–cerebrospinal fluid (CSF) barrier (BCSFB)) and the complexity of the highly regulated brain environment are among the factors responsible for the elevated attrition percentage during the development of these CNS-targeting medicines.
The BBB and BCSFB are of utmost importance to pharmaceutical drug discovery, as both barriers provide obstacles to the penetration and delivery of therapeutics and diagnostics designed for the management of CNS complications and disorders. Presently, leading research has been concentrated on the brain vascular endothelium as a medicinal target to improve the delivery of brain therapeutics. However, the more poorly noticed BCSFB is also an essential gateway to the CNS, since it is broadly accepted that most substances, even macromolecules, that successfully reach the CSF, can find their way further to the brain parenchyma. Taken together with its considerable surface area, which is estimated to be ~25 to 50% the size of the inner capillary surface area of the brain [1], the BCSFB can be regarded as a target for the CNS delivery of (neuro)therapeutics [2,3]. Notably, the unique role of this barrier to regulate the composition of the CSF offers the BCSFB also the privilege of augmenting the levels of certain compounds and drug candidates in the CSF to accomplish therapeutic benefits and availability to the brain parenchyma.
Species relevant in vitro model systems of the BCSFB play an essential role in enabling the understanding of this barrier’s properties and developing potential interventional techniques and therapeutics. In the pharmaceutical, pharmacological, and toxicological fields, these models harbor the potential to be applied for investigational drugs/compounds screenings, permeability and transport assays, (neuro)toxicological evaluations, and mechanistic molecular pharmacological studies and related assays at the pre-clinical level. Due to many economic concerns, ethical issues, and the necessity to comply with the three R rules (the replacement, reduction, and refinement of animal subject experiments) and bypassing animal testing as far as possible, there has been a great effort in past decades toward establishing robust and convenient models to study basic characteristics of the barriers and also to facilitate the research and pre-clinical investigations of neurotherapeutics in the laboratory under controlled conditions. Such models should authentically mimic the in vivo microenvironment phenotype of the BCSFB and demonstrate representative properties of functional elements of tight junctions (TJs) leading to high transepithelial electrical resistance (TEER), restricted paracellular permeability, low non-specific pinocytic activity, and the expression of receptors and transporters.
With regard to the brief introduction, and despite the challenges and drawbacks inherited by these in vitro model systems, it is noticeable that the area has gained significant attention over the previous decades. These in vitro platforms offer a less resource-intensive alternative approach compared to animal models, in addition to an ease of manipulation, which gives rise to their exploitation during high-throughput end-point drug screenings for the development of drug candidates, and also for providing insights into the mechanistic principles governing CNS drug delivery. Furthermore, such models can be helpful to enlighten the principal translocation routes of small-molecule drugs and constantly increasing number of biopharmaceuticals. To this end, the present review sets out to summarize the efforts and progresses made to this research area with a notable focus on the attribution of these models and techniques to the pharmaceutical sector and the development of neuropharmacological therapeutic and diagnostic agents. It is hoped that researchers new to the field find this paper to be a straightforward guide to figure out the suitable in vitro models and optimal techniques for their particular interest.

2. Structure and Physio-Anatomical Features of the BCSFB

A more precise understanding of the BCSFBs physiological and anatomical attributes will pave the way toward the rational development of CNS-targeting small and large molecule therapies, aiming to treat a wide range of neurological conditions. The structure and physio-anatomical features of the BCSFB have been the subject of numerous excellent and comprehensive reviews [1,4,5,6,7,8], so only those features relevant to pharmaceutical/pharmacological research and neurotherapeutics development will be briefly discussed here.
The choroid plexus, as a villous connective and highly vascularized structure, is the major component of the BCSFB system that connects the two principal physiological circulatory systems, namely the peripheral blood circulation and the CSF bulk flow. The choroid plexus convolute consists of two tissue layers, an outer secretory roughly cuboidal epithelium and an inner underlying stromal core comprising an immense network of fenestrated leaky blood vessels with a rich extracellular matrix. Cellular and sub-cellular foundations of choroid plexus barrier properties are non-neuronal epithelial cells (defined as a subtype of macroglia, derived from neuroectoderm) and their adjacent TJs (Figure 1). From an anatomical point of view, this ventricular structure is protruded in the brain lateral, third, and fourth ventricles. The choroid plexus epithelium is in line with the ependyma, a cuboidal epithelium interconnected by gap junctions covering the lumen of the cerebral ventricles; however, despite sharing a common embryological origin, these two cell types are quite distinct [9].
The choroid plexus contributes to the production and secretion of CSF (approximately 500 mL per day in humans, or more precisely ~0.4 mL/min/g choroid plexus in adult mammals) and cerebral homeostasis by regulation of the blood–CSF exchange taking advantage of numerous transport systems allocated in a polarized configuration among the apical or brush border (luminal, CSF-facing) and the basolateral (abluminal, stroma-, or blood-facing) membranes of the choroidal epithelial cells [10]. The choroid plexus epithelium supplies micronutrients, hormones, growth factors, neurotrophins, and neuroprotective proteins to the CSF–brain nexus and uniquely provides micronutrients, such as ascorbic acid (vitamin C) and folate, for neuronal networks and glia [11,12,13,14]. Taken together, the choroid plexus acts in a complementary collaboration with brain microvessels to furnish regulatory factors and essential substances to meet the cerebral metabolism requirements.

2.1. Tight Junctions

As a formidable structural and metabolic barrier, the BCSFB, similar to the BBB, prevents toxic substances in the circulation from reaching the brain. The structural elements of this barrier are dense bands of impermeable TJs near the apical (CSF-facing) surface of adjacent cells of the choroid plexus epithelium. TJs are elaborate networks of specialized membrane microdomains, which tighten adjacent epithelial cells together and consequently limit the free diffusion of hydrophilic and polar molecules. Morphologically and structurally, TJ complexes comprise transmembrane (integral) proteins as occludin, the claudins, or junctional adhesion molecules (JAMs), and the associated cytoplasmic proteins of the membrane-associated guanylate kinase-like homologues (MAGUKs) family, termed zonula occludens proteins ZO-1, ZO-2, and ZO-3. Non-MAGUK peripheral junction proteins, such as cingulin, AF-6, symplekin, 7H6, and 4.1R have also been shown to be associated with TJs [15]. Claudins 1, 2, 3, and 11 have been described as the dominant members of this family present in choroid plexus epithelial cells [3].

2.2. Specific Markers and Receptors

A large number of transport proteins and receptors are expressed in choroid plexus epithelial cells for mediating and controlling multiple functions of this tissue and acting to transduce key humoral signals between the blood and the CNS. For instance, the choroid plexus serves to transport an assortment of amino acids bidirectionally (e.g., glycine, alanine), hormones (growth hormone, thyroid hormones, insulin, and melatonin), proteins, and growth factors (transthyretin or TTR, transferrin, prolactin, vasopressin, leptin, insulin-like growth factor 1 and 2, or IGF-1 and IGF-2, respectively) and pharmacologically active agents (e.g., β-lactam antibiotics, cimetidine, and benzylpenicillin) [16]. The choroid plexus also expresses high levels of receptors for low-density lipoprotein (LDLR), LDLR-related protein 1 and 2 (LRP1 and LRP2), endothelin, serotonin (5-HT), formylpeptide receptor-like 1 (FPRL1), arginine vasopressin (AVP), and atrial natriuretic peptide (ANP) [17,18,19]. Among the markers mentioned above, transthyretin is considered a specialized feature of choroid plexus epithelial cells.

2.3. Transporters and Ion Channels

Choroid plexus epithelial cells possess multiple structural and functional attributes of a transporting epithelium and express a broad spectrum of uni- and bi-directional transporters and ion channels, distributed in a polarized fashion at the apical and basolateral surfaces. This polarized distribution between luminal/apical and abluminal/basolateral membranes enables a heavy yet regulated and directional movement of molecules/compounds across the BCSFB. Multiple transporter proteins, classified as solute carrier (SLC) transporters, are present and functional at the epithelial cells of the BCSFB for glucose, amino acids (acidic, basic, and neutral), peptides, organic anions, monocarboxylic acids, and a myriad of solutes and ions [20,21,22,23,24,25,26,27,28,29,30,31,32]. A summarized list of SLC transporters at the BCSFB is provided in Table 1.

2.4. Xeno- and Endobiotic Efflux Systems

One vital physiological commitment of the choroid plexus is to furnish effectual clearance of xenobiotics and potentially deleterious metabolic end products out of the CSF back to the circulatory system, and the choroid plexus is consequently nominated as kidney of the brain [33]. This aim is achieved through the high capacity of the choroid plexus for drug metabolism using metabolizing (reducing, hydrolyzing, or conjugating) enzymes and the existence of specialized transporters.
The principal choroidal drug transporters accepting a broad range of substrates belong to two superfamilies of transporters, namely ATP-binding cassette (ABC) carrier transporters and the SLC family [34]. The superfamily of ABC transporters functions as ATP-driven efflux pumps, playing an essential role in the barrier functionality of the BCSFB. The human genome embraces 49 genes encoding ABC transporters, arranged in distinct subfamilies of A to G. The three subfamilies B, C, and G contain transporters that essentially handle xenobiotics and are expressed to a greater extent in the barrier and excretory tissues compared to all other cells [35]. From a pharmacological point of view, these transporters can be regarded as a double sword, being neuro- and xeno-protective on one hand, when they limit neurotoxicants entry into the brain, whereas they might hinder effective drug delivery for CNS pharmacotherapy on the other hand. Multiple ABC transporters are expressed in the choroid plexus epithelium, playing a pivotal role in CNS therapeutics delivery, and absorption, distribution, metabolism, and excretion (ADME) of drugs. ABCC1/MRP1, ABCC4/MRP4, and ABCC5/MRP5 are located in the basolateral membranes, enabling active transport from the cytosol toward underlying fenestrated capillaries, while the efflux pump P-glycoprotein (P-gp/ABCB1/MDR1), ABCC2/MRP2, and ABCG2/BCRP are positioned in the apical membrane, allowing transport from the cytosol toward the CSF [28,36,37,38,39,40,41,42,43].
In addition to the aforementioned efflux pumps expressed at luminal and/or basolateral membranes, diverse drug-metabolizing enzymes, responsible for inactivating endogenous and exogenous molecules, are expressed in the choroid plexus, including glutathione S-transferases (GSTs), UDP-glucuronosyltransferases (UGTs), flavin-containing monooxygenases (FMOs), epoxide hydrolases (EHs), and phase I metabolizing enzymes, namely multiple isoforms of the cytochrome P450 family [4,41,44,45,46,47,48,49,50,51].

3. Survey of Available Platforms

Given the principal role of the BCSFB and restrictions of animal and human investigations, there is a continually increasing demand for new and advanced model systems and in vitro techniques. The following sections describe and summarize the current knowledge related to in vitro models of the BCSFB and focus on their relevance to (neuro)therapeutics research and development. These in vitro model systems are arranged in a diverse range of technical, biological, and physiological complexities (Figure 2). It should be commented that no single model is a perfect representation of the in vivo complexity, and each model inherits pros and cons, which should be considered upon investigations (Table 2).

3.1. Static Monolayer Cultures Using Bicameral Systems

Basic 2D models are static systems of choroid plexus epithelial cell monocultures, predominantly based on semipermeable microporous membranes. The static culture systems are defined as those in which cultured cells are kept in the absence of physiological fluid dynamics. In this context of negligible fluid shear stress, the exchange of nutrients and waste takes place using diffusion [52]. These in vitro BCSFB epithelial barrier models have conventionally been founded on bicameral systems (alternatively called cell culture inserts), which possess a cell culture support fabricated from microporous permeable polymer membrane, generating distinct environments on opposing surfaces of a cell monolayer [53]. Upon cultivation, choroidal epithelial cells usually form a tight and polarized confluent monolayer, which can be regarded as a physiologically active cell culture model of the blood–CSF barrier. If cells are grown on top of the semipermeable membranes, this system’s upper or apical compartment simulates the ventricular space, and the basolateral/lower reservoir mimics the blood/stroma side. In the case of the inverted culture of epithelial cells on the lower face of inserts, such as in the method described by Dinner et al. [54], the apical and basolateral directions would be switched. In this manner, either transfer directions (blood-to-CSF and CSF-to-blood) are easily and independently accessible for further analyses.
The facile processing of filters makes this system an appropriate candidate for high-throughput screening (HTS) studies of drug permeability, binding affinity measurements, and transport kinetics. In general, this model type is well suited to explore in detail transport mechanisms (ranging from passive diffusion, facilitated transport, receptor-mediated transport) and cellular transmigration processes, and to investigate the permeability of potential drug candidates as well as the transepithelial resistance. Other applications include the analysis of the metabolism of molecules en passage and the determination of the ensuing metabolites [10]. Based on an experimental design proposed by Strazielle and Preston [10], these model systems can be used to confirm the active transport of an investigational compound, by observing an imbalance across both compartments containing an initial equimolar concentration of the compound and its accumulation against a concentration gradient.
In spite of the advantages pertained to the ease of manipulations of static bicameral model systems, there are disadvantages mainly due to the two-dimensional nature, the absence of fluid-induced shear stress, and the lack of fundamental features, as 3D structure, cell–cell or cell–matrix interactions, comparable TEER, fluid-induced shear stress, vasculature, and the presence of other accompanying cells, necessary to establish a flawless and authentic in vitro model [55]. The “edge effect” arising from the higher permeability of inserts at the outer rim of the membrane is another drawback that should be considered [56]. In addition, cell culture insert-based model designs are not specialized for microscopy and, therefore, do not provide an ideal platform for direct visual inspection of pharmaceuticals’ transport behavior and mechanisms. According to Larsen et al. [53], the main technical hindrance lies in the distance separating the cell monolayer and the below chamber that is not in compliance with the working distance of high magnification, high numerical aperture objectives. As an alternative solution, excised fixed semi-permeable membranes can be mounted on coverslips for imaging; still, this approach provides static snapshots of the transport status and is incapable of capturing the dynamics of (drug)substances’ cellular trafficking. However, commercially available image-compatible platforms can circumvent this obstacle and facilitate high-resolution (live)-cell imaging.
Special attention should be taken for the proper selection of the type of culture inserts for the experiments and the coating of filters prior to cells seeding.
Selection of the culture insert: the choice of the cell culture insert depends on the application and desired outcomes of the study. A broad range of inserts with varying permeable membrane materials (polycarbonate (PC), polyester polyethylene terephthalate (PET), and collagen-coated polytetrafluoroethylene (PTFE)) and different porosities (pore size and pore density) are available commercially from several manufacturers. The selection of the cell culture insert is a crucial step in the design of preliminary experiments. It is recommended that the adsorption and retainment profile of investigational compounds to different types of filter materials are clarified in each experimental setting. The optical properties of filters should also be considered. Transparent membranes offer the advantage of assessing cell culture monolayers using phase-contrast microscopy. A 0.4 µm pore size is well suited to examine the transfer of solutes and a large number of proteins. Theoretically, free diffusion is not restricted in the case that the pore diameter is wider than 20-fold the effective diameter of the compound [10].
Coating material: precoating the semi-permeable membrane support using different basal lamina components and comparable compounds (laminin, collagen, fibronectin, Matrigel, and poly-D-lysine) can enhance cell attachment, growth, and spreading to form a confluent monolayer. As the most appropriate matrix component for culturing choroid plexus epithelial cells, laminin is considered the principal glycoprotein component of the basement membrane serving to expedite cell attachment, spreading, and growth [57]. It was highlighted that a laminin coating ensured the formation of confluent monolayers of porcine choroid plexus epithelial cells, owing to improved adherence and proliferation [58]. Alternatively, collagen is more cost-effective yet still equally efficient. Nevertheless, collagen might contribute to enhancing the adhesion and growth of fibroblasts available in the cell suspension, consequently disturbing the purity of the cultures [59].

3.2. Co-Culture Models

To generate a model resembling more closely the in vivo scenario, possessing functional TJs, high TEER, and expression of specific transporters and enzymes, co-culture systems can be developed [60,61,62,63,64]. To enhance the quality of BCSFB in vitro models, cell culture barrier models can be configured in complex co-culture systems, in which choroid plexus epithelial cells are grown on porous cell culture inserts alongside endothelial cells, mesenchymal (e.g., pericytes), and/or glial cells either cultivated on the bottom of a multi-well plate into which the insert is located (non-contact) or seeded on the opposite side of the inserts containing epithelial cells (leading to a so-called back-to-back contact co-culture).
A co-culture model of choroid plexus epithelial and endothelial cells cultivated on opposing surfaces of cell culture filter inserts has recently been described [65]. To our knowledge, it is the first report of establishing a co-culture in vitro model of the BCSFB. Introducing the vascular component to the model systems, not only results in an upgrade of the model to a more physiologically related one, but also can provide a platform to study the emerging roles of the choroid plexus vasculature in organ function. This can further pave the way for studying dysregulated interactions, such as the gut–brain axis and even clarifying therapeutic targets in various disorders [66]. Despite exhibiting often excellent barrier properties and a more resemblance to an in vivo phenotype, co-culture models are in general labor- and capital-intensive.

3.3. 3D Cultures and Organoids

State-of-the-art three-dimensional (3D) culture systems, including matrix-based and matrix-free models, such as organoids and spheroids, have been recognized as the next advancement from static mono- and co-cultures. These 3D platforms with diverse features constitute promising alternatives to animal models and 2D cell culture systems in an in vitro tool to recapitulate the complex features of cerebral barriers [67,68]. In order to recapitulate the choroid plexus cytoarchitectural arrangement and a model resembling physiological conditions more closely, 3D cultures and organoids can be approached. Compared to other, less sophisticated in vitro counterparts, these models most accurately reflect the BCSFB properties and are valuable alternative tools to the use of animal subjects in CNS-oriented drug discovery programs.

3.3.1. 3D Explants and Cultured Cells in a Scaffold System

The simple design of appropriate cells cultured in a suitable scaffold or gel system, can be instrumental in the generation of high-throughput in vitro BCSFB models. In spite of few experimental experiences with the BCSFB, the currently available knowledge of BBB cells cultured in 3D platforms can be extended to the BCSFB field [69,70,71]. Three-dimensional explants of the choroid plexus can be generated from fragments of choroid plexuses, dissected from animals, human postmortem tissue, or human surgical samples cultured in suitable matrices, such as Matrigel and other commercially available hydrogels [72,73]. Three-dimensional explant platforms of mouse, rat, and human choroid plexus have been well prepared and maintained in the culture [72,74,75,76].

3.3.2. Organoids and Self-Organized 3D Models

Organoids, as defined by Huch et al., are 3D structures derived from either pluripotent stem cells or neonatal or adult stem/progenitor cells, in which cells spontaneously self-organize into properly differentiated functional cell types, and which recapitulates at least some function of the organ [77]. In the case of brain organoids, these accurate and versatile in vitro models reproduce several attributes of the brain barriers, including the expression of tight junctions, molecular transporters, and drug efflux pumps, and are critical tools for the study of brain barriers transport and the development of theranostics that can reach the CNS [78,79]. The potential to be scaled up to a high-throughput format, the ease of culture, and the miniature size nominate these multicellular organoids as robust, reliable, and predictive platforms to analyze and screen brain-penetrating compounds for the discovery of new and optimized treatment approaches for various neuropathologies.
Organoids of the choroid plexus as in vitro research platforms, derived from human pluripotent stem cells (human embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs)) recapitulate fundamental morphological and functional attributes of this organ [80]. These sophisticated models overcome the disadvantage of species-to-species differences and can improve our understanding of the development/function of the human choroid plexus, which due to the lack of experimental access to this vital brain tissue, is still elusive. A pluripotent stem-cell-derived organoid model of the choroid plexus has been developed by Pellegrini et al. [81,82], which brings about a tight barrier and reliably exhibits features of CSF production/secretion and the selective transport of molecules. According to the authors, this model could predict the permeability of pharmacologically relevant compounds qualitatively and quantitatively.
The aforementioned organoid models are devoid of vasculature, which can be regarded as both an advantage (in case of isolated investigations in a context free of interfering secondary complications) or a disadvantage (resulting in limited oxygen and nutrient delivery to the inner-most regions) depending on the study purpose. Despite the apparent drawbacks of lacking fluid flow-induced shear stress and burdensome establishment, this model platform is cost-effective, reproducible, and can serve as a pre-clinical array to narrow the translational gap between animals (mainly rodents) and human clinical trials. As a pre-clinical model scheme, patient-derived organoids may also pave the way toward the individualized translation of therapeutics [83]. Hypothetically, these 3D in vitro models offer an acceptable homogeneity and complexity level, enabling low- to a medium-throughput screening of a substantial number of investigational compounds to be evaluated in the process of novel therapeutics development. Other potential applications embrace evaluation of the (neuro)toxicological profiles of investigational compounds, lead compound identification/optimization phase studies, and structure–activity relationships (SAR) analyses. Taking advantage of automated microscopy and robotic-assisted technologies, the throughput of this model platform can be further enhanced [56].

3.3.3. Three-Dimensional Bioprinting Strategies

Three-dimensional bioprinting, as an additive manufacturing technology for modeling of user-defined biological samples, has emerged as a promising tool for the expansion of the BBB models. There is no known report of the in vitro 3D-bioprinted models of the BCSFB until now, but there are studied cases of the BBB [84,85,86,87,88], which have the potential to be adapted by the researchers in the field of the BCSFB. In principle, major printing modalities of inkjet-based, extrusion-based, and light-assisted bioprinting can be exploited to establish models with a high level of heterogeneity and biomimicry, which possess great potential as drug screening platforms [84]. Indeed, 3D printed models provide advantages of uniform and reproducible manufacture, minimal operating time, diminished user error, precisely controlled size, flexibility, and a high throughput compared to traditional techniques [89].

3.4. Dynamic Models and Microfluidic Platforms

To generate experimental conditions largely comparable to the in vivo environment, the impact of hydrostatic pressure and fluid-induced shear stress can be incorporated into in vitro model platforms. In the light of this phenomenon and with further technological advances, in addition to static models, dynamic systems and culture in microfluidic chamber devices have been evolved, in which a tunable shear stress is induced by a continuous flow of culture medium, in either pump-based or pumpless dynamic configurations [52].
Microfluidic device-based models, or so-called organ-on-a-chip systems, are dynamic models with a precisely controlled periodic physiological fluid flow that tend to augment the survival of three-dimensional cultures and organoid models, with improved nutrients/wastes and oxygen exchange, and more realistic dimensions and geometries. Microfluidics-based approaches, as next generation drug testing tools, rely on spatially resolved compartments joined together via microgrooves, allowing cell-to-cell interactions, precise control of the 3D cellular and extracellular matrix, and the flow of small amounts of fluids [67]. The integration of functional organ elements onto these structures enables the study of multi-organ interactions and dynamics of drug activities [90]. The rationale is that these emerging in vitro model platforms reconstitute the choroid plexus-mimetic microenvironment more accurately, which enables effective modeling of this tissue for (neuro)therapeutics development and research. Furthermore, the advancement in the 3D printing technology, nanofabrication, integrated sensors, and the versatility of human-derived stem cells have contributed largely to boost the field [91,92]. However, the system setup is sophisticated, time-consuming, and requires specialized equipment, significant resources, and technical skills compared to conventional static models, leading to low-throughput screening capabilities and hindering their broad application. Despite the widespread use of dynamic models, such as cone–plate apparatuses, microporous hollow fibers, and microfluidic-based devices in the generation of in vitro BBB models [93,94], their application and usefulness remain to be adapted for their BCSFB counterparts.
In principle, the high-fidelity microfluidic designs used for the development of BBB-on-chip models, have the potential to be tailored as BCSFB models. They are suitable for co-culturing various cell types, are compatible with high-resolution imaging modalities, allow the monitoring of intracellular and extracellular responses, and have the potential to incorporate patient-specific stem cells towards personalized human brain barrier chips. Accordingly, complex microfluidic BBB devices are described, which can be used as preclinical models to screen brain-targeting drugs, evaluate neurotoxicity, and recapitulate transport processes in the brain under recirculating perfusion [90,95,96]. These BBB-on-a-chip microfluidic models combine the benefits of both in vitro and in vivo models, show significant barrier integrity, and demonstrate a high capacity as drug permeability models using representative drugs and compounds, such as caffeine, cimetidine, doxorubicin, propranolol, antipyrine, carbamazepine, nitrofurantoin, and fluorescent-tagged dextrans [86,90,97,98,99,100].

4. Survey of Available Cells

Concerning cell type (origin and species), cell culture-based models of the BCSFB can be based on two different cell varieties, namely cerebral-derived or noncerebral-derived epithelial cells obtained from mammalian and non-mammalian species. Each category can be further divided into primary or continuous cell lines/immortalized cells in terms of proliferative potential. The time-consuming and laborious preparation process of primary and low passage cells has paved the way for employing either cell lines established without genetic manipulation (e.g., from tumors), or laboratory-generated/manipulated immortalized and transfected cell line models. Besides immortalization, these kinds of genetic manipulations can reintroduce the vanished characteristics of the models compared to their respective in vivo tissue. Here, various main cell source details are discussed and summarized in Table 3; however, it is essential to be aware that the selection of cell origin and cell type may have a decisive impact on the experimental costs and outcomes, as well as the reproducibility of obtained results.

4.1. Cerebral Originating Cells

The application of human-origin primary cells as in vitro models is restricted due to ethical and technical issues. Therefore, small and large animals have been sources of primary choroid plexus epithelium for various studies. Primary cultures of choroid plexus epithelial cells have been established from various species, including mouse [9,101,102,103,104], rat [16,105,106,107,108], pig [33,58,109,110,111,112,113,114,115,116,117], cow [118,119], sheep [120,121], rabbit [122,123,124], and non-human primates such as rhesus macaque [125]. Canine choroid plexus cells have been isolated as well, though being more challenging compared to other species [16].
Human primary choroid plexus epithelial cells can be obtained from aborted embryos, directly after surgical removal, or postmortem [3,126,127]. However, postmortem-derived human samples may have disturbed viability and functionality depending on the time elapsed after death and could be impacted by the health status of the subject in terms of infections, disorders, injuries, and medication history [3,128,129], and, due to limited applicability for functional assays and usual vectorial transport studies, are gradually abandoned by the researchers in the course of time. HCPEpiC from ScienCell Research Laboratories (also obtainable from other companies) are another human source for primary epithelial cells of the choroid plexus [130,131]. According to the manufacturer, further expansion of these cell populations for 15 doublings is guaranteed. However, multiple concerns are still inherited by this commercially available culture in terms of its origin, nature, and morphology [3].
The experimental procedure to isolate primary choroid plexus epithelial cells from different species and establish a pure cell culture devoid of contaminating cell types has been described comprehensively elsewhere and is not brought up here for the sake of brevity [9,132,133]. Principally, the freshly isolated primary cells retain the differentiated properties of the choroidal epithelium and exhibit a myriad of morphological- and biochemical-desired properties, yet the disadvantages are a small culture size, a limited proliferative potential, and a diminished viability over time after dissection. Therefore, prompt transfer to a suitable in vitro setting is crucial for establishing viable cultures [9]. Likewise, their purity, functionality, recovery, and survival rate explicitly rely on the animals’ health status, isolation approaches/techniques, and applied cultivation conditions [128].
Despite the outstanding contribution of primary cells in comprehensive studies of transport mechanisms and the molecular pharmacology of therapeutics, their potential to be exploited in high-throughput screenings of CNS-acting drugs is rather limited. Accordingly, spontaneous continuous cell lines, immortal/tumor cell lines, and immortalized cells overcome this issue inherited by primary cells and can be propagated easier and with minor limitations.
Z310 and TR-CSFB cells, as rat immortalized cell lines carrying the simian virus 40 large T-antigen gene, are well-characterized choroid plexus epithelial cells that have found their way as suitable cells [25,134,135,136,137,138,139,140]. TR-CSFB cells (five cell lines, TR-CSFB 1–5) exhibit a polygonal-shaped morphology comparable to primary cultured rat choroid plexus epithelial cells. Immunohistochemically, the TR-CSFB cell lines express TTR, apically located Na+, K+-ATPase, and the efflux transporters ABCB1/MDR1a, ABCC1/MRP1, and ABCG2/BCRP [141]. However, it has been reported that TR-CSFB cells seeded on culture inserts develop a net TEER of only approximately 50 Ω⸱cm2. The in vitro transepithelial electrical resistance of the Z310 cells monolayer is declared to reach 150–200 Ω⸱cm2 [142]. According to Strazielle and Ghersi-Egea [59], Z310 cells do not affirm a high degree of structural diffusion characteristics indispensable for an eligible BSCFB model and to investigate transepithelial transport processes. Upon culturing on permeable filters, the paracellular diffusion barrier exhibited by Z310 cells is unsatisfactory compared to that of pig counterparts [135,143].
Murine choroid plexus carcinoma cell lines ECPC3 and ECPC4 are isolated from transgenic mice harboring the viral simian virus 40 large T oncogene under the transcriptional control of an intronic enhancer region from the human immunoglobulin heavy chain gene. These cells have shown acceptable stability in culture in a time span of a year [144,145]. The SV11 is another mouse choroid cell line obtained from transgenic mice expressing the SV40 antigen in the choroid plexus [146,147]. The latter has not been characterized in terms of choroidal differentiation status.
The PCP-R cell line has been established based on primary porcine choroid plexus epithelial cells (PCPEC). The cell line exhibits a regular polygonal pattern, expresses junctional proteins, and develops morphologically dense TJs. When cultured on cell culture inserts, this cell line exhibits characteristic barrier properties of TEER above 600 Ω × cm2 and a restricted permeability for macromolecular paracellular markers [143,148].
SCP, a sheep choroid plexus epithelial cell line, is another alternative option on hand [13,149,150,151,152,153,154]. A variant of this ovine finite cell line (prepared from brain choroid plexus of Ovis aries) is listed in the American Type Culture Collection (ATCC) as cell line SCP No. CRL-1700®, or as catalog number 89101302 in the European Collection of Authenticated Cell Cultures (ECACC). However, its applicability as a BSCFB model remains to be validated, since the establishment of junctional complexes and diffusion barrier characteristics essential for a BCSFB model is still an undetermined field for this cell line [59].
Human choroid plexus epithelial papilloma (HIBCPP) cells have been isolated by Ishwata et al. from a 29-year-old Japanese woman, being spindle, oval, and polygonal in shape with neoplastic and pleomorphic features [122]. The cell line prevails over the ethical and technical challenges with primary human cells, and thanks to its human origin, negates any species differences in studies. The advantage of this cell line is that it provides an inexhaustible source of proliferating cells, while preserving the differentiation properties after consecutive passages. On the other hand, the contact inhibition has possibly vanished in these tumor cells leading to disturbed basolateral/apical orientation due to potential overgrowing in multiple layers; nevertheless, there have been reports of establishing optimized protocols endeavored to address this drawback [54].
Other cells, derived from a fragment of a fourth-ventricle choroid plexus papilloma (originated from a 28-year-old male patient) [155], and the CPC-2 cell line, resected from a choroid plexus carcinoma in the left cerebral hemisphere of a 2-month-old boy [156,157,158], have also been reported. The usefulness of these two latter cell lines as a tight and impermeable model of the BCSFB is improbable and not validated.
An immortalized human choroid plexus endothelial cell line (iHCPEnC), expressing pan-endothelial markers and presenting characteristic plasmalemma vesicle-associated protein-containing structures has been generated by Muranyi et al. using transduction of primary human choroid plexus endothelial cells with the human telomerase reverse transcriptase. The resulting cell line grows as a monolayer with contact inhibition and is regarded as invaluable for the generation of in vitro BCSFB co-culture models, as well as contribution to the clarification of the choroid plexus endothelial–epithelial interplay [65].

4.2. Noncerebral-Based Cells (Surrogate Models)

Noncerebral-originating cells are considered surrogates in blood–CSF barrier modeling. Madin-Darby canine kidney (MDCK and MDCKII) cells, human colonic epithelial cell line Caco-2 cells, Ralph Russ canine kidney cells (RRCK), Lilly Laboratories Culture-Porcine Kidney 1 epithelial cells (LLC-PK1), and cells transfected with specific efflux transporters or pharmacologic targets can be employed as a surrogate cell-based model to assess the permeability of selected compounds in the presence and absence of overexpressed efflux transporters or to evaluate their mechanism of action and effectiveness [159,160,161,162,163,164,165,166]. The spontaneously immortalized MDCK-MDR1 cell line expressing P-gp/ABCB1, MDCK2-ABCB1 cells expressing ABCB1, and LLC-PK1/BCRP cells expressing the efflux transporter ABCG2/BCRP are examples of surrogate cells, which can be used [167].

5. Models Validation Criteria

An effective in vitro model should imitate essential characteristics of an in vivo setting, namely the reproducibility of solute permeability, a restrictive paracellular route, a realistic physiological architecture, the functional expression of transporters/enzymes/receptors, and the ease of culture [55]. This section delineates several standard methods used to qualify in vitro BCSFB models and to evaluate the extent to which the models retain their phenotype and differentiated functions. Thus far, no single model can attain all the strict requirements, but rather the most relevant model configurations for the particular experimental goal should be approached. Certainly, the morphology and quality of in vitro cell-based models can be assured by ultrastructural evaluations, including transmission electron microscopy (TEM) imaging [168]. The determination of the integrity and characteristics of in vitro paracellular barriers can be achieved by evaluating three distinct readily accessible parameters, which are TEER, the permeability for tracer hydrophilic molecules of various but known (predetermined) concentrations and molecular weights, and the qualitative/quantitative expression of TJ proteins and customary markers as well.

5.1. Barrier Morphology

Once confluent, an in vitro choroid plexus epithelial cell barrier represents a differentiated polarized morphology, whose ultrastructure can be assessed using standard imaging modalities, including immunofluorescence microscopy and TEM. The latter allows direct visualization of the cells, their organizational structure, and any possible defect or imperfection in the cellular monolayer. Nonetheless, only few studies have used morphology imaging of in vitro cellular barriers as a routine quality validation to assure a closeness to in vivo conditions [168].

5.2. Barrier Properties

The reproducible tightness of models can be ascertained by convenient measuring of the TEER. The TEER value is a well-acknowledged measure to appraise the ion permeability of cell layers, reflecting the passive conductance of the TJs to small inorganic electrolytes, and impedance analysis manifests the electrical capacitance of the barrier likewise. Since the TEER can be correlated to the amount and degree of complexity of functional TJs and the expression of microvilli and other membrane invaginations, a high TEER might reflect the resemblance to in vivo situations. As a non-invasive quantitative method, the TEER measurement provides the most selective approach towards evaluating barrier integrity [115,169,170]. Currently, two strategies are employed to measure the TEER, either by using tissue Volt–Ohm resistance meters (voltage-measuring chopstick electrodes, such as the commercially available instruments Millicel-ERS or EVOM) to measure ohmic resistance or impedance spectroscopy systems (such as the CellZscope chamber-type electrodes from nanoAnalytics) to measure impedance across a broad spectrum of frequencies. There are technical variations between the two methods; while CellZscope yields continuous real-time monitoring of electrical resistance, chopstick electrodes of Volt–Ohm meters can record measurements solely at defined time points. The resulting TEER measures are expressed in an ohm centimeter square (Ω⸱cm2) unit. Considering the effect of Ca2+ ion concentrations, electrode level of submersion, and electrode position relative to the filter, TEER values can differ across distinct systems.
The analysis of cell monolayers using impedance-based biosensor technologies commercially available (such as xCELLigence technology, which uses proprietary microplates E-Plates) can be regarded as a label-free and real-time measure. The electric impedance-based workflow for monitoring of barrier function shows the advantages of being non-invasive, fast, easy, and real-time compared to end-point assays.

5.3. Exogenous Tracer Permeability

Exogenous tracers or inert paracellular flux markers compatible with analytical conditions can provide beneficial information on the permeability status of model barriers towards lipid insoluble organic compounds. However, one should be aware of potential side effects and avoid unnecessary interactions of the tracers with experimental elements. Exogenous tracers come in various physicochemical properties, ranging from proteins and polysaccharides to small polar compounds (such as, but not limited to, mannitol, sucrose, inulin, and dextran) [171]. Most routinely used protein markers are purified plasma proteins such as bovine albumin (~66 kDa), human albumin (~66 kDa), bovine fetuin (~49 kDa), and horseradish peroxidase (40 kDa). It should always be noted that the choice of the protein marker relies upon the biological process being studied and the experimental design. Still, it should be antigenically distinguished from the cells under study. A wide selection of dextran conjugates in a diverse molecular weight range (MW 3, 10, 40, 70, 150, 500, and 2000 kDa) is also available. Owing to satisfactory water solubility, low toxicity, limited immunogenicity, and biologically inertness, dextrans are among the most popular exogenous tracers used to determine barrier tightness.
The permeation of such aptly labeled probes through the cell monolayer as a function of time can be quantitively measured and be relied on as a model validation factor. Using an optimized quantification method (can range from colorimetry, fluorescence spectroscopy, and radioactive scintillation) to measure the concentration of the tracer, the permeability coefficient ( P ) for each probe can be calculated based on the data obtained from a standard curve and the following equation (based on the assumption that the concentration of the tracer at the donor side remains relatively constant due to the restricted flux and short time frame of the assays).
d Q d T = ( P ) ( A ) ( C d )
where d Q / d T is the slope of the linear segments of the (standard) curve, A is the surface area of the filter, and C d is the tracer concentration in the donor reservoir. The permeability of the cell monolayer ( P e ), generally expressed in cm s−1 (or cm min−1), can be calculated using the following equation:
1 P e = 1 P t 1 P f
where P f is the permeability of the filters without seeded cells (empty filters) and P t is the total or collective permeability of cells cultured on filters.
Besides providing a BCSFB model quality index and an evaluation of the time-dependent establishment of the barrier, the obtained data from exogenous tracers’ permeability can shed light on the relative contribution of the paracellular compared to transcellular pathways to the overall permeability of an investigational compound. However, attention must be paid that the paracellular marker is chosen appropriately to match the size of the compound [59]. In the following, available dyes and tracers are discussed, and a summarized list of various tracers with their molecular weights and hydrodynamic radii is provided in Table 4.
Dyes: The movement of certain dyes, including trypan blue (an azo dye of 0.96 kDa) and Evans blue (a synthetic dye and a non-specific albumin binder, with a molecular weight of 67 kDa, which produces the Evans blue dye–albumin complex), and phenol red (phenol sulfonphtalein, active transport against concentration gradient) across choroid plexus epithelial cells represents the barrier capacity status of the models [109]. However, since such dyes bind to albumin, their application is more restricted to in vivo models, and since superior and more sensitive tracers are at hand, their use is limited [172,173].
Fluorescent tracers: FITC (fluorescein isothiocyanate), TAMARA (Tetramethylrhodamine), Texas Red®, and Alexa Fluor®-conjugated dextrans and inulin with various molecular weights are amongst the most popular fluorescent tracers. Conventionally, fluorescently conjugated dextrans, depending on their molecular size, can be used to assess the permeability of models to an ion (low molecular weights variants) and proteins/macromolecules (high molecular weight variants) [174]. Sodium fluorescein (376 Da) is a small molecular-sized marker hydrophilic in nature; with the excitation and emission wavelengths being 494 nm and 525 nm, respectively. Lucifer yellow (absorption 428 nm, emission 533–535 nm), as a small (potassium salt 522 Da, lithium salt 457 Da, and ammonium salt 479 Da) hydrophilic molecule can be used as a tracer for fluid-phase pinocytosis. The assessment of fluorescence intensity can be performed by fluorescence microscopy and/or measurement in a fluorescence plate reader. Working concentrations exhibiting adequate fluorescent signals must be empirically determined for each set of experiments.
Radiolabeled tracers: Small polar compounds radiolabeled with radioactive isotopes can be used, such as [3H]-mannitol (182 Da), [14C]-sucrose (342 Da), [3H]-sucrose, [3H]-inulin (~ 5 kDa), [11C]-inulin, [125I]-albumin (67 kDa), or polyethylene glycol (~4 kDa) [175]. The quantification of radiolabeled tracers is attained by measuring radiation counts by a liquid scintillation system or the quantitative autoradiographic (QAR) method. Despite providing superior sensitivity, radiolabeled markers necessitate special safety precautions and equipment.
Biotin tracers: Besides fluorophores and radioactive isotopes tags, biotin is another compound that can be used in conjugation with tracers to monitor the permeability of in vitro BCSFB models. Biotin-ethylenediamine (287 Da), biotin-dextran, or biotinylated-dextran amine (BDA) with various molecular sizes (3–70 kDa) are exemplary choices. Biotin-labeled exogenous tracers can be visualized both by light- and electron microscopy imaging techniques [172,176,177].

5.4. Functional Junctional Proteins and Transporters

Analysis of the expression and localization of junctional proteins and relevant carriers/transporters and their accordance with the choroidal epithelium in vivo can reveal information regarding the quality index of the model. Depending on the individual experimental goal, the characterization of specific markers, enzymes, receptors, adhesion molecules, and specific proteins/polypeptides, can be performed. Immunostaining of cell-type-specific markers and junctional molecules leads to a qualitative confirmation of barrier integrity of an epithelial monolayer.
To assess the efflux transporters’ activity, substrate accumulation assays are exploited. Various efflux inhibitors (such as cyclosporin A and MK-571) are used to perform such substrate accumulation assays, assuming that substrate-inhibitor pairs are appropriately matched for each efflux transporter. In this assay configuration, model cells are incubated with the desired substrate either with or without their respective inhibitors, and at the culmination of the experiment, normalized fluorescence of the cells is calculated. In the case of the functional expression of transporters, the uptake of substrate should be increased in the presence of a corresponding transporter inhibitor [178].
Directional transport: in another experiment variation, transporter activities can be evaluated using directional transport assays, in which inhibitors are only added on the side of the cells grown on cell culture inserts where directional transport is being assessed. Here, again, model cells are incubated with the desired substrate either with or without their respective inhibitors, and the fluorescence signal on the opposite filter side is measured. If efflux transporters are expressed functionally at apical or basolateral surfaces, then the directional transport of substrate in the presence of the corresponding inhibitor is enhanced [178].

5.5. Factors Critical to Cell Selection and Culture Conditions

The barrier-forming characteristics and functions of choroid plexus epithelial cells in vitro and their degree of differentiation are closely related to the culture conditions applied. There have been lines of evidence supporting that certain agents and conditions may induce tightening and strengthening of the barrier. The factors critical to cell type selection and culture conditions are discussed in this section.
Serum withdrawal also seems to impact choroid plexus cells to reach the full barrier function. Serum deprivation can tighten the epithelial monolayer and improve the cellular polarity [132,179]. However, the barrier-dismantling effect of serum is believed to be compartment-specific, meaning that it depends on the apical or basolateral exposure of the cell layer to serum. In the case of serum exposure to the apical (comparable to CSF side in vivo) surface of the choroid plexus epithelial cell monolayer, the barrier function is diminished vastly. On the other hand, when serum is applied to the basolateral (corresponding to the stroma/blood side in vivo) surface, the epithelial barrier function is scarcely affected [33,117].
Due to the fact that barrier properties of epithelia are modulated by cAMP-dependent pathways, the presence of membrane-permeable cAMP analogs such as 8-(4-chlorophenylthio)-cAMP (CPT-cAMP) or the adenylate cyclase activator forskolin can have an augmenting effect on TEER values of choroid plexus epithelial cell layers [33,180,181].
Corticosteroids (hydrocortisone, dexamethasone) application to in vitro models may increase the barrier tightness of the epithelial and endothelial cells monolayer. This phenomenon is proposed to be a consequence of the regulation of the expression and distribution of tight junction proteins upon corticosteroids treatment [182]. Dexamethasone, as a synthetic glucocorticoid, has been shown to improve barrier strength and can be exploited as a positive control to investigate the effect of various conditions on the barrier integrity [183,184,185].

6. Applications in (Neuro)Therapeutics Development and Research

Several lines of evidence propose that unique transport and barrier attributes of the choroid plexus, in conjunction with the blood–brain barrier, are among the decisive factors establishing cerebral bioavailability of therapeutics and diagnostics. A large number of innovative and highly specific active compounds and investigational pharmaceutical preparations are continuously invented/developed/manufactured during the research and development cycles by the pharmaceutical industry, yet the necessity to discover a practical strategy for the effective delivery of them to the CNS target site remains to be addressed properly. Leveraging a robust and authentic in vitro model of the BCSFB can diminish the time and effort spent on unproductive or redundant development activities by a preliminary assessment of desired physiochemical behavior of an agent toward this barrier. In addition, these model systems can prove helpful in elucidating the nature of permeability and transport mechanisms, with a resolution at the cellular and molecular level, as a focal step during the long journey of neurotherapeutics research and development.

6.1. Permeability Screenings and Studies

Any rational drug discovery project dealing with candidates targeted to the brain or requiring exclusion from the CNS to prevent possible side effects should contemplate permeability measurements and prediction studies. Hence, profiling brain/CSF permeability of investigational novel molecular species at preliminary stages of the drug development track is game-changing. To this end, models can predict the CNS permeability of novel or known compounds in relation to both the route and rate.
Most in vitro BCSFB models used for permeability studies/screenings and understanding the pharmacokinetics of CNS active compounds are variants of 2D static bicameral inserts, employed to measure compound movement across the cell monolayer cultured on the porous membrane support. When using these models, a possible disturbance of the mass balance of the initial amount of drug added to the donor chamber with the amount recovered on the opposite reservoir would provide rough evidence of either cellular accumulation or intracellular metabolism of the compound (unspecific adsorption to filter membrane or well plastic should be ruled out). Measuring the concentration of the test compound, using an appropriate method (ranging from chromatography techniques, mass spectrometry, and biological detections methods), as a function of time in one of the compartments and knowing the initial concentration on the opposing compartment yields to the clarification of active transport rate and direction. Subsequently, the permeability coefficient ( P e ) or apparent permeability coefficient ( P a p p ) (the latter not corrected for permeability of the filter) can be calculated. P a p p values, expressed in cm s−1 or cm min−1 unit, can be calculated according to the following equation:
P a p p   [ cm / s   o r   cm / min ] = ( d Q d t ) × 1 C 0 × 1 A
where d Q / d t is the permeability rate or rate of transfer to the receiver compartment, t is the incubation time (seconds or minutes), C 0 is the initial concentration in the donor compartment, and A is the filter surface area in cm 2 .
The above equation can be rearranged in terms of the concentration and expressed as follows:
P a p p   [ cm / s   o r   cm / min ] = V d × Δ M r A × M d × Δ t
where V d denotes the volume in the donor compartment in cm3; Δ M r is the total amount of compound in the receiver compartment after time t (seconds or minutes); M d is the donor amount (added at time 0); Δ t indicates the incubation time measured in seconds or minutes; and A is the filter surface area in cm2.
Eventually, the contribution of both filter and any coating/substrate can be subtracted from the total permeability P a p p or P t using the following equation leading to P e estimation:
1 P e = 1 P t 1 P f
where P f is the permeability of the filters without seeded cells (empty filters) and P t or P a p p is the total or collective permeability of cells cultured on the filters.
When applying dynamic microfluidic platforms to measure the permeability of a test (drug)substance added to the basal (mimicking blood side) channel toward the luminal (presenting CSF side) channel, the permeability coefficient P a p p can be determined using the following equation:
P a p p [ cm / s ] = C l × Q A × ( C b C l )
where P a p p is the measured or apparent permeability coefficient in the cm / s unit, C l is the luminal concentration, C b is the basolateral concentration, Q is the applied flow rate in the channel ( mL / s ), the term C l × Q   ( m ˙ a ) represents the mass transport rate ( mol / s ) across the membrane, and A   ( cm 2 ) is the membrane surface area through which the transport occurs.
The permeability coefficient of a cell layer-free device (blank, or P 0 ) can be subtracted from the measured P a p p to calculate the P e of the model toward the compound, according to the following equation:
1 P e = 1 P a p p 1 P 0
Alternatively, the transport of a given compound can also be calculated from the incremental clearance volume ( Δ V C L ) for each time point from the following equation [186]:
V o l u m e   C l e a r e d   ( Δ V C L ) = C a × V a C d
where C a and C d are concentrations in the acceptor/receiver and donor chambers at sampling time, and V a and V d are acceptor/receiver and donor solutions volumes at sampling time, respectively. Accordingly, the slope of the linear segment of the Δ V C L versus the time curve leads to the total P S product ( P S t ) in unit of volume/time, assuming that the (drug)substance concentrations in the acceptor/receiver chamber remain small. P S product P S e and finally P e can be obtained from the following equations, leading to the same results as previous equations.
1 P S e = 1 P S t 1 P S f
P e = P S e S
where P S t and P S f are the P S products for total model system and cell-free blank filter insert, respectively, and S is the membrane surface area in cm 2 .
An advantage, in the case of measuring the permeability using microfluidic devices compared to static bicameral cell culture filter inserts, is that the drug or substance can be supplied to the basal channel at a constant flow rate and the transported amount can be sampled from the apical/luminal channel with a constant flow rate. Consequently, the assumption that the concentration differences beyond the filter membranes remain constant throughout the experimental read-out is met, albeit in the static bicameral devices this difference decreases over time [187].
The permeation rates of various drugs (ranging from diazepam, propranolol, and morphine, cefadroxil, cyclosporin A, to antiretroviral therapies) [33,188,189], novel central-acting cholinesterase inhibitors (2-phenoxy-indan-1-one derivatives or PIOs) [190], proteins/hormones (thyroxine, leptin, β-amyloids, and TTR) [191], as well as transport kinetics of metals (manganese, iron, and copper) [192] have been assessed effectually with compartmentalized culture insert models.

6.2. Transport Mechanisms Studies and (Targeted)Drug Delivery

Thanks to their physicochemical nature, hydrophobic/lipophilic substances of a low molecular weight are capable of free diffusion across cell membranes. In contrast, as mentioned in previous sections, the unrestricted diffusion of hydrophilic chemical species is substantially hindered due to the effective closure and sealing of the paracellular shunt by TJs. Accordingly, access to the CSF is granted exclusively to those compounds that are transported actively by the corresponding transport systems in the plasma membrane of choroid plexus epithelial cells. Theoretically, therapeutic- and diagnostic agents that are effectively and successfully transported by the choroid plexus and remain to some extent unaffected by the metabolizing enzymes and efflux transporters are rapidly distributed throughout the CNS using the bulk flow of CSF. This is due to the fact that at the brain ventricles, the extracellular/interstitial fluid (ISF) and the CSF are separated from each other by the non-barrier/non-restrictive permeable layer of ependymal cells, leading to a direct continuity of these two components and the subsequent free exchange of substances within the extracellular space of CNS [193].
Models expressing the specific transporters found in choroid plexus epithelium could be harnessed to establish whether the permeation of a compound of interest is impacted by a specific carrier system (e.g., P-gp) and to provide comprehensive knowledge on the physiology and modulation of such transporters. From another point of view, several neurological disorders are assumed to be associated with the dysfunction of transporters in the brain, and the detailed region-specific knowledge of transporters and their interaction with investigational therapeutics can prove helpful. In the case that transporters govern the permeability or uptake of a test compound, Michaelis–Menten kinetics can be obtained using nonlinear regression analysis of the concentration dependence influx. In vitro models have proven advantageous and constructive in studies of thyroxine [194], leptin [195], taurine [124], ascorbic acid [112], creatinine [196], and the neuroactive flavonoid resveratrol [197] transport across epithelial cells of choroid plexus. The in vitro BCSFB models have also proven helpful in library screening and identification of cell specific penetrating peptides for the choroid plexus epithelium using phage display techniques [76,198].
Therapeutic and diagnostic agents can pass the BCSFB via the transcellular route by employing one of the possible pathways, which are passive diffusion, facilitated diffusion, or vesicular transfer or transcytosis mechanisms. Generally, cellular uptake mechanisms based on endocytosis/transcytosis are the preferred cell entry route for many compounds. The endocytosis process can be categorized into two broad divisions of phagocytosis and pinocytosis. While the former is restricted to specialized cell types, the latter occurs in all cell types and can be subdivided into macropinocytosis, clathrin-dependent endocytosis (CDE), and clathrin-independent endocytosis (CIE) [199]. Contrary to endocytosis, transcytosis mechanisms are not well understood. Endocytosis and transcytosis across choroid plexus epithelium, studied at the subcellular level, can shed light on mechanisms of targeted delivery of therapeutics across this target site. These studies may rely on endocytosis inhibitors (including, but not limited to, chlorpromazine, genistein, methyl-β-cyclodextrin, and potassium depletion) to provide information concerning the endocytic pathway of compounds under study.

Drug Delivery Employing Transport Mechanisms at the BCSFB

Designing novel CNS therapeutics and pharmacologically active candidates that exploit the intrinsic transporting capacity of choroid plexus epithelial cells will prove helpful toward the generation of CNS drug delivery approaches through the BCSFB. Nevertheless, the choroid plexus has received little attention as a potential gateway for drug delivery to the brain, and there remain debates about its eligibility for this aim [200]. An evaluation of the design process effectiveness and rate of successful transport across this barrier can be achieved through preliminary mechanistic and transport studies adopting the available in vitro BCSFB models. Potential transporters that can be targeted to mediate drug delivery to the CNS across the BCSFB belong to the major categories of protein/hormone receptors, solute carriers, and amino acid transporters [201].
Targeting transcytosed receptors (receptor-mediated transcytosis (RMT)): From a pharmacological viewpoint, although most efforts have been concentrated on the BBB, yet the epithelial barrier of the choroid plexus is of considerable importance as a potential target for drug delivery to the CNS, and the subsequent maintenance of effective drug concentrations needed to treat conditions and disorders. Hitherto, transcytosis-mediated transport by interacting with hormone/protein receptors expressed on choroid plexus epithelial cells seems to be the most favorable mechanism exploited for this purpose. The conjugation of BCSFB-penetrating antibodies/peptides with drugs or the engineering of bispecific antibodies, harboring both a therapeutic arm and a BCSFB-transcytosing arm, are among the most promising attempts toward neurotherapeutics-developing pipelines. Other alternatives are vectors based on the native ligand or a fragment of the ligand of the transcytosed receptor.
CNS shuttling of therapeutic and diagnostic macromolecules can be achieved through RMT, an active transport mechanism involving transcytosed receptors, including transferrin receptor (TfR), insulin receptor (IR), insulin-like growth factor receptors (IGF1R and IGF2R), folate receptor FRα, and receptors responsible for lipoprotein transport, namely the low-density lipoprotein receptors (LDLRs) [202,203,204,205,206]. The RMT process is initiated upon ligand binding to the receptors present on the stroma/blood-facing pole of the epithelial cells, consequent ligand/receptor complex internalization and endosomal sorting, followed by ligand release on the apical membrane of the epithelial cells, and receptor recycling back to the basolateral surface. The targeting of transferrin receptors using anti-transferrin receptor antibodies has been the most studied approach so far.
Contrary to solute carrier-mediated transcytosis, targeting an RMT pathway circumvents the size constraints of therapeutic cargos, since it employs a vesicle-based transport rather than a stereoselective carrier. This Trojan horse approach, although being currently studied by many research groups, holds a few technical drawbacks, which need to be addressed. The saturation of transferrin receptors with endogenous transferrin from the circulation requires the transferrin receptor-targeted compound to compete with the natural ligand [207]. As well, the aforementioned RMT targets are also highly and broadly expressed in peripheral tissues and being involved in metabolically critical cellular tasks, arising off-target effects and safety concerns [208,209].
There are reports of enhanced CNS penetration of antineoplastic drugs (methotrexate), peptides (vasointestinal peptide, nerve growth factor), and tracer protein native horseradish peroxidase using anti-transferrin receptor antibody OX26, the mouse monoclonal antibody against the TfR [210,211]. The conjugation of macromolecules to a peptidomimetic monoclonal antibody with an affinity to the human insulin receptor (HIRMAb) has been the topic of multiple experiments using a similar paradigm [212,213]. A HIRMAb-iduronidase fusion protein, valanafusp alpha, is another attempt for efficient brain drug delivery as a therapeutic option for the Mucopolysaccharidosis Type I (MPSI) [214]. The in vitro models of the BCSFB can be helpful to evaluate the transport of anti-TfR antibodies (and obviously other antibodies directed against RMT targets) as vectors for the delivery of a non-lipid-soluble macromolecule into the CNS. The rate of transcytosis can be predicted according to the equations described in the previous sub-sections.
Solute Carrier-Mediated Transcytosis (CMT): Harnessing in vitro BCSFB models, mechanisms pertaining to solute CMT exploited as a drug delivery approach to the CNS, using various solute carrier transporters (SLC) present on apical and/or basolateral membranes of choroid plexus epithelial cells, can be substantiated. In this context, the SLC proteins experience a conformational alteration from an outward to inward-facing orientation and consequently translocate their substrates across the cellular barrier. Generally, small di- and tripeptides can be transferred using this mechanism through the SLC15A family members [202,215]. Although being a promising target, CMT approaches inherit a few disadvantages, such as susceptibility to vector’s stereochemical configuration, rigidity, conjugation regiochemistry, and competition between the drug and original substrate. Furthermore, since SLC-mediated systems are considered portals for the trafficking of small molecule drugs with comparable size and structure to the original ligands, care should be taken upon the cargo’s size to mimic SLC’s original substrates and the vector-drug ligation design to prevent the loss of vector affinity towards the protein carrier [201]. There are few well known examples of drugs that take advantage of this pathway at the BBB, including the anti-parkinsonian drug L-DOPA (through SLC7A5/SLC3A2), antiepileptic agents gabapentin, pregabalin, and valproic acid, choline esterase inhibitor donepezil (through SLC22 subfamily), and N-methyl-D-aspartate receptor antagonist memantine (through SLC22 subfamily). However, the distinct role of active transporters at the BCSFB for this delivery strategy remains to be further clarified [200].
Nano-therapeutics: Nanotherapeutics represent a promising area of CNS drug delivery approaches and have revolutionized the concept of drug delivery across brain barriers with some successful exemplars currently in the clinic [216,217,218]. Nanoparticles (NPs)-based platforms of various size ranges and designs can overcome intact or impaired brain barriers and facilitate efficient CNS drug delivery leveraging transport via multiple mechanisms, including paracellular pathway, cell-mediated transport, adsorptive-mediated transcytosis, RMT, CMT or ligand–receptor interactions [219,220]. Extensive and comprehensive knowledge within the nanomedicine field and available designs of nanocarrier-based drug delivery systems (such as, but not limited to, liposomes, dendrimers, polymeric NPs, polymeric micelles, nanocapsules, lipid-based nanoparticles, inorganic and gold NPs, quantum dots) are summarized elsewhere [219,220,221,222,223,224] and, therefore, are not discussed in detail here.
In principle, all the BCSFB in vitro model systems described so far have the potential to be explored to shed light on the mechanisms of nanotherapeutics interactions with the BCSFB, identify the impact of biological variables on NPs transport across the BCSFB, evaluate the biocompatibility of nanomaterials, help design nanomaterials interacting selectively with the epithelium of the choroid plexus, assess the safety and efficiency of pre-clinical formulations, model the effects of social/stress/immune/environmental factors on NPs transport across BCSFB, and meet the ongoing need of research in neurotherapeutics’ research and development [225,226].

6.3. Metabolites/Xenobiotics Transport(er) Regulation

The fate of drugs and xenobiotics metabolites, affiliated to several chemical classes, generated at choroidal epithelial cells and the relative contribution of various efflux pumps present at this barrier can also be investigated using these in vitro models. In addition, the potential effect of multiple metabolites, xenobiotics, and compounds on both influx into and efflux out of the CSF can be studied. Referring to static bicameral device models, efflux transport assays can be performed, and the efflux ratio is estimated by dividing the permeability value in the apical to basolateral (A−B) direction by the permeability value in the basolateral to apical (B−A) direction. With respect to efflux transport assays, the transport of the compounds under study is assessed in the presence of inhibitors of relevant transporter proteins and compared to the non-inhibited conditions. Namely, verapamil, GF120918, PSC833, and N-desmethyl-loperamide as inhibitors of ABCB1/P-gp, Ko143, and fumitremorgin C as an inhibitor of ABCG2/BCRPs, tariquidar and elacridar as inhibitors of both P-gp and BCRP, or MK571 as an inhibitor of various MRPs are examples that can be applied for such experiments [45]. As an alternative strategy, the inhibition of these efflux transporters potentially improves the pharmacokinetics of CNS therapeutic candidates. To this end, the efficacy of a range of compounds (mainly phytoestrogens) as modulators of BCRP/ABCG2 has been evaluated using in vitro BCSFB models [227]. Accordingly, genetic manipulation approaches, such as overexpression, knock-out, or knock-down of specific transporters can be approached.

6.4. In Vitro Molecular Verification of Pharmacological Activity

One of the main thrusts behind the development of in vitro BCSFB models has been to shed light on the pharmacological molecular mechanisms of compounds toward this interface. Using cell-based BCSFB models, in vitro activity, potency, and mechanisms of actions of investigational CNS therapeutics and compounds under study can be demonstrated in preclinical drug evaluations. Excerpts of case studies from distinct pharmacological categories describe investigations of the anti-inflammatory activity of synthetic matrix metalloproteinase (MMP) inhibitors [102], and events of receptor activation for drugs of potential abuse and hallucinogens have been studied [228].
Cell cultures of the rat choroid plexus have been used to elucidate the regulation mechanisms and phenomena of various receptors, including 5-hydroxytryptamine1c (5HT1c) (using mianserin as an antagonist, or (–)-1-(4-bromo-2,5-dimethoxyphenyl)-2-aminopropane as an agonist) [229].
Investigational anti-viral compounds can be tested in vitro in the BCSFB model platforms. In these experiments, the choroid plexus cells lines could be used as hosts and cellular substrates for viruses to perform a wide range of assays, including the cytopathic effects inhibition assay, the virus titer reduction assay, cytotoxicity assays, for the determination of the effective concentration of compounds under study, and even molecular mechanistic assays. For instance, the activity of nucleoside cytidine analogs as possible anti-HIV compounds was investigated in sheep choroid plexus cells as the cellular substrate for the Maedi-MVV virus [121]. In another study, the potential antiviral activity of teriflunomide against DNA viruses was shown using a choroid plexus epithelial cell model [230].

6.5. (Neuro)Toxicological Studies

Harnessing appropriate in vitro BCSFB models, the ever-increasing concern regarding hazardous compounds, such as environmental toxins, metals, pesticides, solvents, as well as the broad field of study of neurotoxicants, can be the target of both exposure (amounts permeated) and impact (effects exerted) investigations. The models have proven valuable in evaluating the influx/efflux of (neuro)toxins, understanding the associated neuroprotective mechanisms, elucidating the alteration of the BCSFB function by toxic compounds, and the involvement of the BCSFB in neuroinflammation [59].
Choroid plexus cells and models have been used to study underlying mechanisms of metal-induced toxicity and the transport of neurotoxicants metals including, but not limited to, mercury (Hg), as presumably the most studied neurotoxic metal, lead (Pb) [231,232,233,234,235], arsenic (As) [236], iron (Fe), manganese (Mn) [237,238,239], copper (Cu) [240], and cadmium (Cd) [241,242].

6.6. Pharmacological Interventions at the BCSFB

Transport data acquired through pharmacological interventive measures and modulation by hormones and drugs contribute to understanding the BCSFB homeostatic phenomena and barrier/transporting functions. In addition, establishing approaches for restoring aberrant choroid plexus and CSF dynamics during disease has been of great interest to researchers in pharmacological, neurological, and neurosurgical fields. In this sub-section, major pharmacological mediations used to study this barrier are reviewed.
Carbonic anhydrase inhibitors (acetazolamide, benzolamide): Pharmacological agents of different classes altering CSF dynamics are exploited to study CSF production and secretion. Prototype pharmacological agents to reduce the CSF production rate in the range of 50–100% are inhibitors of carbonic anhydrases, the enzymes responsible for the hydration of CO2 to produce carbonic acid (H2CO3), which spontaneously dissociates into HCO3ˉ and H+ [23,243,244,245,246]. Acetazolamide is one of the commonly employed drugs in this pharmacological category believed to lessen CSF secretion via its effect on the choroid plexus, and modulation of aquaporin-4 and perhaps aquaporin-1 [247].
Ouabain is a potent inhibitor of the Na+-K+-ATPase pump and consequently ion and fluid transport. This cardiac glycoside reduces the CSF production rate to a half basal level when placed on the luminal side of the choroid plexus epithelial cells. This aspect along with its neurotoxic side effects hinders its clinical application [11,22,248].
Amiloride, the NHE (basolateral Na+-H+ exchanger) inhibitor, blocks Na+ distribution into the choroid plexus from the blood-facing side [23,249]. This pyrazine diuretic can be exploited to study CSF dynamics.
Loop diuretics (furosemide, ethacrynic acid, and bumetanide) likewise attenuate Na+, K+, and Cl transport at choroidal epithelium by inhibiting NKCCs transporters [250,251,252], thereby assisting transport mechanisms studies at this barrier.
Disulfonic stilbene (DIDS) is an anion transporter blocker, inhibiting Cl transport into the CSF. DIDS is a stilbene-type inhibitor of anion exchangers, which can be used as a pharmacological tool to evaluate the activity of these ion exchangers at the choroid plexus. However, as a disadvantage, its effects are not specific, and a wide range of other transporters can be affected [253,254,255].
Ouabain, acetazolamide, furosemide, bumetanide, and DIDS inhibit CSF secretion from the luminal membrane of choroid plexus. Acetazolamide, DIDS, and amiloride are inhibitors acting on the basolateral/blood-facing membrane [7,256]. In other words, bumetanide and ouabain need to be administered intracerebroventricularly to access their apical target transporters. Such a limitation has suspended their current clinical application; however, effective delivery methods to the CSF would be promising.
The regulation of apical ion channels of choroid plexus by various substances, including serotonin, mianserin, and mesulergine, has been the topic of research [106,257,258]. The integrity of the BCSFB and transepithelial protein transport have been evaluated in the presence of an agonist (GSK1016790A) and an antagonist (HC067047) of the polymodally gated divalent cation channel TRPV4 (transient receptor potential vanilloid 4) [259,260]. Such interventional studies lead to mechanistic clues of the regulation/modulation of receptors and ion channels by investigational compounds.

6.7. The BCSFB and Choroid Plexus as a Drug Target in Various Diseases

In this subheading, the role of choroid plexus dysfunction in various diseases and in conditions where itself is a target, for instance, choroid plexus carcinomas, is discussed. Choroid plexus breakdown has been proposed and hypothesized in a wide range of neurological conditions, including aging, Alzheimer’s disease, Parkinson’s disease, epilepsy, stroke, neoplasms, perhaps psychiatric disorders, intracranial hypertension, and certainly varying types of hydrocephalus. Therefore, targeting this tissue may offer opportunities to translate neurotherapeutics from the lab to the clinic, and provide insights toward rational drug therapy to restore barrier function [261,262,263,264].
Strategies to diminish choroidal fluid turnover or to develop efficacious agents, which can manage altered CSF dynamics in disorders, including hydrocephalus, ventriculomegaly, cerebral edema, and intracranial hypertension, can be designed based on the obtained transport data at choroid plexus. In spite of a large body of knowledge of choroid plexus transporters and ion channels that mediate CSF secretion, effective pharmacological agents are sparse, and a gap regarding pharmacological substances capable of interfering still remains.
Supportive evidence proposes the involvement of the choroid plexus in neuroinflammation and neurodegeneration [265,266,267]. Examining the anti-inflammatory potential of a novel or already approved repurposed therapeutics continues to be a subject for major research [268]. Likewise, in vitro cell culture models of the choroid plexus can be approached to answer research questions of the therapeutic efficacy and associated mechanisms of the action/resistance of chemotherapeutic agents to find the right treatment options for the rare brain tumor group of choroid plexus malignancies [147,269].

7. Conclusions and Future Outlook

This review set out with the aim of rendering an overview of the up-to-date status of in vitro BCSFB models currently exploited, with a special emphasis on their applicability for research and development of CNS-affecting therapeutics and drug delivery to the brain. Taken together, a suitable model should exhibit the features of the restricted paracellular shunt, the expression of TJ proteins and choroidal markers, and the functional and polarized expression of BCSFB-specific transport mechanisms. In addition, the model should meet the features of availability, convenience, predictability, reproducibility, and extrapolation to human and in vivo settings.
Although a large body of knowledge on transporters, receptors, and drugs pharmaco-kinetics and -dynamics are derived from in vivo animal models and invasive techniques, the fragility of choroid plexus tissue, its small mass, and hardly accessible location along with ethical constraints leads to technical challenges during isolation and further experimental interventions. On the other side, a challenge to reproduce the complexity of the in vivo environment with in vitro models as much as possible still exists. It is generally recognized that the reproducibility of cell-based models can be imperfect, with day-to-day and lab-to-lab inconsistencies being observed. In the case of static monoculture/coculture bicameral devices models, technical challenges may prevail, e.g., the cells can have an incomplete surface coverage or even overgrow and subsequently produce multiple layers instead of monolayers.
Nevertheless, despite all the mentioned shortcomings, in vitro models remain desirable workhorses for routine applications in industrial and academic pharmaceutical studies. Still, it should be emphasized that since no single model can imitate the absolute in vivo physiological environments, for each experimental condition based on the desired outcomes, the question under investigation and the throughput level, a suitable configuration of model system-cell type-validation criteria should be chosen. In other words, by weighing the pros and cons of various platforms, a translationally relevant model of the choroid plexus appropriate for the desired research needs should be selected.
Despite the fact that in vivo, ex vivo, and in situ models (such as models based on isolated choroid plexus in extracorporeal perfusion systems, microdialysis, ventriculocisternal perfusion) may be considered as gold standards and ultimate test systems for exploring drugs transport and pharmaceutically related events, integrated and physiologically related in vitro models are more promising to eliminate the effect of species-specific differences. Furthermore, these in vitro models can be captivating when the aim is to survey transport and other pharmacological mechanisms solely at the choroid plexus in the absence of interfering biochemical effects and traces of the BBB and brain parenchyma. In these model systems, secondary effects originating from drug pharmacokinetics, distribution, and tissue blood flow are eliminated exceedingly.

Author Contributions

Conceptualization: F.D., H.S. and C.S.; literature review, writing and original draft preparation: F.D.; revision: F.D., H.S. and C.S. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by funding from the German Federal Ministry of Education and Research, BMBF (FKZ: 16GW0325 (BrainAim)). For the APC we acknowledge financial support by Deutsche Forschungsgemeinschaft within the funding programme “Open Access Publikationskosten” as well as by Heidelberg University.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

Abbreviations

2Dtwo dimensional
3Dthree dimensional
5-HT5-hydroxytryptamine
ABCATP-binding cassette
ADMEabsorption, distribution, metabolism, excretion
ANPatrial natriuretic peptide
ATPaseadenosine triphosphatase
AVParginine vasopressin
BBBblood–brain barrier
BCRPbreast cancer resistance protein
BCSFBblood–cerebrospinal fluid barrier
CDEclathrin-dependent endocytosis
CIEclathrin-independent endocytosis
CMTcarrier-mediated transport
CNScentral nervous system
CPchoroid plexus
CSFcerebrospinal fluid
EHsepoxide hydrolases
ESCsembryonic stem cells
FITCfluorescein isothiocyanate
FMOsflavin-containing monooxygenases
FPRL1formylpeptide receptor-like 1
GSTsglutathione S-transferases
HTShigh throughput screening
IGFinsulin-like growth factor
IGFRinsulin-like growth factor receptor
iPSCsinduced pluripotent stem cells
IRinsulin receptor
ISFinterstitial fluid
JAMsjunctional adhesion molecules
LDLRlow-density lipoprotein receptor
LLC-PK1Lilly laboratories culture-porcine kidney 1
LRPLDLR-related protein
MAGUKsmembrane-associated guanylate kinase-like homologues
MDCKMadin-Darby canine kidney
MDRmultidrug resistance protein
MMPmatrix metalloproteinase
MPSMucopolysaccharidosis
MRPmultidrug resistance-associated protein
NPsNanoparticles
NHENa+-H+ exchanger
PCpolycarbonate
PCPECporcine choroid plexus epithelial cells
PETpolyethylene terephthalate
P-gpP-glycoprotein
PIOs2-phenoxy-indan-1-one
PTFEpolytetrafluoroethylene
QARquantitative autoradiographic
RMTreceptor-mediated transcytosis
RRCKRalph Russ canine kidney
SARstructure–activity relationships
SLCsolute carrier
TAMARAtetramethylrhodamine
TEERtransepithelial electrical resistance
TEMtransmission electron microscopy
Tftransferrin
TfRtransferrin receptor
TJtight junction
TTRtransthyretin
UGTsUDP-glucuronosyltransferases
ZOZonula occludens

References

  1. Spector, R.; Keep, R.F.; Snodgrass, S.R.; Smith, Q.R.; Johanson, C.E. A balanced view of choroid plexus structure and function: Focus on adult humans. Exp. Neurol. 2015, 267, 78–86. [Google Scholar] [CrossRef] [PubMed]
  2. Johanson, C.E.; Duncan, J.A.; Stopa, E.G.; Baird, A. Enhanced prospects for drug delivery and brain targeting by the choroid plexus—CSF route. Pharm. Res. 2005, 22, 1011–1037. [Google Scholar] [CrossRef] [PubMed]
  3. Redzic, Z.B. Studies on the human choroid plexus in vitro. Fluids Barriers CNS 2013, 10, 10. [Google Scholar] [CrossRef]
  4. Kratzer, I.; Ek, J.; Stolp, H. The molecular anatomy and functions of the choroid plexus in healthy and diseased brain. Biochim. Biophys. Acta Biomembr. 2020, 1862, 183430. [Google Scholar] [CrossRef]
  5. MacAulay, N.; Keep, R.F.; Zeuthen, T. Cerebrospinal fluid production by the choroid plexus: A century of barrier research revisited. Fluids Barriers CNS 2022, 19, 26. [Google Scholar] [CrossRef] [PubMed]
  6. Ghersi-Egea, J.F.; Strazielle, N.; Catala, M.; Silva-Vargas, V.; Doetsch, F.; Engelhardt, B. Molecular anatomy and functions of the choroidal blood-cerebrospinal fluid barrier in health and disease. Acta Neuropathol. 2018, 135, 337–361. [Google Scholar] [CrossRef]
  7. Praetorius, J.; Damkier, H.H. Transport across the choroid plexus epithelium. Am. J. Physiol. Physiol. 2017, 312, C673–C686. [Google Scholar] [CrossRef]
  8. Benarroch, E.E. Choroid plexus--CSF system: Recent developments and clinical correlations. Neurology 2016, 86, 286–296. [Google Scholar] [CrossRef]
  9. Menheniott, T.R.; Charalambous, M.; Ward, A. Derivation of primary choroid plexus epithelial cells from the mouse. Methods Mol. Biol. 2010, 633, 207–220. [Google Scholar] [CrossRef]
  10. Strazielle, N.; Preston, J.E.; Nag, S. Transport across the choroid plexuses in vivo and in vitro. Methods Mol. Med. 2003, 89, 291–304. [Google Scholar] [CrossRef]
  11. Johanson, C.E.; Keep, R.F. Blending established and new perspectives on choroid plexus-CSF dynamics. In Role of the Choroid Plexus in Health and Disease; Praetorius, J., Blazer-Yost, B., Damkier, H., Eds.; Springer: New York, NY, USA, 2020; pp. 35–81. [Google Scholar]
  12. Spector, R.; Johanson, C.E. The nexus of vitamin homeostasis and DNA synthesis and modification in mammalian brain. Mol. Brain 2014, 7, 3. [Google Scholar] [CrossRef] [PubMed]
  13. Gee, P.; Rhodes, C.H.; Fricker, L.D.; Angeletti, R.H. Expression of neuropeptide processing enzymes and neurosecretory proteins in ependyma and choroid plexus epithelium. Brain Res. 1993, 617, 238–248. [Google Scholar] [CrossRef]
  14. Stopa, E.G.; Berzin, T.M.; Kim, S.; Song, P.; Kuo-LeBlanc, V.; Rodriguez-Wolf, M.; Baird, A.; Johanson, C.E. Human choroid plexus growth factors: What are the implications for CSF dynamics in Alzheimer’s disease? Exp. Neurol. 2001, 167, 40–47. [Google Scholar] [CrossRef] [PubMed]
  15. Bauer, H.-C.; Traweger, A.; Bauer, H. 1—Proteins of the tight junction in the blood-brain barrier. In Blood-Spinal Cord and Brain Barriers in Health and Disease; Sharma, H.S., Westman, J., Eds.; Academic Press: San Diego, CA, USA, 2004; pp. 1–10. [Google Scholar]
  16. Zheng, W.; Zhao, Q. The blood-CSF barrier in culture: Development of a primary culture and transepithelial transport model from choroidal epithelial cells. Methods Mol. Biol. 2002, 188, 99–114. [Google Scholar] [CrossRef] [PubMed]
  17. Nilsson, C.; Lindvall-Axelsson, M.; Owman, C. Neuroendocrine regulatory mechanisms in the choroid plexus-cerebrospinal fluid system. Brain Res. Brain Res. Rev. 1992, 17, 109–138. [Google Scholar] [CrossRef]
  18. Ueno, M.; Chiba, Y.; Murakami, R.; Matsumoto, K.; Kawauchi, M.; Fujihara, R. Blood-brain barrier and blood-cerebrospinal fluid barrier in normal and pathological conditions. Brain Tumor Pathol. 2016, 33, 89–96. [Google Scholar] [CrossRef]
  19. Redzic, Z.B.; Segal, M.B. The structure of the choroid plexus and the physiology of the choroid plexus epithelium. Adv. Drug Deliv. Rev. 2004, 56, 1695–1716. [Google Scholar] [CrossRef]
  20. Saunders, N.R.; Daneman, R.; Dziegielewska, K.M.; Liddelow, S.A. Transporters of the blood-brain and blood—CSF interfaces in development and in the adult. Mol. Asp. Med. 2013, 34, 742–752. [Google Scholar] [CrossRef]
  21. Wright, E.M. Effect of bicarbonate and other buffers on choroid plexus Na+/K+ pump. Biochim. Biophys. Acta 1977, 468, 486–489. [Google Scholar] [CrossRef]
  22. Zeuthen, T.; Wright, E.M. An electrogenic Na+/K+ pump in the choroid plexus. Biochim. Biophys. Acta 1978, 511, 517–522. [Google Scholar] [CrossRef]
  23. Barbuskaite, D.; Damkier, H.; Praetorius, J. Acid/base transporters in CSF secretion and pH regulation. In Role of the Choroid Plexus in Health and Disease; Praetorius, J., Blazer-Yost, B., Damkier, H., Eds.; Springer: New York, NY, USA, 2020; pp. 149–171. [Google Scholar]
  24. Shen, H.; Smith, D.E.; Keep, R.F.; Xiang, J.; Brosius, F.C., 3rd. Targeted disruption of the PEPT2 gene markedly reduces dipeptide uptake in choroid plexus. J. Biol. Chem. 2003, 278, 4786–4791. [Google Scholar] [CrossRef] [PubMed]
  25. Wang, X.; Li, G.J.; Zheng, W. Upregulation of DMT1 expression in choroidal epithelia of the blood—CSF barrier following manganese exposure in vitro. Brain Res. 2006, 1097, 1–10. [Google Scholar] [CrossRef]
  26. Choi, B.S.; Zheng, W. Copper transport to the brain by the blood-brain barrier and blood-CSF barrier. Brain Res. 2009, 1248, 14–21. [Google Scholar] [CrossRef] [PubMed]
  27. Zheng, W.; Monnot, A.D. Regulation of brain iron and copper homeostasis by brain barrier systems: Implication in neurodegenerative diseases. Pharmacol. Ther. 2012, 133, 177–188. [Google Scholar] [CrossRef] [PubMed]
  28. Sun, A.; Wang, J. Choroid plexus and drug removal mechanisms. AAPS J. 2021, 23, 61. [Google Scholar] [CrossRef] [PubMed]
  29. Ghersi-Egea, J.F.; Strazielle, N. Choroid plexus transporters for drugs and other xenobiotics. J. Drug Target. 2002, 10, 353–357. [Google Scholar] [CrossRef]
  30. Hladky, S.B.; Barrand, M.A. Fluid and ion transfer across the blood-brain and blood-cerebrospinal fluid barriers; a comparative account of mechanisms and roles. Fluids Barriers CNS 2016, 13, 19. [Google Scholar] [CrossRef]
  31. Gao, B.; Meier, P.J. Organic anion transport across the choroid plexus. Microsc. Res. Tech. 2001, 52, 60–64. [Google Scholar] [CrossRef]
  32. Morris, M.E.; Rodriguez-Cruz, V.; Felmlee, M.A. SLC and ABC transporters: Expression, localization, and species differences at the blood-brain and the blood-cerebrospinal fluid barriers. AAPS J. 2017, 19, 1317–1331. [Google Scholar] [CrossRef]
  33. Angelow, S.; Wegener, J.; Galla, H.-J. 4—Transport and permeability characteristics of the blood-cerebrospinal fluid barrier in vitro. In Blood-Spinal Cord and Brain Barriers in Health and Disease; Sharma, H.S., Westman, J., Eds.; Academic Press: San Diego, CA, USA, 2004; pp. 33–45. [Google Scholar]
  34. Strazielle, N.; Khuth, S.T.; Ghersi-Egea, J.F. Detoxification systems, passive and specific transport for drugs at the blood-CSF barrier in normal and pathological situations. Adv. Drug Deliv. Rev. 2004, 56, 1717–1740. [Google Scholar] [CrossRef]
  35. Miller, D.S. ABC transporter regulation by signaling at the blood-brain barrier: Relevance to pharmacology. Adv. Pharmacol. 2014, 71, 1–24. [Google Scholar] [CrossRef] [PubMed]
  36. Rao, V.V.; Dahlheimer, J.L.; Bardgett, M.E.; Snyder, A.Z.; Finch, R.A.; Sartorelli, A.C.; Piwnica-Worms, D. Choroid plexus epithelial expression of MDR1 P glycoprotein and multidrug resistance-associated protein contribute to the blood-cerebrospinal-fluid drug-permeability barrier. Proc. Natl. Acad. Sci. USA 1999, 96, 3900–3905. [Google Scholar] [CrossRef] [PubMed]
  37. Ghersi-Egea, J.F.; Mönkkönen, K.S.; Schmitt, C.; Honnorat, J.; Fèvre-Montange, M.; Strazielle, N. Blood-brain interfaces and cerebral drug bioavailability. Rev. Neurol. 2009, 165, 1029–1038. [Google Scholar] [CrossRef] [PubMed]
  38. Begley, D.J. 9—Efflux mechanisms in the central nervous system: A powerful influence on drug distribution within the brain. In Blood-Spinal Cord and Brain Barriers in Health and Disease; Sharma, H.S., Westman, J., Eds.; Academic Press: San Diego, CA, USA, 2004; pp. 83–97. [Google Scholar]
  39. Bernd, A.; Ott, M.; Ishikawa, H.; Schroten, H.; Schwerk, C.; Fricker, G. Characterization of efflux transport proteins of the human choroid plexus papilloma cell line HIBCPP, a functional in vitro model of the blood-cerebrospinal fluid barrier. Pharm. Res. 2015, 32, 2973–2982. [Google Scholar] [CrossRef]
  40. Graff, C.L.; Pollack, G.M. Drug transport at the blood-brain barrier and the choroid plexus. Curr. Drug Metab. 2004, 5, 95–108. [Google Scholar] [CrossRef]
  41. Wang, Q.; Zuo, Z. Impact of transporters and enzymes from blood-cerebrospinal fluid barrier and brain parenchyma on CNS drug uptake. Expert Opin. Drug Metab. Toxicol. 2018, 14, 961–972. [Google Scholar] [CrossRef]
  42. Spector, R. Nature and consequences of mammalian brain and CSF efflux transporters: Four decades of progress. J. Neurochem. 2010, 112, 13–23. [Google Scholar] [CrossRef]
  43. De Lange, E.C. Potential role of ABC transporters as a detoxification system at the blood—CSF barrier. Adv. Drug Deliv. Rev. 2004, 56, 1793–1809. [Google Scholar] [CrossRef]
  44. Strazielle, N.; Ghersi-Egea, J.F. Demonstration of a coupled metabolism—Efflux process at the choroid plexus as a mechanism of brain protection toward xenobiotics. J. Neurosci. 1999, 19, 6275–6289. [Google Scholar] [CrossRef]
  45. Gomez-Zepeda, D.; Taghi, M.; Scherrmann, J.M.; Decleves, X.; Menet, M.C. ABC transporters at the blood-brain interfaces, their study models, and drug delivery implications in gliomas. Pharmaceutics 2019, 12, 20. [Google Scholar] [CrossRef]
  46. Ghersi-Egea, J.F.; Leininger-Muller, B.; Cecchelli, R.; Fenstermacher, J.D. Blood-brain interfaces: Relevance to cerebral drug metabolism. Toxicol. Lett. 1995, 82–83, 645–653. [Google Scholar] [CrossRef]
  47. Ghersi-Egea, J.F.; Strazielle, N.; Belin, M.F. Neuroprotective and detoxifying mechanisms at the blood-brain interfaces. In Blood—Brain Barrier: Drug Delivery and Brain Pathology; Kobiler, D., Lustig, S., Shapira, S., Eds.; Springer: Boston, MA, USA, 2001; pp. 19–25. [Google Scholar]
  48. Declèves, X.; Strazielle, N.; Scherrmann, J.-M.; Ghersi-Egea, J.-F. Drug metabolism at the blood–brain and blood–CSF barriers. In Drug Delivery to the Brain: Physiological Concepts, Methodologies and Approaches; Hammarlund-Udenaes, M., de Lange, E.C.M., Thorne, R.G., Eds.; Springer: New York, NY, USA, 2014; pp. 101–124. [Google Scholar]
  49. Ghersi-Egea, J.F.; Strazielle, N. Brain drug delivery, drug metabolism, and multidrug resistance at the choroid plexus. Microsc. Res. Tech. 2001, 52, 83–88. [Google Scholar] [CrossRef]
  50. Ghersi-Egea, J.F.; Strazielle, N.; Murat, A.; Jouvet, A.; Buénerd, A.; Belin, M.F. Brain protection at the blood-cerebrospinal fluid interface involves a glutathione-dependent metabolic barrier mechanism. J. Cereb. Blood Flow Metab. 2006, 26, 1165–1175. [Google Scholar] [CrossRef]
  51. Niehof, M.; Borlak, J. Expression of HNF4alpha in the human and rat choroid plexus: Implications for drug transport across the blood-cerebrospinal-fluid (CSF) barrier. BMC Mol. Biol. 2009, 10, 68. [Google Scholar] [CrossRef]
  52. Tan, H.Y.; Cho, H.; Lee, L.P. Human mini-brain models. Nat. Biomed. Eng. 2021, 5, 11–25. [Google Scholar] [CrossRef] [PubMed]
  53. Larsen, J.B.; Taebnia, N.; Dolatshahi-Pirouz, A.; Eriksen, A.Z.; Hjørringgaard, C.; Kristensen, K.; Larsen, N.W.; Larsen, N.B.; Marie, R.; Mündler, A.K.; et al. Imaging therapeutic peptide transport across intestinal barriers. RSC Chem. Biol. 2021, 2, 1115–1143. [Google Scholar] [CrossRef]
  54. Dinner, S.; Borkowski, J.; Stump-Guthier, C.; Ishikawa, H.; Tenenbaum, T.; Schroten, H.; Schwerk, C. A choroid plexus epithelial cell-based model of the human blood-cerebrospinal fluid barrier to study bacterial infection from the basolateral side. J. Vis. Exp. 2016, 11, 54061. [Google Scholar] [CrossRef]
  55. Kara, A.; Ozturk, N.; Vural, I. Chapter 8—In vitro CNS models. In Nanotechnology Methods for Neurological Diseases and Brain Tumors; Gürsoy-Özdemir, Y., Bozdağ-Pehlivan, S., Sekerdag, E., Eds.; Academic Press: Cambridge, MA, USA, 2017; pp. 151–185. [Google Scholar]
  56. Bhalerao, A.; Sivandzade, F.; Archie, S.R.; Chowdhury, E.A.; Noorani, B.; Cucullo, L. In vitro modeling of the neurovascular unit: Advances in the field. Fluids Barriers CNS 2020, 17, 22. [Google Scholar] [CrossRef]
  57. Mayer, S.E.; Sanders-Bush, E. Sodium-dependent antiporters in choroid plexus epithelial cultures from rabbit. J. Neurochem. 1993, 60, 1308–1316. [Google Scholar] [CrossRef]
  58. Haselbach, M.; Wegener, J.; Decker, S.; Engelbertz, C.; Galla, H.J. Porcine choroid plexus epithelial cells in culture: Regulation of barrier properties and transport processes. Microsc. Res. Tech. 2001, 52, 137–152. [Google Scholar] [CrossRef]
  59. Strazielle, N.; Ghersi-Egea, J.-F. In vitro models of the blood-cerebrospinal fluid barrier and their use in neurotoxicological research. In Cell Culture Techniques. Neuromethods; Aschner, M., Suñol, C., Bal-Price, A., Eds.; Humana Press: Totowa, NJ, USA, 2011; Volume 56. [Google Scholar]
  60. Dehouck, M.P.; Méresse, S.; Delorme, P.; Fruchart, J.C.; Cecchelli, R. An easier, reproducible, and mass-production method to study the blood-brain barrier in vitro. J. Neurochem. 1990, 54, 1798–1801. [Google Scholar] [CrossRef] [PubMed]
  61. Gaillard, P.J.; Voorwinden, L.H.; Nielsen, J.L.; Ivanov, A.; Atsumi, R.; Engman, H.; Ringbom, C.; de Boer, A.G.; Breimer, D.D. Establishment and functional characterization of an in vitro model of the blood-brain barrier, comprising a co-culture of brain capillary endothelial cells and astrocytes. Eur. J. Pharm. Sci. 2001, 12, 215–222. [Google Scholar] [CrossRef]
  62. Reichel, A.; Begley, D.J.; Abbott, N.J. An overview of in vitro techniques for blood-brain barrier studies. Methods Mol. Med. 2003, 89, 307–324. [Google Scholar] [CrossRef] [PubMed]
  63. Di Marco, A.; Gonzalez Paz, O.; Fini, I.; Vignone, D.; Cellucci, A.; Battista, M.R.; Auciello, G.; Orsatti, L.; Zini, M.; Monteagudo, E.; et al. Application of an in vitro blood-brain barrier model in the selection of experimental drug candidates for the treatment of Huntington’s disease. Mol. Pharm. 2019, 16, 2069–2082. [Google Scholar] [CrossRef]
  64. Deli, M.A.; Abrahám, C.S.; Kataoka, Y.; Niwa, M. Permeability studies on in vitro blood-brain barrier models: Physiology, pathology, and pharmacology. Cell Mol. Neurobiol. 2005, 25, 59–127. [Google Scholar] [CrossRef]
  65. Muranyi, W.; Schwerk, C.; Herold, R.; Stump-Guthier, C.; Lampe, M.; Fallier-Becker, P.; Weiß, C.; Sticht, C.; Ishikawa, H.; Schroten, H. Immortalized human choroid plexus endothelial cells enable an advanced endothelial-epithelial two-cell type in vitro model of the choroid plexus. iScience 2022, 25, 104383. [Google Scholar] [CrossRef]
  66. Carloni, S.; Bertocchi, A.; Mancinelli, S.; Bellini, M.; Erreni, M.; Borreca, A.; Braga, D.; Giugliano, S.; Mozzarelli, A.M.; Manganaro, D.; et al. Identification of a choroid plexus vascular barrier closing during intestinal inflammation. Science 2021, 374, 439–448. [Google Scholar] [CrossRef]
  67. von Maydell, D.; Jorfi, M. A synergistic engineering approach to build human brain spheroids. Methods Mol. Biol. 2021, 2258, 151–169. [Google Scholar] [CrossRef]
  68. Logan, S.; Arzua, T.; Canfield, S.G.; Seminary, E.R.; Sison, S.L.; Ebert, A.D.; Bai, X. Studying human neurological disorders using induced pluripotent stem cells: From 2D monolayer to 3D organoid and blood brain barrier models. Compr. Physiol. 2019, 9, 565–611. [Google Scholar] [CrossRef]
  69. Ahn, S.I.; Sei, Y.J.; Park, H.J.; Kim, J.; Ryu, Y.; Choi, J.J.; Sung, H.J.; MacDonald, T.J.; Levey, A.I.; Kim, Y. Microengineered human blood-brain barrier platform for understanding nanoparticle transport mechanisms. Nat. Commun. 2020, 11, 175. [Google Scholar] [CrossRef]
  70. Katt, M.E.; Linville, R.M.; Mayo, L.N.; Xu, Z.S.; Searson, P.C. Functional brain-specific microvessels from iPSC-derived human brain microvascular endothelial cells: The role of matrix composition on monolayer formation. Fluids Barriers CNS 2018, 15, 7. [Google Scholar] [CrossRef]
  71. Potjewyd, G.; Kellett, K.A.B.; Hooper, N.M. 3D hydrogel models of the neurovascular unit to investigate blood-brain barrier dysfunction. Neuronal Signal. 2021, 5, Ns20210027. [Google Scholar] [CrossRef] [PubMed]
  72. Petersen, N.; Torz, L.; Jensen, K.H.R.; Hjortø, G.M.; Spiess, K.; Rosenkilde, M.M. Three-dimensional explant platform for studies on choroid plexus epithelium. Front. Cell. Neurosci. 2020, 14, 108. [Google Scholar] [CrossRef] [PubMed]
  73. Ulloa, V.; Saldivia, N.; Ferrada, L.; Salazar, K.; Martínez, F.; Silva-Alvarez, C.; Magdalena, R.; Oviedo, M.J.; Montecinos, H.; Torres-Vergara, P.; et al. Basal sodium-dependent vitamin C transporter 2 polarization in choroid plexus explant cells in normal or scorbutic conditions. Sci. Rep. 2019, 9, 14422. [Google Scholar] [CrossRef]
  74. Pellegrini, L.; Lancaster, M.A. Breaking the barrier: In vitro models to study choroid plexus development. Curr. Opin. Cell. Biol. 2021, 73, 41–49. [Google Scholar] [CrossRef] [PubMed]
  75. Dragunow, M.; Feng, S.; Rustenhoven, J.; Curtis, M.; Faull, R. Studying human brain inflammation in leptomeningeal and choroid plexus explant cultures. Neurochem. Res. 2016, 41, 579–588. [Google Scholar] [CrossRef] [PubMed]
  76. Gonzalez, A.M.; Leadbeater, W.E.; Burg, M.; Sims, K.; Terasaki, T.; Johanson, C.E.; Stopa, E.G.; Eliceiri, B.P.; Baird, A. Targeting choroid plexus epithelia and ventricular ependyma for drug delivery to the central nervous system. BMC Neurosci. 2011, 12, 4. [Google Scholar] [CrossRef]
  77. Huch, M.; Knoblich, J.A.; Lutolf, M.P.; Martinez-Arias, A. The hope and the hype of organoid research. Development 2017, 144, 938–941. [Google Scholar] [CrossRef]
  78. Bergmann, S.; Lawler, S.E.; Qu, Y.; Fadzen, C.M.; Wolfe, J.M.; Regan, M.S.; Pentelute, B.L.; Agar, N.Y.R.; Cho, C.F. Blood-brain-barrier organoids for investigating the permeability of CNS therapeutics. Nat. Protoc. 2018, 13, 2827–2843. [Google Scholar] [CrossRef]
  79. Simonneau, C.; Duschmalé, M.; Gavrilov, A.; Brandenberg, N.; Hoehnel, S.; Ceroni, C.; Lassalle, E.; Kassianidou, E.; Knoetgen, H.; Niewoehner, J.; et al. Investigating receptor-mediated antibody transcytosis using blood-brain barrier organoid arrays. Fluids Barriers CNS 2021, 18, 43. [Google Scholar] [CrossRef]
  80. Jacob, F.; Pather, S.R.; Huang, W.K.; Zhang, F.; Wong, S.Z.H.; Zhou, H.; Cubitt, B.; Fan, W.; Chen, C.Z.; Xu, M.; et al. Human pluripotent stem cell-derived neural cells and brain organoids reveal SARS-CoV-2 neurotropism predominates in choroid plexus epithelium. Cell Stem Cell 2020, 27, 937–950.e939. [Google Scholar] [CrossRef] [PubMed]
  81. Pellegrini, L.; Bonfio, C.; Chadwick, J.; Begum, F.; Skehel, M.; Lancaster, M.A. Human CNS barrier-forming organoids with cerebrospinal fluid production. Science 2020, 369, eaaz5626. [Google Scholar] [CrossRef] [PubMed]
  82. Pellegrini, L.; Albecka, A.; Mallery, D.L.; Kellner, M.J.; Paul, D.; Carter, A.P.; James, L.C.; Lancaster, M.A. SARS-CoV-2 infects the brain choroid plexus and disrupts the blood-CSF barrier in human brain organoids. Cell Stem Cell 2020, 27, 951–961.e955. [Google Scholar] [CrossRef] [PubMed]
  83. Xu, H.; Lehtinen, M.K. Choroid plexus organoids: Harnessing CSF gatekeepers for brain therapeutics. Cell Stem Cell 2020, 27, 191–192. [Google Scholar] [CrossRef]
  84. Tang, M.; Rich, J.N.; Chen, S. Biomaterials and 3D bioprinting strategies to model glioblastoma and the blood-brain barrier. Adv. Mater. 2021, 33, e2004776. [Google Scholar] [CrossRef]
  85. Marino, A.; Tricinci, O.; Battaglini, M.; Filippeschi, C.; Mattoli, V.; Sinibaldi, E.; Ciofani, G. A 3D real-scale, biomimetic, and biohybrid model of the blood-brain barrier fabricated through two-photon lithography. Small 2018, 14, 1702959. [Google Scholar] [CrossRef]
  86. Kim, J.A.; Kim, H.N.; Im, S.K.; Chung, S.; Kang, J.Y.; Choi, N. Collagen-based brain microvasculature model in vitro using three-dimensional printed template. Biomicrofluidics 2015, 9, 024115. [Google Scholar] [CrossRef]
  87. Qi, D.; Wu, S.; Lin, H.; Kuss, M.A.; Lei, Y.; Krasnoslobodtsev, A.; Ahmed, S.; Zhang, C.; Kim, H.J.; Jiang, P.; et al. Establishment of a human iPSC- and nanofiber-based microphysiological blood-brain barrier system. ACS Appl. Mater. Interfaces 2018, 10, 21825–21835. [Google Scholar] [CrossRef]
  88. Galpayage Dona, K.N.U.; Hale, J.F.; Salako, T.; Anandanatarajan, A.; Tran, K.A.; DeOre, B.J.; Galie, P.A.; Ramirez, S.H.; Andrews, A.M. The use of tissue engineering to fabricate perfusable 3D brain microvessels in vitro. Front. Physiol. 2021, 12, 715431. [Google Scholar] [CrossRef]
  89. Sivandzade, F.; Cucullo, L. In-vitro blood-brain barrier modeling: A review of modern and fast-advancing technologies. J. Cereb Blood Flow Metab. 2018, 38, 1667–1681. [Google Scholar] [CrossRef]
  90. Wang, Y.I.; Abaci, H.E.; Shuler, M.L. Microfluidic blood-brain barrier model provides in vivo-like barrier properties for drug permeability screening. Biotechnol. Bioeng. 2017, 114, 184–194. [Google Scholar] [CrossRef] [PubMed]
  91. Musafargani, S.; Mishra, S.; Gulyás, M.; Mahalakshmi, P.; Archunan, G.; Padmanabhan, P.; Gulyás, B. Blood brain barrier: A tissue engineered microfluidic chip. J. Neurosci. Methods 2020, 331, 108525. [Google Scholar] [CrossRef] [PubMed]
  92. Oddo, A.; Peng, B.; Tong, Z.; Wei, Y.; Tong, W.Y.; Thissen, H.; Voelcker, N.H. Advances in microfluidic blood-brain barrier (BBB) models. Trends Biotechnol. 2019, 37, 1295–1314. [Google Scholar] [CrossRef] [PubMed]
  93. Bagchi, S.; Chhibber, T.; Lahooti, B.; Verma, A.; Borse, V.; Jayant, R.D. In-vitro blood-brain barrier models for drug screening and permeation studies: An overview. Drug Des. Dev. Ther. 2019, 13, 3591–3605. [Google Scholar] [CrossRef]
  94. Yu, F.; Selva Kumar, N.D.; Choudhury, D.; Foo, L.C.; Ng, S.H. Microfluidic platforms for modeling biological barriers in the circulatory system. Drug Discov. Today 2018, 23, 815–829. [Google Scholar] [CrossRef]
  95. Campisi, M.; Lim, S.H.; Chiono, V.; Kamm, R.D. 3D self-organized human blood-brain barrier in a microfluidic chip. Methods Mol. Biol. 2021, 2258, 205–219. [Google Scholar] [CrossRef]
  96. Vatine, G.D.; Barrile, R.; Workman, M.J.; Sances, S.; Barriga, B.K.; Rahnama, M.; Barthakur, S.; Kasendra, M.; Lucchesi, C.; Kerns, J.; et al. Human iPSC-derived blood-brain barrier chips enable disease modeling and personalized medicine applications. Cell Stem Cell 2019, 24, 995–1005.e1006. [Google Scholar] [CrossRef]
  97. Campisi, M.; Shin, Y.; Osaki, T.; Hajal, C.; Chiono, V.; Kamm, R.D. 3D self-organized microvascular model of the human blood-brain barrier with endothelial cells, pericytes and astrocytes. Biomaterials 2018, 180, 117–129. [Google Scholar] [CrossRef]
  98. Yeon, J.H.; Na, D.; Choi, K.; Ryu, S.W.; Choi, C.; Park, J.K. Reliable permeability assay system in a microfluidic device mimicking cerebral vasculatures. Biomed. Microdevices 2012, 14, 1141–1148. [Google Scholar] [CrossRef]
  99. Adriani, G.; Ma, D.; Pavesi, A.; Kamm, R.D.; Goh, E.L. A 3D neurovascular microfluidic model consisting of neurons, astrocytes and cerebral endothelial cells as a blood-brain barrier. Lab Chip 2017, 17, 448–459. [Google Scholar] [CrossRef]
  100. Peng, B.; Tong, Z.; Tong, W.Y.; Pasic, P.J.; Oddo, A.; Dai, Y.; Luo, M.; Frescene, J.; Welch, N.G.; Easton, C.D.; et al. In situ surface modification of microfluidic blood-brain-barriers for improved screening of small molecules and nanoparticles. ACS Appl. Mater. Interfaces 2020, 12, 56753–56766. [Google Scholar] [CrossRef] [PubMed]
  101. Thomas, T.; Stadler, E.; Dziadek, M. Effects of the extracellular matrix on fetal choroid plexus epithelial cells: Changes in morphology and multicellular organization do not affect gene expression. Exp. Cell Res. 1992, 203, 198–213. [Google Scholar] [CrossRef]
  102. Aerts, J.; Vandenbroucke, R.E.; Dera, R.; Balusu, S.; Van Wonterghem, E.; Moons, L.; Libert, C.; Dehaen, W.; Arckens, L. Synthesis and validation of a hydroxypyrone-based, potent, and specific matrix metalloproteinase-12 inhibitor with anti-inflammatory activity in vitro and in vivo. Mediat. Inflamm. 2015, 2015, 510679. [Google Scholar] [CrossRef]
  103. Barkho, B.Z.; Monuki, E.S. Proliferation of cultured mouse choroid plexus epithelial cells. PLoS ONE 2015, 10, e0121738. [Google Scholar] [CrossRef] [PubMed]
  104. Péraldi-Roux, S.; Nguyen-Than Dao, B.; Hirn, M.; Gabrion, J. Choroidal ependymocytes in culture: Expression of markers of polarity and function. Int. J. Dev. Neurosci. 1990, 8, 575–588. [Google Scholar] [CrossRef]
  105. Zheng, W.; Zhao, Q.; Graziano, J.H. Primary culture of choroidal epithelial cells: Characterization of an in vitro model of blood-CSF barrier. Vitr. Cell Dev. Biol. Anim. 1998, 34, 40–45. [Google Scholar] [CrossRef]
  106. Watson, J.A.; Elliott, A.C.; Brown, P.D. Serotonin elevates intracellular Ca2+ in rat choroid plexus epithelial cells by acting on 5-HT2C receptors. Cell Calcium 1995, 17, 120–128. [Google Scholar] [CrossRef]
  107. Batisson, M.; Strazielle, N.; Hejmadi, M.; Thomas, D.; Ghersi-Egea, J.F.; Etienne, J.; Vandenesch, F.; Lina, G. Toxic shock syndrome toxin-1 challenges the neuroprotective functions of the choroidal epithelium and induces neurotoxicity. J. Infect. Dis. 2006, 194, 341–349. [Google Scholar] [CrossRef]
  108. Villalobos, A.R.; Parmelee, J.T.; Pritchard, J.B. Functional characterization of choroid plexus epithelial cells in primary culture. J. Pharmacol. Exp. Ther. 1997, 282, 1109–1116. [Google Scholar]
  109. Hakvoort, A.; Haselbach, M.; Galla, H.J. Active transport properties of porcine choroid plexus cells in culture. Brain Res. 1998, 795, 247–256. [Google Scholar] [CrossRef]
  110. Nilsson, C.; Fahrenkrug, J.; Lindvall-Axelsson, M.; Owman, C. Epithelial cells purified from choroid plexus have receptors for vasoactive intestinal polypeptide. Brain Res. 1991, 542, 241–247. [Google Scholar] [CrossRef]
  111. Angelow, S.; Zeni, P.; Galla, H.J. Usefulness and limitation of primary cultured porcine choroid plexus epithelial cells as an in vitro model to study drug transport at the blood—CSF barrier. Adv. Drug Deliv. Rev. 2004, 56, 1859–1873. [Google Scholar] [CrossRef] [PubMed]
  112. Angelow, S.; Haselbach, M.; Galla, H.J. Functional characterisation of the active ascorbic acid transport into cerebrospinal fluid using primary cultured choroid plexus cells. Brain Res. 2003, 988, 105–113. [Google Scholar] [CrossRef]
  113. Angelow, S.; Zeni, P.; Höhn, B.; Galla, H.J. Phorbol ester induced short- and long-term permeabilization of the blood—CSF barrier in vitro. Brain Res. 2005, 1063, 168–179. [Google Scholar] [CrossRef]
  114. Tenenbaum, T.; Essmann, F.; Adam, R.; Seibt, A.; Jänicke, R.U.; Novotny, G.E.; Galla, H.J.; Schroten, H. Cell death, caspase activation, and HMGB1 release of porcine choroid plexus epithelial cells during Streptococcus suis infection in vitro. Brain Res. 2006, 1100, 1–12. [Google Scholar] [CrossRef]
  115. Baehr, C.; Reichel, V.; Fricker, G. Choroid plexus epithelial monolayers—A cell culture model from porcine brain. Cerebrospinal Fluid Res. 2006, 3, 13. [Google Scholar] [CrossRef]
  116. Gath, U.; Hakvoort, A.; Wegener, J.; Decker, S.; Galla, H.J. Porcine choroid plexus cells in culture: Expression of polarized phenotype, maintenance of barrier properties and apical secretion of CSF-components. Eur. J. Cell Biol. 1997, 74, 68–78. [Google Scholar]
  117. Hakvoort, A.; Haselbach, M.; Wegener, J.; Hoheisel, D.; Galla, H.J. The polarity of choroid plexus epithelial cells in vitro is improved in serum-free medium. J. Neurochem. 1998, 71, 1141–1150. [Google Scholar] [CrossRef]
  118. Crook, R.B.; Kasagami, H.; Prusiner, S.B. Culture and characterization of epithelial cells from bovine choroid plexus. J. Neurochem. 1981, 37, 845–854. [Google Scholar] [CrossRef]
  119. Crook, R.B.; Prusiner, S.B. Vasoactive intestinal peptide stimulates cyclic AMP metabolism in choroid plexus epithelial cells. Brain Res. 1986, 384, 138–144. [Google Scholar] [CrossRef]
  120. Holm, N.R.; Hansen, L.B.; Nilsson, C.; Gammeltoft, S. Gene expression and secretion of insulin-like growth factor-II and insulin-like growth factor binding protein-2 from cultured sheep choroid plexus epithelial cells. Brain Res. Mol. Brain Res. 1994, 21, 67–74. [Google Scholar] [CrossRef]
  121. Salvatori, D.; Vincenzetti, S.; Maury, G.; Gosselin, G.; Gaubert, G.; Vita, A. Maedi-visna virus, a model for in vitro testing of potential anti-HIV drugs. Comp. Immunol. Microbiol. Infect. Dis. 2001, 24, 113–122. [Google Scholar] [CrossRef]
  122. Ishiwata, I.; Ishiwata, C.; Ishiwata, E.; Sato, Y.; Kiguchi, K.; Tachibana, T.; Hashimoto, H.; Ishikawa, H. Establishment and characterization of a human malignant choroids plexus papilloma cell line (HIBCPP). Hum. Cell 2005, 18, 67–72. [Google Scholar] [CrossRef]
  123. Ramanathan, V.K.; Hui, A.C.; Brett, C.M.; Giacomini, K.M. Primary cell culture of the rabbit choroid plexus: An experimental system to investigate membrane transport. Pharm. Res. 1996, 13, 952–956. [Google Scholar] [CrossRef]
  124. Ramanathan, V.K.; Chung, S.J.; Giacomini, K.M.; Brett, C.M. Taurine transport in cultured choroid plexus. Pharm. Res. 1997, 14, 406–409. [Google Scholar] [CrossRef]
  125. Delery, E.C.; MacLean, A.G. Culture model for non-human primate choroid plexus. Front. Cell. Neurosci. 2019, 13, 396. [Google Scholar] [CrossRef] [PubMed]
  126. Greenwood, S.; Swetloff, A.; Wade, A.M.; Terasaki, T.; Ferretti, P. Fgf2 is expressed in human and murine embryonic choroid plexus and affects choroid plexus epithelial cell behaviour. Cerebrospinal Fluid Res. 2008, 5, 20. [Google Scholar] [CrossRef]
  127. Swetloff, A.; Ferretti, P. Changes in E2F5 intracellular localization in mouse and human choroid plexus epithelium with development. Int. J. Dev. Biol. 2005, 49, 859–865. [Google Scholar] [CrossRef] [PubMed]
  128. Erb, U.; Schwerk, C.; Schroten, H.; Karremann, M. Review of functional in vitro models of the blood-cerebrospinal fluid barrier in leukaemia research. J. Neurosci. Methods 2020, 329, 108478. [Google Scholar] [CrossRef]
  129. Madea, B.; Musshoff, F. Postmortem biochemistry. Forensic Sci. Int. 2007, 165, 165–171. [Google Scholar] [CrossRef]
  130. Sauer, S.W.; Opp, S.; Mahringer, A.; Kamiński, M.M.; Thiel, C.; Okun, J.G.; Fricker, G.; Morath, M.A.; Kölker, S. Glutaric aciduria type I and methylmalonic aciduria: Simulation of cerebral import and export of accumulating neurotoxic dicarboxylic acids in in vitro models of the blood-brain barrier and the choroid plexus. Biochim. Biophys. Acta 2010, 1802, 552–560. [Google Scholar] [CrossRef] [PubMed]
  131. Kuplennik, N.; Lang, K.; Steinfeld, R.; Sosnik, A. Folate receptor α-modified nanoparticles for targeting of the central nervous system. ACS Appl. Mater. Interfaces 2019, 11, 39633–39647. [Google Scholar] [CrossRef] [PubMed]
  132. Strazielle, N.; Ghersi-Egea, J.-F. In vitro investigation of the blood-cerebrospinal fluid barrier properties: Primary cultures and immortalized cell lines of the choroidal epithelium. In The Blood-Cerebrospinal Fluid Barrier; Zheng, W., Chodobski, A., Eds.; CRC Press: Boca Raton, FL, USA, 2005. [Google Scholar]
  133. Lallai, V.; Ahmed, A.; Fowler, C.D. Method for primary epithelial cell culture from the rat choroid plexus. Bio Protoc. 2020, 10, e3532. [Google Scholar] [CrossRef]
  134. Kitazawa, T.; Hosoya, K.; Watanabe, M.; Takashima, T.; Ohtsuki, S.; Takanaga, H.; Ueda, M.; Yanai, N.; Obinata, M.; Terasaki, T. Characterization of the amino acid transport of new immortalized choroid plexus epithelial cell lines: A novel in vitro system for investigating transport functions at the blood-cerebrospinal fluid barrier. Pharm. Res. 2001, 18, 16–22. [Google Scholar] [CrossRef] [PubMed]
  135. Zheng, W.; Zhao, Q. Establishment and characterization of an immortalized Z310 choroidal epithelial cell line from murine choroid plexus. Brain Res. 2002, 958, 371–380. [Google Scholar] [CrossRef]
  136. Kläs, J.; Wolburg, H.; Terasaki, T.; Fricker, G.; Reichel, V. Characterization of immortalized choroid plexus epithelial cell lines for studies of transport processes across the blood-cerebrospinal fluid barrier. Cerebrospinal Fluid Res. 2010, 7, 11. [Google Scholar] [CrossRef]
  137. Fujiyoshi, M.; Ohtsuki, S.; Hori, S.; Tachikawa, M.; Terasaki, T. 24S-hydroxycholesterol induces cholesterol release from choroid plexus epithelial cells in an apical- and apoE isoform-dependent manner concomitantly with the induction of ABCA1 and ABCG1 expression. J. Neurochem. 2007, 100, 968–978. [Google Scholar] [CrossRef]
  138. Li, G.J.; Zhao, Q.; Zheng, W. Alteration at translational but not transcriptional level of transferrin receptor expression following manganese exposure at the blood-CSF barrier in vitro. Toxicol. Appl. Pharmacol. 2005, 205, 188–200. [Google Scholar] [CrossRef]
  139. Tachikawa, M.; Fujinawa, J.; Takahashi, M.; Kasai, Y.; Fukaya, M.; Sakai, K.; Yamazaki, M.; Tomi, M.; Watanabe, M.; Sakimura, K.; et al. Expression and possible role of creatine transporter in the brain and at the blood-cerebrospinal fluid barrier as a transporting protein of guanidinoacetate, an endogenous convulsant. J. Neurochem. 2008, 107, 768–778. [Google Scholar] [CrossRef]
  140. Shi, L.Z.; Li, G.J.; Wang, S.; Zheng, W. Use of Z310 cells as an in vitro blood-cerebrospinal fluid barrier model: Tight junction proteins and transport properties. Toxicol. In Vitro 2008, 22, 190–199. [Google Scholar] [CrossRef]
  141. Terasaki, T.; Hosoya, K. Conditionally immortalized cell lines as a new in vitro model for the study of barrier functions. Biol. Pharm. Bull. 2001, 24, 111–118. [Google Scholar] [CrossRef] [PubMed]
  142. Hosoya, K.; Hori, S.; Ohtsuki, S.; Terasaki, T. A new in vitro model for blood-cerebrospinal fluid barrier transport studies: An immortalized choroid plexus epithelial cell line derived from the tsA58 SV40 large T-antigen gene transgenic rat. Adv. Drug Deliv. Rev. 2004, 56, 1875–1885. [Google Scholar] [CrossRef] [PubMed]
  143. Schroten, M.; Hanisch, F.G.; Quednau, N.; Stump, C.; Riebe, R.; Lenk, M.; Wolburg, H.; Tenenbaum, T.; Schwerk, C. A novel porcine in vitro model of the blood-cerebrospinal fluid barrier with strong barrier function. PLoS ONE 2012, 7, e39835. [Google Scholar] [CrossRef] [PubMed]
  144. Enjoji, M.; Iwaki, T.; Hara, H.; Sakai, H.; Nawata, H.; Watanabe, T. Establishment and characterization of choroid plexus carcinoma cell lines: Connection between choroid plexus and immune systems. Jpn. J. Cancer Res. 1996, 87, 893–899. [Google Scholar] [CrossRef]
  145. Enjoji, M.; Iwaki, T.; Nawata, H.; Watanabe, T. IgH intronic enhancer element HE2 (μB) functions as a cis-activator in choroid plexus cells at the cellular level as well as in transgenic mice. J. Neurochem. 1995, 64, 961–966. [Google Scholar] [CrossRef] [PubMed]
  146. Schell, T.D.; Mylin, L.M.; Georgoff, I.; Teresky, A.K.; Levine, A.J.; Tevethia, S.S. Cytotoxic T-lymphocyte epitope immunodominance in the control of choroid plexus tumors in simian virus 40 large T antigen transgenic mice. J. Virol. 1999, 73, 5981–5993. [Google Scholar] [CrossRef]
  147. Prasad, P.; Vasquez, H.; Das, C.M.; Gopalakrishnan, V.; Wolff, J.E. Histone acetylation resulting in resistance to methotrexate in choroid plexus cells. J. Neurooncol. 2009, 91, 279–286. [Google Scholar] [CrossRef]
  148. Lauer, A.N.; März, M.; Meyer, S.; Meurer, M.; de Buhr, N.; Borkowski, J.; Weiß, C.; Schroten, H.; Schwerk, C. Optimized cultivation of porcine choroid plexus epithelial cells, a blood-cerebrospinal fluid barrier model, for studying granulocyte transmigration. Lab. Investig. 2019, 99, 1245–1255. [Google Scholar] [CrossRef]
  149. Thörnwall, M.; Chhajlani, V.; Le Grevès, P.; Nyberg, F. Detection of growth hormone receptor mRNA in an ovine choroid plexus epithelium cell line. Biochem. Biophys. Res. Commun. 1995, 217, 349–353. [Google Scholar] [CrossRef]
  150. Oelschlegel, A.M.; Geissen, M.; Lenk, M.; Riebe, R.; Angermann, M.; Schatzl, H.; Groschup, M.H. A bovine cell line that can be infected by natural sheep scrapie prions. PLoS ONE 2015, 10, e0117154. [Google Scholar] [CrossRef]
  151. Albert, O.; Ancellin, N.; Preisser, L.; Morel, A.; Corman, B. Serotonin, bradykinin and endothelin signalling in a sheep choroid plexus cell line. Life Sci. 1999, 64, 859–867. [Google Scholar] [CrossRef]
  152. Merino, B.; Díez-Fernández, C.; Ruiz-Gayo, M.; Somoza, B. Choroid plexus epithelial cells co-express the long and short form of the leptin receptor. Neurosci. Lett. 2006, 393, 269–272. [Google Scholar] [CrossRef] [PubMed]
  153. Angelova, K.; Fralish, G.B.; Puett, D.; Narayan, P. Identification of conventional and novel endothelin receptors in sheep choroid plexus cells. Mol. Cell. Biochem. 1996, 159, 65–72. [Google Scholar] [CrossRef] [PubMed]
  154. Dickinson, K.E.; Baska, R.A.; Cohen, R.B.; Bryson, C.C.; Smith, M.A.; Schroeder, K.; Lodge, N.J. Identification of [3H]P1075 binding sites and P1075-activated K+ currents in ovine choroid plexus cells. Eur. J. Pharmacol. 1998, 345, 97–101. [Google Scholar] [CrossRef]
  155. Nakashima, N.; Goto, K.; Tsukidate, K.; Sobue, M.; Toida, M.; Takeuchi, J. Choroid plexus papilloma. Light and electron microscopic study. Virchows Arch. A Pathol Anat 1983, 400, 201–211. [Google Scholar] [CrossRef]
  156. Takahashi, K.; Satoh, F.; Hara, E.; Murakami, O.; Kumabe, T.; Tominaga, T.; Kayama, T.; Yoshimoto, T.; Shibahara, S. Production and secretion of adrenomedullin by cultured choroid plexus carcinoma cells. J. Neurochem. 1997, 68, 726–731. [Google Scholar] [CrossRef]
  157. Kumabe, T.; Tominaga, T.; Kondo, T.; Yoshimoto, T.; Kayama, T. Intraoperative radiation therapy and chemotherapy for huge choroid plexus carcinoma in an infant—Case report. Neurol. Med. Chir. 1996, 36, 179–184. [Google Scholar] [CrossRef]
  158. Szmydynger-Chodobska, J.; Pascale, C.L.; Pfeffer, A.N.; Coulter, C.; Chodobski, A. Expression of junctional proteins in choroid plexus epithelial cell lines: A comparative study. Cerebrospinal Fluid Res. 2007, 4, 11. [Google Scholar] [CrossRef]
  159. Callegari, E.; Malhotra, B.; Bungay, P.J.; Webster, R.; Fenner, K.S.; Kempshall, S.; LaPerle, J.L.; Michel, M.C.; Kay, G.G. A comprehensive non-clinical evaluation of the CNS penetration potential of antimuscarinic agents for the treatment of overactive bladder. Br. J. Clin. Pharmacol. 2011, 72, 235–246. [Google Scholar] [CrossRef]
  160. Fischer, H.; Senn, C.; Ullah, M.; Cantrill, C.; Schuler, F.; Yu, L. Calculation of an apical efflux ratio from P-plycoprotein (P-gp) in vitro transport experiments shows an improved correlation with in vivo cerebrospinal fluid measurements in rats: Impact on P-gp screening and compound optimization. J. Pharmacol. Exp. Ther. 2021, 376, 322–329. [Google Scholar] [CrossRef]
  161. Braun, A.; Hämmerle, S.; Suda, K.; Rothen-Rutishauser, B.; Günthert, M.; Krämer, S.D.; Wunderli-Allenspach, H. Cell cultures as tools in biopharmacy. Eur. J. Pharm. Sci. 2000, 11 (Suppl. 2), S51–S60. [Google Scholar] [CrossRef]
  162. Fischer, H.; Ullah, M.; de la Cruz, C.C.; Hunsaker, T.; Senn, C.; Wirz, T.; Wagner, B.; Draganov, D.; Vazvaei, F.; Donzelli, M.; et al. Entrectinib, a TRK/ROS1 inhibitor with anti-CNS tumor activity: Differentiation from other inhibitors in its class due to weak interaction with P-glycoprotein. Neuro Oncol. 2020, 22, 819–829. [Google Scholar] [CrossRef] [PubMed]
  163. Mangas-Sanjuan, V.; González-Álvarez, I.; González-Álvarez, M.; Casabó, V.G.; Bermejo, M. Innovative in vitro method to predict rate and extent of drug delivery to the brain across the blood-brain barrier. Mol. Pharm. 2013, 10, 3822–3831. [Google Scholar] [CrossRef]
  164. Terasaki, T.; Ohtsuki, S.; Hori, S.; Takanaga, H.; Nakashima, E.; Hosoya, K. New approaches to in vitro models of blood-brain barrier drug transport. Drug Discov. Today 2003, 8, 944–954. [Google Scholar] [CrossRef]
  165. Feng, B.; Doran, A.C.; Di, L.; West, M.A.; Osgood, S.M.; Mancuso, J.Y.; Shaffer, C.L.; Tremaine, L.; Liras, J. Prediction of human brain penetration of P-glycoprotein and breast cancer resistance protein substrates using in vitro transporter studies and animal models. J. Pharm. Sci. 2018, 107, 2225–2235. [Google Scholar] [CrossRef] [PubMed]
  166. Tachibana, K.; Hashimoto, Y.; Shirakura, K.; Okada, Y.; Hirayama, R.; Iwashita, Y.; Nishino, I.; Ago, Y.; Takeda, H.; Kuniyasu, H.; et al. Safety and efficacy of an anti-claudin-5 monoclonal antibody to increase blood-brain barrier permeability for drug delivery to the brain in a non-human primate. J. Control. Release 2021, 336, 105–111. [Google Scholar] [CrossRef]
  167. Nicolas, J.M.; Chanteux, H.; Nicolaï, J.; Brouta, F.; Viot, D.; Rosseels, M.L.; Gillent, E.; Bonnaillie, P.; Mathy, F.X.; Long, J.; et al. Role of P-glycoprotein in the brain disposition of seletalisib: Evaluation of the potential for drug-drug interactions. Eur. J. Pharm. Sci. 2020, 142, 105122. [Google Scholar] [CrossRef]
  168. Ye, D.; Dawson, K.A.; Lynch, I. A TEM protocol for quality assurance of in vitro cellular barrier models and its application to the assessment of nanoparticle transport mechanisms across barriers. Analyst 2015, 140, 83–97. [Google Scholar] [CrossRef]
  169. Srinivasan, B.; Kolli, A.R.; Esch, M.B.; Abaci, H.E.; Shuler, M.L.; Hickman, J.J. TEER measurement techniques for in vitro barrier model systems. J. Lab. Autom. 2015, 20, 107–126. [Google Scholar] [CrossRef]
  170. Vigh, J.P.; Kincses, A.; Ozgür, B.; Walter, F.R.; Santa-Maria, A.R.; Valkai, S.; Vastag, M.; Neuhaus, W.; Brodin, B.; Dér, A.; et al. Transendothelial electrical resistance measurement across the blood-brain barrier: A critical review of methods. Micromachines 2021, 12, 685. [Google Scholar] [CrossRef]
  171. Nag, S. Blood-brain barrier permeability using tracers and immunohistochemistry. Methods Mol. Med. 2003, 89, 133–144. [Google Scholar] [CrossRef] [PubMed]
  172. Saunders, N.R.; Dziegielewska, K.M.; Møllgård, K.; Habgood, M.D. Markers for blood-brain barrier integrity: How appropriate is Evans blue in the twenty-first century and what are the alternatives? Front. Neurosci. 2015, 9, 385. [Google Scholar] [CrossRef] [PubMed]
  173. Wu, M.C.; Hsu, J.L.; Lai, T.W. Evans blue dye as an indicator of albumin permeability across a brain endothelial cell monolayer in vitro. Neuroreport 2021, 32, 957–964. [Google Scholar] [CrossRef] [PubMed]
  174. Sharma, B.; Luhach, K.; Kulkarni, G.T. In vitro and in vivo models of BBB to evaluate brain targeting drug delivery. In Brain Targeted Drug Delivery System; Academic Press: Cambridge, MA, USA, 2019; pp. 53–101. [Google Scholar] [CrossRef]
  175. Fossan, G.; Cavanagh, M.E.; Evans, C.A.; Malinowska, D.H.; Møllgård, K.; Reynolds, M.L.; Saunders, N.R. CSF-brain permeability in the immature sheep fetus: A CSF-brain barrier. Dev. Brain Res. 1985, 350, 113–124. [Google Scholar] [CrossRef]
  176. Liddelow, S.A.; Dziegielewska, K.M.; Ek, C.J.; Johansson, P.A.; Potter, A.M.; Saunders, N.R. Cellular transfer of macromolecules across the developing choroid plexus of Monodelphis domestica. Eur. J. Neurosci. 2009, 29, 253–266. [Google Scholar] [CrossRef]
  177. Ek, C.J.; Habgood, M.D.; Dziegielewska, K.M.; Saunders, N.R. Structural characteristics and barrier properties of the choroid plexuses in developing brain of the opossum (Monodelphis domestica). J. Comp. Neurol. 2003, 460, 451–464. [Google Scholar] [CrossRef]
  178. Neal, E.H.; Shi, Y.; Lippmann, E.S. In vitro blood-brain barrier functional assays in a human iPSC-based model. In Cell Culture Techniques; Aschner, M., Costa, L., Eds.; Springer: New York, NY, USA, 2019; pp. 1–15. [Google Scholar]
  179. Nitz, T.; Eisenblätter, T.; Haselbach, M.; Galla, H.-J. Recent advances in the development of cell culture models for the blood-brain- and blood-CSF-barrier. In Blood—Brain Barrier: Drug Delivery and Brain Pathology; Kobiler, D., Lustig, S., Shapira, S., Eds.; Springer: Boston, MA, USA, 2001; pp. 45–62. [Google Scholar]
  180. Wegener, J.; Hakvoort, A.; Galla, H.J. Barrier function of porcine choroid plexus epithelial cells is modulated by cAMP-dependent pathways in vitro. Brain Res. 2000, 853, 115–124. [Google Scholar] [CrossRef]
  181. Duffey, M.E.; Hainau, B.; Ho, S.; Bentzel, C.J. Regulation of epithelial tight junction permeability by cyclic AMP. Nature 1981, 294, 451–453. [Google Scholar] [CrossRef]
  182. Tenenbaum, T.; Matalon, D.; Adam, R.; Seibt, A.; Wewer, C.; Schwerk, C.; Galla, H.J.; Schroten, H. Dexamethasone prevents alteration of tight junction-associated proteins and barrier function in porcine choroid plexus epithelial cells after infection with Streptococcus suis in vitro. Brain Res. 2008, 1229, 1–17. [Google Scholar] [CrossRef]
  183. Stone, N.L.; England, T.J.; O’Sullivan, S.E. A novel Transwell blood brain barrier model using primary human cells. Front. Cell. Neurosci. 2019, 13, 230. [Google Scholar] [CrossRef]
  184. Shi, L.Z.; Zheng, W. Establishment of an in vitro brain barrier epithelial transport system for pharmacological and toxicological study. Brain Res. 2005, 1057, 37–48. [Google Scholar] [CrossRef] [PubMed]
  185. Förster, C.; Burek, M.; Romero, I.A.; Weksler, B.; Couraud, P.O.; Drenckhahn, D. Differential effects of hydrocortisone and TNFalpha on tight junction proteins in an in vitro model of the human blood-brain barrier. J. Physiol. 2008, 586, 1937–1949. [Google Scholar] [CrossRef] [PubMed]
  186. Neumaier, F.; Zlatopolskiy, B.D.; Neumaier, B. Drug penetration into the central nervous system: Pharmacokinetic concepts and in vitro model systems. Pharmaceutics 2021, 13, 1542. [Google Scholar] [CrossRef] [PubMed]
  187. Van der Helm, M.W.; van der Meer, A.D.; Eijkel, J.C.T.; van den Berg, A.; Segerink, L.I. Microfluidic organ-on-chip technology for blood-brain barrier research. Tissue Barriers 2016, 4, e1142493. [Google Scholar] [CrossRef]
  188. Strazielle, N.; Belin, M.F.; Ghersi-Egea, J.F. Choroid plexus controls brain availability of anti-HIV nucleoside analogs via pharmacologically inhibitable organic anion transporters. Aids 2003, 17, 1473–1485. [Google Scholar] [CrossRef]
  189. Shen, H.; Keep, R.F.; Hu, Y.; Smith, D.E. PEPT2 (Slc15a2)-mediated unidirectional transport of cefadroxil from cerebrospinal fluid into choroid plexus. J. Pharmacol. Exp. Ther. 2005, 315, 1101–1108. [Google Scholar] [CrossRef]
  190. Hu, H.H.; Bian, Y.C.; Liu, Y.; Sheng, R.; Jiang, H.D.; Yu, L.S.; Hu, Y.Z.; Zeng, S. Evaluation of blood-brain barrier and blood-cerebrospinal fluid barrier permeability of 2-phenoxy-indan-1-one derivatives using in vitro cell models. Int. J. Pharm. 2014, 460, 101–107. [Google Scholar] [CrossRef]
  191. Crossgrove, J.S.; Li, G.J.; Zheng, W. The choroid plexus removes beta-amyloid from brain cerebrospinal fluid. Exp. Biol. Med. 2005, 230, 771–776. [Google Scholar] [CrossRef]
  192. Wang, X.; Li, G.J.; Zheng, W. Efflux of iron from the cerebrospinal fluid to the blood at the blood-CSF barrier: Effect of manganese exposure. Exp. Biol. Med. 2008, 233, 1561–1571. [Google Scholar] [CrossRef]
  193. Siegal, T. Strategies for increasing drug delivery to the brain. In Blood—Brain Barrier: Drug Delivery and Brain Pathology; Kobiler, D., Lustig, S., Shapira, S., Eds.; Springer: Boston, MA, USA, 2001; pp. 251–271. [Google Scholar]
  194. Southwell, B.R.; Duan, W.; Alcorn, D.; Brack, C.; Richardson, S.J.; Köhrle, J.; Schreiber, G. Thyroxine transport to the brain: Role of protein synthesis by the choroid plexus. Endocrinology 1993, 133, 2116–2126. [Google Scholar] [CrossRef]
  195. Peiser, C.; McGregor, G.P.; Lang, R.E. Binding and internalization of leptin by porcine choroid plexus cells in culture. Neurosci. Lett. 2000, 283, 209–212. [Google Scholar] [CrossRef]
  196. Tachikawa, M.; Kasai, Y.; Takahashi, M.; Fujinawa, J.; Kitaichi, K.; Terasaki, T.; Hosoya, K. The blood-cerebrospinal fluid barrier is a major pathway of cerebral creatinine clearance: Involvement of transporter-mediated process. J. Neurochem. 2008, 107, 432–442. [Google Scholar] [CrossRef] [PubMed]
  197. Duarte, A.C.; Rosado, T.; Costa, A.R.; Santos, J.; Gallardo, E.; Quintela, T.; Ishikawa, H.; Schwerk, C.; Schroten, H.; Gonçalves, I.; et al. The bitter taste receptor TAS2R14 regulates resveratrol transport across the human blood-cerebrospinal fluid barrier. Biochem. Pharmacol. 2020, 177, 113953. [Google Scholar] [CrossRef] [PubMed]
  198. Baird, A.; Eliceiri, B.P.; Gonzalez, A.M.; Johanson, C.E.; Leadbeater, W.; Stopa, E.G. Targeting the choroid plexus-CSF-brain nexus using peptides identified by phage display. Methods Mol. Biol. 2011, 686, 483–498. [Google Scholar] [CrossRef]
  199. Vercauteren, D.; Vandenbroucke, R.E.; Jones, A.T.; Rejman, J.; Demeester, J.; De Smedt, S.C.; Sanders, N.N.; Braeckmans, K. The use of inhibitors to study endocytic pathways of gene carriers: Optimization and pitfalls. Mol. Ther. 2010, 18, 561–569. [Google Scholar] [CrossRef]
  200. Strazielle, N.; Ghersi-Egea, J.F. Potential pathways for CNS drug delivery across the blood-cerebrospinal fluid barrier. Curr. Pharm. Des. 2016, 22, 5463–5476. [Google Scholar] [CrossRef]
  201. Abdul Razzak, R.; Florence, G.J.; Gunn-Moore, F.J. Approaches to CNS drug delivery with a focus on transporter-mediated transcytosis. Int. J. Mol. Sci. 2019, 20, 3108. [Google Scholar] [CrossRef]
  202. Bryniarski, M.A.; Ren, T.; Rizvi, A.R.; Snyder, A.M.; Morris, M.E. Targeting the choroid plexuses for protein drug delivery. Pharmaceutics 2020, 12, 963. [Google Scholar] [CrossRef]
  203. Schulingkamp, R.J.; Pagano, T.C.; Hung, D.; Raffa, R.B. Insulin receptors and insulin action in the brain: Review and clinical implications. Neurosci. Biobehav. Rev. 2000, 24, 855–872. [Google Scholar] [CrossRef]
  204. Johnsen, K.B.; Burkhart, A.; Thomsen, L.B.; Andresen, T.L.; Moos, T. Targeting the transferrin receptor for brain drug delivery. Prog. Neurobiol. 2019, 181, 101665. [Google Scholar] [CrossRef]
  205. Bien-Ly, N.; Yu, Y.J.; Bumbaca, D.; Elstrott, J.; Boswell, C.A.; Zhang, Y.; Luk, W.; Lu, Y.; Dennis, M.S.; Weimer, R.M.; et al. Transferrin receptor (TfR) trafficking determines brain uptake of TfR antibody affinity variants. J. Exp. Med. 2014, 211, 233–244. [Google Scholar] [CrossRef] [PubMed]
  206. Molino, Y.; David, M.; Varini, K.; Jabès, F.; Gaudin, N.; Fortoul, A.; Bakloul, K.; Masse, M.; Bernard, A.; Drobecq, L.; et al. Use of LDL receptor-targeting peptide vectors for in vitro and in vivo cargo transport across the blood-brain barrier. FASEB J. 2017, 31, 1807–1827. [Google Scholar] [CrossRef] [PubMed]
  207. Jones, A.R.; Shusta, E.V. Blood-brain barrier transport of therapeutics via receptor-mediation. Pharm. Res. 2007, 24, 1759–1771. [Google Scholar] [CrossRef] [PubMed]
  208. Couch, J.A.; Yu, Y.J.; Zhang, Y.; Tarrant, J.M.; Fuji, R.N.; Meilandt, W.J.; Solanoy, H.; Tong, R.K.; Hoyte, K.; Luk, W.; et al. Addressing safety liabilities of TfR bispecific antibodies that cross the blood-brain barrier. Sci. Transl. Med. 2013, 5, 183ra157. [Google Scholar] [CrossRef] [PubMed]
  209. Farrington, G.K.; Caram-Salas, N.; Haqqani, A.S.; Brunette, E.; Eldredge, J.; Pepinsky, B.; Antognetti, G.; Baumann, E.; Ding, W.; Garber, E.; et al. A novel platform for engineering blood-brain barrier-crossing bispecific biologics. FASEB J. 2014, 28, 4764–4778. [Google Scholar] [CrossRef]
  210. Broadwell, R.D.; Baker-Cairns, B.J.; Friden, P.M.; Oliver, C.; Villegas, J.C. Transcytosis of protein through the mammalian cerebral epithelium and endothelium: III. Receptor-mediated transcytosis through the blood–brain barrier of blood-borne transferrin and antibody against the transferrin receptor. Exp. Neurol. 1996, 142, 47–65. [Google Scholar] [CrossRef]
  211. Thom, G.; Burrell, M.; Haqqani, A.S.; Yogi, A.; Lessard, E.; Brunette, E.; Delaney, C.; Baumann, E.; Callaghan, D.; Rodrigo, N.; et al. Enhanced delivery of galanin conjugates to the brain through bioengineering of the anti-transferrin receptor antibody OX26. Mol. Pharm. 2018, 15, 1420–1431. [Google Scholar] [CrossRef]
  212. Boado, R.J.; Hui, E.K.; Lu, J.Z.; Pardridge, W.M. Drug targeting of erythropoietin across the primate blood-brain barrier with an IgG molecular Trojan horse. J. Pharmacol. Exp. Ther. 2010, 333, 961–969. [Google Scholar] [CrossRef]
  213. Boado, R.J.; Zhang, Y.; Zhang, Y.; Pardridge, W.M. Humanization of anti-human insulin receptor antibody for drug targeting across the human blood-brain barrier. Biotechnol. Bioeng. 2007, 96, 381–391. [Google Scholar] [CrossRef]
  214. Giugliani, R.; Giugliani, L.; de Oliveira Poswar, F.; Donis, K.C.; Corte, A.D.; Schmidt, M.; Boado, R.J.; Nestrasil, I.; Nguyen, C.; Chen, S.; et al. Neurocognitive and somatic stabilization in pediatric patients with severe Mucopolysaccharidosis Type I after 52 weeks of intravenous brain-penetrating insulin receptor antibody-iduronidase fusion protein (valanafusp alpha): An open label phase 1-2 trial. Orphanet J. Rare Dis. 2018, 13, 110. [Google Scholar] [CrossRef]
  215. Shu, C.; Shen, H.; Teuscher, N.S.; Lorenzi, P.J.; Keep, R.F.; Smith, D.E. Role of PEPT2 in peptide/mimetic trafficking at the blood-cerebrospinal fluid barrier: Studies in rat choroid plexus epithelial cells in primary culture. J. Pharmacol. Exp. Ther. 2002, 301, 820–829. [Google Scholar] [CrossRef] [PubMed]
  216. Joseph, A.; Nance, E. Nanotherapeutics and the brain. Annu. Rev. Chem. Biomol. Eng. 2022, 13, 325–346. [Google Scholar] [CrossRef] [PubMed]
  217. Sharma, M.; Dube, T.; Chibh, S.; Kour, A.; Mishra, J.; Panda, J.J. Nanotheranostics, a future remedy of neurological disorders. Expert Opin. Drug Deliv. 2019, 16, 113–128. [Google Scholar] [CrossRef] [PubMed]
  218. Dong, X. Current strategies for brain drug delivery. Theranostics 2018, 8, 1481–1493. [Google Scholar] [CrossRef]
  219. Kim, J.; Ahn, S.I.; Kim, Y. Nanotherapeutics engineered to cross the blood-brain barrier for advanced drug delivery to the central nervous system. J. Ind. Eng. Chem. 2019, 73, 8–18. [Google Scholar] [CrossRef]
  220. Tang, W.; Fan, W.; Lau, J.; Deng, L.; Shen, Z.; Chen, X. Emerging blood-brain-barrier-crossing nanotechnology for brain cancer theranostics. Chem. Soc. Rev. 2019, 48, 2967–3014. [Google Scholar] [CrossRef]
  221. Tang, L.; Feng, Y.; Gao, S.; Mu, Q.; Liu, C. Nanotherapeutics overcoming the blood-brain barrier for glioblastoma treatment. Front. Pharmacol. 2021, 12, 786700. [Google Scholar] [CrossRef]
  222. Alotaibi, B.S.; Buabeid, M.; Ibrahim, N.A.; Kharaba, Z.J.; Ijaz, M.; Noreen, S.; Murtaza, G. Potential of nanocarrier-based drug delivery systems for brain targeting: A current review of literature. Int. J. Nanomed. 2021, 16, 7517–7533. [Google Scholar] [CrossRef]
  223. Nehra, M.; Uthappa, U.T.; Kumar, V.; Kumar, R.; Dixit, C.; Dilbaghi, N.; Mishra, Y.K.; Kumar, S.; Kaushik, A. Nanobiotechnology-assisted therapies to manage brain cancer in personalized manner. J. Control. Release 2021, 338, 224–243. [Google Scholar] [CrossRef]
  224. Furtado, D.; Björnmalm, M.; Ayton, S.; Bush, A.I.; Kempe, K.; Caruso, F. Overcoming the blood-brain barrier: The role of nanomaterials in treating neurological diseases. Adv. Mater. 2018, 30, e1801362. [Google Scholar] [CrossRef]
  225. Lynch, M.J.; Gobbo, O.L. Advances in non-animal testing approaches towards accelerated clinical translation of novel nanotheranostic therapeutics for central nervous system disorders. Nanomaterials 2021, 11, 2632. [Google Scholar] [CrossRef] [PubMed]
  226. Kumar, A.; Chaudhary, R.K.; Singh, R.; Singh, S.P.; Wang, S.Y.; Hoe, Z.Y.; Pan, C.T.; Shiue, Y.L.; Wei, D.Q.; Kaushik, A.C.; et al. nanotheranostic applications for detection and targeting neurodegenerative diseases. Front. Neurosci. 2020, 14, 305. [Google Scholar] [CrossRef] [PubMed]
  227. Kaur, M.; Badhan, R.K. Phytoestrogens modulate breast cancer resistance protein expression and function at the blood-cerebrospinal fluid barrier. J. Pharm. Pharm. Sci. 2015, 18, 132–154. [Google Scholar] [CrossRef]
  228. Sanders-Bush, E.; Breeding, M. Choroid plexus epithelial cells in primary culture: A model of 5HT1C receptor activation by hallucinogenic drugs. Psychopharmacology 1991, 105, 340–346. [Google Scholar] [CrossRef] [PubMed]
  229. Barker, E.L.; Sanders-Bush, E. 5-hydroxytryptamine1C receptor density and mRNA levels in choroid plexus epithelial cells after treatment with mianserin and (-)-1-(4-bromo-2,5-dimethoxyphenyl)-2-aminopropane. Mol. Pharmacol. 1993, 44, 725–730. [Google Scholar] [PubMed]
  230. O’Hara, B.A.; Gee, G.V.; Haley, S.A.; Morris-Love, J.; Nyblade, C.; Nieves, C.; Hanson, B.A.; Dang, X.; Turner, T.J.; Chavin, J.M.; et al. Teriflunomide inhibits JCPyV infection and spread in glial cells and choroid plexus epithelial cells. Int. J. Mol. Sci. 2021, 22, 9809. [Google Scholar] [CrossRef]
  231. Song, H.; Zheng, G.; Liu, Y.; Shen, X.F.; Zhao, Z.H.; Aschner, M.; Luo, W.J.; Chen, J.Y. Cellular uptake of lead in the blood-cerebrospinal fluid barrier: Novel roles of Connexin 43 hemichannel and its down-regulations via Erk phosphorylation. Toxicol. Appl. Pharmacol. 2016, 297, 1–11. [Google Scholar] [CrossRef]
  232. Zheng, G.; Zhang, J.; Xu, Y.; Shen, X.; Song, H.; Jing, J.; Luo, W.; Zheng, W.; Chen, J. Involvement of CTR1 and ATP7A in lead (Pb)-induced copper (Cu) accumulation in choroidal epithelial cells. Toxicol. Lett. 2014, 225, 110–118. [Google Scholar] [CrossRef]
  233. Zhao, Q.; Slavkovich, V.; Zheng, W. Lead exposure promotes translocation of protein kinase C activities in rat choroid plexus in vitro, but not in vivo. Toxicol. Appl. Pharmacol. 1998, 149, 99–106. [Google Scholar] [CrossRef]
  234. Shi, L.Z.; Zheng, W. Early lead exposure increases the leakage of the blood-cerebrospinal fluid barrier, in vitro. Hum. Exp. Toxicol. 2007, 26, 159–167. [Google Scholar] [CrossRef]
  235. Lohren, H.; Bornhorst, J.; Galla, H.J.; Schwerdtle, T. The blood-cerebrospinal fluid barrier—First evidence for an active transport of organic mercury compounds out of the brain. Metallomics 2015, 7, 1420–1430. [Google Scholar] [CrossRef]
  236. Zheng, W. Toxicology of choroid plexus: Special reference to metal-induced neurotoxicities. Microsc. Res. Tech. 2001, 52, 89–103. [Google Scholar] [CrossRef]
  237. Bornhorst, J.; Wehe, C.A.; Hüwel, S.; Karst, U.; Galla, H.J.; Schwerdtle, T. Impact of manganese on and transfer across blood-brain and blood-cerebrospinal fluid barrier in vitro. J. Biol. Chem. 2012, 287, 17140–17151. [Google Scholar] [CrossRef] [PubMed]
  238. Schmitt, C.; Strazielle, N.; Richaud, P.; Bouron, A.; Ghersi-Egea, J.F. Active transport at the blood-CSF barrier contributes to manganese influx into the brain. J. Neurochem. 2011, 117, 747–756. [Google Scholar] [CrossRef] [PubMed]
  239. Morgan, S.E.; Schroten, H.; Ishikawa, H.; Zhao, N. Localization of ZIP14 and ZIP8 in HIBCPP Cells. Brain Sci. 2020, 10, 534. [Google Scholar] [CrossRef]
  240. Monnot, A.D.; Zheng, G.; Zheng, W. Mechanism of copper transport at the blood-cerebrospinal fluid barrier: Influence of iron deficiency in an in vitro model. Exp. Biol. Med. 2012, 237, 327–333. [Google Scholar] [CrossRef]
  241. Zheng, W.; Aschner, M.; Ghersi-Egea, J.F. Brain barrier systems: A new frontier in metal neurotoxicological research. Toxicol. Appl. Pharmacol. 2003, 192, 1–11. [Google Scholar] [CrossRef]
  242. Young, R.K.; Villalobos, A.R. Stress-induced stimulation of choline transport in cultured choroid plexus epithelium exposed to low concentrations of cadmium. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2014, 306, R291–R303. [Google Scholar] [CrossRef]
  243. Johanson, C.E.; Murphy, V.A. Acetazolamide and insulin alter choroid plexus epithelial cell [Na+], pH, and volume. Am. J. Physiol. 1990, 258, F1538–F1546. [Google Scholar] [CrossRef]
  244. Uldall, M.; Botfield, H.; Jansen-Olesen, I.; Sinclair, A.; Jensen, R. Acetazolamide lowers intracranial pressure and modulates the cerebrospinal fluid secretion pathway in healthy rats. Neurosci. Lett. 2017, 645, 33–39. [Google Scholar] [CrossRef]
  245. Collins, P.; Morriss, G.M. Changes in the surface features of choroid plexus of the rat following the administration of acetazolamide and other drugs which affect CSF secretion. J. Anat. 1975, 120, 571–579. [Google Scholar] [PubMed]
  246. Supuran, C.T. Acetazolamide for the treatment of idiopathic intracranial hypertension. Expert Rev. Neurother. 2015, 15, 851–856. [Google Scholar] [CrossRef] [PubMed]
  247. Ameli, P.A.; Madan, M.; Chigurupati, S.; Yu, A.; Chan, S.L.; Pattisapu, J.V. Effect of acetazolamide on aquaporin-1 and fluid flow in cultured choroid plexus. Acta Neurochir. Suppl. 2012, 113, 59–64. [Google Scholar] [CrossRef]
  248. Swetloff, A.; Greenwood, S.; Wade, A.M.; Ferretti, P. Growth of choroid plexus epithelium vesicles in vitro depends on secretory activity. J. Cell. Physiol. 2006, 208, 549–555. [Google Scholar] [CrossRef]
  249. Murphy, V.A.; Johanson, C.E. Alteration of sodium transport by the choroid plexus with amiloride. Biochim. Biophys. Acta 1989, 979, 187–192. [Google Scholar] [CrossRef]
  250. Johanson, C.E.; Palm, D.E.; Dyas, M.L.; Knuckey, N.W. Microdialysis analysis of effects of loop diuretics and acetazolamide on chloride transport from blood to CSF. Brain Res. 1994, 641, 121–126. [Google Scholar] [CrossRef]
  251. Löscher, W.; Kaila, K. CNS pharmacology of NKCC1 inhibitors. Neuropharmacology 2022, 205, 108910. [Google Scholar] [CrossRef]
  252. Gregoriades, J.M.C.; Madaris, A.; Alvarez, F.J.; Alvarez-Leefmans, F.J. Genetic and pharmacological inactivation of apical Na+-K+-2Cl cotransporter 1 in choroid plexus epithelial cells reveals the physiological function of the cotransporter. Am. J. Physiol. Cell Physiol. 2019, 316, C525–C544. [Google Scholar] [CrossRef]
  253. Culliford, S.; Ellory, C.; Lang, H.J.; Englert, H.; Staines, H.; Wilkins, R. Specificity of classical and putative Cl transport inhibitors on membrane transport pathways in human erythrocytes. Cell. Physiol. Biochem. 2003, 13, 181–188. [Google Scholar] [CrossRef]
  254. Hughes, A.L.; Pakhomova, A.; Brown, P.D. Regulatory volume increase in epithelial cells isolated from the mouse fourth ventricle choroid plexus involves Na+-H+ exchange but not Na+-K+-2Cl cotransport. Brain Res. 2010, 1323, 1–10. [Google Scholar] [CrossRef]
  255. Bouzinova, E.V.; Praetorius, J.; Virkki, L.V.; Nielsen, S.; Boron, W.F.; Aalkjaer, C. Na+-dependent HCO3 uptake into the rat choroid plexus epithelium is partially DIDS sensitive. Am. J. Physiol. Cell Physiol. 2005, 289, C1448–C1456. [Google Scholar] [CrossRef] [PubMed]
  256. Damkier, H.; Praetorius, J. Structure of the mammalian choroid plexus. In Role of the Choroid Plexus in Health and Disease; Praetorius, J., Blazer-Yost, B., Damkier, H., Eds.; Springer: New York, NY, USA, 2020; pp. 1–33. [Google Scholar]
  257. Hung, B.C.; Loo, D.D.; Wright, E.M. Regulation of mouse choroid plexus apical Cl and K+ channels by serotonin. Brain Res. 1993, 617, 285–295. [Google Scholar] [CrossRef]
  258. Garner, C.; Feniuk, W.; Brown, P.D. Serotonin activates Cl channels in the apical membrane of rat choroid plexus epithelial cells. Eur. J. Pharmacol. 1993, 239, 31–37. [Google Scholar] [CrossRef]
  259. Narita, K.; Sasamoto, S.; Koizumi, S.; Okazaki, S.; Nakamura, H.; Inoue, T.; Takeda, S. TRPV4 regulates the integrity of the blood-cerebrospinal fluid barrier and modulates transepithelial protein transport. FASEB J. 2015, 29, 2247–2259. [Google Scholar] [CrossRef] [PubMed]
  260. Preston, D.; Simpson, S.; Halm, D.; Hochstetler, A.; Schwerk, C.; Schroten, H.; Blazer-Yost, B.L. Activation of TRPV4 stimulates transepithelial ion flux in a porcine choroid plexus cell line. Am. J. Physiol. Cell Physiol. 2018, 315, C357–C366. [Google Scholar] [CrossRef] [PubMed]
  261. Dragunow, M. Meningeal and choroid plexus cells—Novel drug targets for CNS disorders. Brain Res. 2013, 1501, 32–55. [Google Scholar] [CrossRef]
  262. Johanson, C.; Johanson, N. Merging transport data for choroid plexus with blood-brain barrier to model CNS homeostasis and disease more effectively. CNS Neurol. Disord. Drug Targets 2016, 15, 1151–1180. [Google Scholar] [CrossRef]
  263. Marques, F.; Sousa, J.C.; Brito, M.A.; Pahnke, J.; Santos, C.; Correia-Neves, M.; Palha, J.A. The choroid plexus in health and in disease: Dialogues into and out of the brain. Neurobiol. Dis. 2017, 107, 32–40. [Google Scholar] [CrossRef]
  264. Safaee, M.; Oh, M.C.; Bloch, O.; Sun, M.Z.; Kaur, G.; Auguste, K.I.; Tihan, T.; Parsa, A.T. Choroid plexus papillomas: Advances in molecular biology and understanding of tumorigenesis. Neuro Oncol. 2013, 15, 255–267. [Google Scholar] [CrossRef]
  265. Keep, R.F.; Jones, H.C.; Drewes, L.R. Brain barriers and brain fluids research in 2020 and the fluids and barriers of the CNS thematic series on advances in in vitro modeling of the blood-brain barrier and neurovascular unit. Fluids Barriers CNS 2021, 18, 24. [Google Scholar] [CrossRef]
  266. Rodríguez-Lorenzo, S.; Konings, J.; van der Pol, S.; Kamermans, A.; Amor, S.; van Horssen, J.; Witte, M.E.; Kooij, G.; de Vries, H.E. Inflammation of the choroid plexus in progressive multiple sclerosis: Accumulation of granulocytes and T cells. Acta Neuropathol. Commun. 2020, 8, 9. [Google Scholar] [CrossRef] [PubMed]
  267. Solár, P.; Zamani, A.; Kubíčková, L.; Dubový, P.; Joukal, M. Choroid plexus and the blood-cerebrospinal fluid barrier in disease. Fluids Barriers CNS 2020, 17, 35. [Google Scholar] [CrossRef] [PubMed]
  268. Jansson, D.; Dieriks, V.B.; Rustenhoven, J.; Smyth, L.C.D.; Scotter, E.; Aalderink, M.; Feng, S.; Johnson, R.; Schweder, P.; Mee, E.; et al. Cardiac glycosides target barrier inflammation of the vasculature, meninges and choroid plexus. Commun. Biol. 2021, 4, 260. [Google Scholar] [CrossRef] [PubMed]
  269. Krzyzankova, M.; Mertsch, S.; Koos, B.; Jeibmann, A.; Kruse, A.; Kordes, U.; Frühwald, M.C.; Wolff, J.E.; Paulus, W.; Hasselblatt, M. Loss of TP53 expression in immortalized choroid plexus epithelial cells results in increased resistance to anticancer agents. J. Neurooncol. 2012, 109, 449–455. [Google Scholar] [CrossRef]
Figure 1. Schematic representation of anatomical location, physiological properties, and pharmacologically related transport systems of the BCSFB. (A) The BCSFB structure is comprised of the choroid plexus polarized cuboidal epithelial cells surrounding the highly permeable fenestrated capillaries of stromal core and tight junctional strands uniting adjacent epithelial cells. The innermost capillary (containing red blood cells), with leaky inter-endothelial gap junctions, is alongside the underlying stroma/basement membrane extracellular matrix. The epithelial basolateral surface faces stroma/blood and is in contact with interstitial fluid (ISF). The brush border apical membrane containing microvilli faces the adjacent CSF. (B) The main transport-relevant features of the BCSFB in terms of influx and efflux transport systems responsible for supplying nutrients, hormones, and therapeutics to the brain/CSF or acting to eliminate metabolites, xenobiotics, and neurotoxic compounds, respectively, are depicted.
Figure 1. Schematic representation of anatomical location, physiological properties, and pharmacologically related transport systems of the BCSFB. (A) The BCSFB structure is comprised of the choroid plexus polarized cuboidal epithelial cells surrounding the highly permeable fenestrated capillaries of stromal core and tight junctional strands uniting adjacent epithelial cells. The innermost capillary (containing red blood cells), with leaky inter-endothelial gap junctions, is alongside the underlying stroma/basement membrane extracellular matrix. The epithelial basolateral surface faces stroma/blood and is in contact with interstitial fluid (ISF). The brush border apical membrane containing microvilli faces the adjacent CSF. (B) The main transport-relevant features of the BCSFB in terms of influx and efflux transport systems responsible for supplying nutrients, hormones, and therapeutics to the brain/CSF or acting to eliminate metabolites, xenobiotics, and neurotoxic compounds, respectively, are depicted.
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Figure 2. Various possible in vitro BCSFB model platforms are schematically depicted here. (A) Static bicameral devices or alternatively known as cell culture filter inserts monolayers. This compartmentalized model, as the mostly utilized configuration by a preponderance of studies, represents culture of choroid plexus epithelial cells either in standard or inverted format on a suitable permeable filter insert; (B) Co-culture and multi-culture filter inserts. Here, choroid plexus epithelial cells are grown on porous cell culture inserts alongside endothelial, mesenchymal (e.g., pericytes), and/or glial cells either cultivated into the bottom of a multi-well plate in which the insert is located (non-contact) or seeded on the opposite side of the inserts containing epithelial cells (leading to a so-called back-to-back contact co-culture); (C) Three-dimensional cultures and organoids; (D) Dynamic cultures or microfluidic devices. The upper and lower channels of the devices are separated to model the luminal and abluminal membrane interfaces based on cell culture direction and a tunable shear stress is induced by a continuous flow of culture medium.
Figure 2. Various possible in vitro BCSFB model platforms are schematically depicted here. (A) Static bicameral devices or alternatively known as cell culture filter inserts monolayers. This compartmentalized model, as the mostly utilized configuration by a preponderance of studies, represents culture of choroid plexus epithelial cells either in standard or inverted format on a suitable permeable filter insert; (B) Co-culture and multi-culture filter inserts. Here, choroid plexus epithelial cells are grown on porous cell culture inserts alongside endothelial, mesenchymal (e.g., pericytes), and/or glial cells either cultivated into the bottom of a multi-well plate in which the insert is located (non-contact) or seeded on the opposite side of the inserts containing epithelial cells (leading to a so-called back-to-back contact co-culture); (C) Three-dimensional cultures and organoids; (D) Dynamic cultures or microfluidic devices. The upper and lower channels of the devices are separated to model the luminal and abluminal membrane interfaces based on cell culture direction and a tunable shear stress is induced by a continuous flow of culture medium.
Pharmaceutics 14 01729 g002
Table 1. Transporters at the BCSFB.
Table 1. Transporters at the BCSFB.
FamilyTransporter FunctionMembers Present on Choroid Plexus Epithelial Cells (Also-Known-as)
SLC1High-affinity glutamate and neutral amino acidsSLC1A3, SLC1A4, SLC1A5 (ASCT2)
SLC2Facultative GLUT transportersSLC2A1 (GLUT1), SLC2A6, SLC2A10, SLC2A12
SLC4Bicarbonate transporters (anion exchanger)SLC4A1, SLC4A2 (AE2), SLC4A4, SLC4A5 (NBC4/NBCe2), SLC4A8, SLC4A10, SLC4A11
SLC5Sodium glucose cotransportersSLC5A1, SLC5A5, SLC5A6
SLC6Sodium- and chloride-dependent neurotransmitter transportersSLC6A4, SLC6A6, SLC6A8 (Crt), SLC6A9, SLC6A11, SLC6A13, SLC6A14, SLC6A15, SLC6A17, SLC6A20A, SLC6A20B
SLC7Cationic amino acid transporter/glycoprotein- associatedSLC7A1, SLC7A2, SLC7A5 (LAT1), SLC7A6 (LAT2), SLC7A7, SLC7A10
SLC8Na+/Ca2+ exchangersSLC8A1
SLC9Na+/H+ exchangers (antiporter)SLC9A1 (NHE1), SLC9A2, SLC9A6 (NHE6), SLC9A7, SLC9A8, SLC9A9
SLC10Sodium/bile acid co-transporter familySLC10A3
SLC11Proton coupled metal ion transportersSLC11A2
SLC12Electroneutral cation-coupled Cl cotransportersSLC12A2 (NKCC1), SLC12A4 (KCC1)
SLC13Human Na+-sulfate/carboxylate cotransportersSLC13A4, SLC13A5
SLC14Urea transportersSLC14A2
SLC15Proton oligopeptide co-transportersSLC15A2 (PEPT2)
SLC16Monocarboxylate/monocarboxylic acid transporter familySLC16A3, SLC16A4, SLC16A6, SLC16A8, SLC16A9, SLC16A10
SLC17Vesicular glutamate transportersSLC17A6
SLC20Type III Na+-phosphate cotransportersSLC20A1, SLC20A2
SLC21/SLCOOrganic anion transportersSLCO1A5 (OATP1A5), SLCO 1C1, SLCO 2A1 (Pgt), SLCO5A1
SLC22Organic cation/anion/zwitterion transportersSLC22A5 (OCTN2), SLC22A6 (OAT1), SLC22A8 (OAT3), SLC22A17, SLC22A18, SLC22A21, SLC22A23
SLC23Na+-dependent ascorbic acid transportersSLC23A2
SLC24Na+/(Ca2+/K+) exchangersSLC24A3, SLC24A4, SLC24A5
SLC25Mitochondrial carriersSLC25A1, SLC25A10, SLC25A12, SLC25A14, SLC25A16, SLC25A17, SLC25A18, SLC25A21, SLC25A22, SLC25A26, SLC25A27, SLC25A30, SLC25A32, SLC25A33, SLC25A35, SLC25A37, SLC25A38, SLC25A39, SLC25A45
SLC26Multifunctional anion exchangersSLC26A2, SLC26A7
SLC27Fatty acid transportersSLC27A1, SLC27A2, SLC27A3
SLC28Na+-coupled nucleoside transportersSLC28A3
SLC29Facilitative nucleoside transportersSLC29A2, SLC29A4 (PMAT)
SLC30Zn2+ efflux transportersSLC30A3, SLC30A4, SLC30A5, SLC30A6, SLC30A9, SLC30A10
SLC31Cu2+ transportersSLC31A1, SLC31A2
SLC33Acetyl-CoA transportersSLC33A1
SLC35Nucleoside-sugar transportersSLC35A1, SLC35A4, SLC35A5, SLC35D2, SLC35E2, SLC35E4, SLC35F1, SLC35F2, SLC35F3, SLC35F5
SLC37Sugar-phosphate/phosphate exchangersSLC37A1 (G3PP), SLC37A2
SLC38Amino acid transporterSLC38A1, SLC38A3, SLC38A4, SLC38A5, SLC38A11
SLC39Metal ion transportersSLC39A4, SLC39A8, SLC39A10, SLC39A11, SLC39A12, SLC39A14
SLC40Basolateral Fe2+ transportersSLC40A1
SLC41MgtE-like magnesium transportersSLC41A1, SLC41A2
SLC43Na+-independent, system-L-like amino acid transportersSLC43A1, SLC43A2
SLC44Choline-like transportersSLC44A3
SLC45Putative sugar transportersSLC45A4
SLC46Folate transportersSLC46A1, SLC46A3
SLC48Heme transportersSLC48A1
SLC50Sugar efflux transportersSLC50A1
Table 2. Summary of in vitro BCSFB models.
Table 2. Summary of in vitro BCSFB models.
Model and Its DescriptionApplicationsAdvantagesDisadvantagesThroughput
2D static bicameral devices (cell culture inserts) or compartmentalized monocultures.
  • Compartmentalized culturing.
  • Routinely used for permeability studies.
  • Evaluation of (neuro)toxicological profile of investigational compounds.
  • Lead compound identification/optimization phase studies.
  • Structure–activity relationships (SAR) analyses.
  • Being user-friendly, easy to set up, and low labor intensity.
  • Minimal cost.
  • Used when the isolated study of compounds and epithelial cells interaction is the aim.
  • 2D structure.
  • Generally low TEER values.
  • Too simple to fully replicate key features of the BCSFB.
  • Lack of contact with other cells.
  • Fail to mimic CP microenvironment due to lack of shear stress and blood/CSF flow.
  • Real-time readouts are not easily possible.
Moderate (offers HTS capabilities).
Co-culture models.Study drug permeability.
  • Allows co-culture of endothelial cells and other related cells.
  • Takes into account the impact of other elements of the BCSFB.
  • Higher TEER and greater barrier stability.
  • Lack the fluid flow-induced shear stress.
  • Relatively time-consuming.
  • Higher cost.
Moderate.
3D and organoids.
  • Search for therapeutic targets.
  • Study interventional strategies to control drugs and substances entry.
  • Evaluation of (neuro)toxicological profile of investigational compounds.
  • Lead compound identification/optimization phase studies.
  • Structure–activity relationships (SAR) analyses.
  • Human origin cells/tissue can be used.
  • 3D culture model.
  • Reduced re-differentiation.
  • High barrier tightness.
  • Better maintain the tight junction organization compared to bicameral systems.
  • Possibility to be applied for personalized medicine developments by using cells/tissues from a specific donor group.
  • Allow studies on an organ-level in both healthy and diseased conditions.
  • Not applicable for high-throughput quantitative permeability measurements.
  • Lack the fluid flow-induced shear stress.
  • Complicated procedure with greater skill required.
  • High processing time.
  • Differential process relies upon random and continuous addition of differentiation factors.
Low to medium.
Dynamic models
(microfluidic or organ-on-a-chip platforms).
  • Study drug permeability.
  • Study drug’s Pharmacokinetic elements.
  • Possibility of integrating imaging systems and sensors with real-time readouts.
  • Human-derived cells or tissues can be used.
  • Contribution of fluid shear stress as an important factor is considered.
  • Difficult to set up and maintain.
  • High technical prerequisite needed.
Low to medium.
Table 3. Summary of available cells lines.
Table 3. Summary of available cells lines.
Cell TypesMain Advantages and DisadvantagesOriginCellsSpecies Source
Primary cell cultures
  • More representative of the in vivo state.
  • But usually expensive and labor-intensive to isolate.
  • Have limited yield.
  • Exhibit difficulty obtaining from human origin.
CerebralPig primary cells, PCPECPig
Mouse primary cellsMouse
Rat primary cellsRat
Bovine primary cellsCow
Ovine primary cellsSheep
Rabbit primary cellsRabbit
HCPEpiCHuman
Non-cerebralNo Reports
Immortalized and continuous cell lines
  • Affordable and easily accessible.
  • Commercially available.
  • Easy to culture.
  • But usually do not mimic the native CP.
  • Possibly altered genotype/phenotype after many passages.
CerebralZ310Rat
TR-CSFBRat
ECPC3Mouse
ECPC4Mouse
SV11Mouse
PCP-RPig
HIBCPPHuman
CPC-2Human
iHCPEnCHuman
Non-cerebralMDCKDog
MDCK-MDR1Dog
RRCKDog
Caco-2Human
LLC-PK1Pig
Table 4. Exogenous tracers categorized based on target solute/compound permeability.
Table 4. Exogenous tracers categorized based on target solute/compound permeability.
MarkerSize
Molecular Weight (Da)Approximate Hydrodynamic Radius (nm)
Markers of Protein/Macromolecules Permeability
DyesEvans blue960NR 1
Trypan blue961NR
Fluorescent tracersFITC-dextran 150 kDa150,0009.0 ± 0.6
FITC-dextran 70 kDa70,0006
FITC-dextran 40 kDa40,0004.5
FITC-albumin67,0005.4 ± 0.1
Horseradish peroxidase40,0005–6
Microperoxidase19003.0
Radiolabeled Compounds[125I]-albumin~69,0003.5
[14C]-dextran 70 kDa~70,0006
Markers of Solute and Ion Permeability
Ionic Lanthanum138.90.12
Sodium Fluorescein3760.45
Lucifer Yellow4570.42
Biotin ethylenediamine286NR
FITC-dextran 3kDa30001.4
Radiolabeled Compounds[14C]-α-Aminoisobutyric acid103NR
[14C]-Sucrose3420.46
[3H]-mannitol1820.36
[14C]-Methotrexate455NR
[14C]-Inulin50001.3
1 Not Reported.
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Dabbagh, F.; Schroten, H.; Schwerk, C. In Vitro Models of the Blood–Cerebrospinal Fluid Barrier and Their Applications in the Development and Research of (Neuro)Pharmaceuticals. Pharmaceutics 2022, 14, 1729. https://doi.org/10.3390/pharmaceutics14081729

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Dabbagh F, Schroten H, Schwerk C. In Vitro Models of the Blood–Cerebrospinal Fluid Barrier and Their Applications in the Development and Research of (Neuro)Pharmaceuticals. Pharmaceutics. 2022; 14(8):1729. https://doi.org/10.3390/pharmaceutics14081729

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Dabbagh, Fatemeh, Horst Schroten, and Christian Schwerk. 2022. "In Vitro Models of the Blood–Cerebrospinal Fluid Barrier and Their Applications in the Development and Research of (Neuro)Pharmaceuticals" Pharmaceutics 14, no. 8: 1729. https://doi.org/10.3390/pharmaceutics14081729

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