Abstract

Advances in human induced pluripotent stem cell (iPSC) approaches have greatly expanded the use of patient-derived cellular models, including cortical-like neurons and brain organoids, to study genetic epilepsies. New protocols to differentiate iPSCs into various neural cell types, and to generate brain region-specific organoids, have accelerated progress. In addition, the application of gene editing techniques adds rigor to these studies and offers the opportunity to model rare genetic epilepsies by enabling correction or insertion of mutations to generate isogenic controls or virtual patients, respectively. Studies using patient-derived cells have provided insight into the mechanisms underlying seizures and associated comorbidities for an increasing number of epilepsy gene variants. In this chapter, methods for generating iPSCs, the culture and gene editing of human pluripotent stem cells (hPSCs; both iPSCs and human embryonic stem cells), and differentiation strategies to derive cortical-like neurons are described. Next, physiological assays that can be applied to study hPSC-derived neuronal and network activity are discussed. Finally, the chapter reviews studies interrogating the functional effects of specific epilepsy gene variants using hPSC-derived 2D neural and 3D brain organoid cultures, as well as the use of iPSC-derived cardiac myocytes to study the most devastating epilepsy complication, sudden unexpected death in epilepsy (SUDEP).The chapter ends with a description of current challenges and future directions for hPSC modeling of genetic epilepsies.

Introduction

Recent years have seen a surge in the discovery of epilepsy related genes and, with it, a blooming of mechanistic studies. Because of the inaccessibility of human epileptic tissue, the lack of adequate control tissue and the fact that only late-stage specimens are acquired when it is available, model systems are necessary for the study of epilepsy mechanisms and the development of novel therapies. Moreover, as epilepsies typically are associated with comorbidities involving diverse and widespread brain networks, a useful strategy is to incorporate multifaceted approaches that encompass structural, functional, and developmental features to model epilepsies and their associated comorbidities.

The mainstay of genetic epilepsy models over the past five decades has been spontaneous or genetically engineered rodent models harboring genetic abnormalities similar to those identified in human genetic epilepsies. Such models have contributed a wealth of knowledge to the epilepsy field, specifically surrounding the function of neural networks in epilepsy and the underlying in vivo pathophysiological processes. Zebrafish represent a more recent addition to the armamentarium of genetic epilepsy models and are particularly useful for high-throughput drug screening (Baraban, 2021). However, several limitations arise in the use of animal models of epilepsy. Fish and even mice possess important genetic, structural, and neurodevelopmental differences from humans. And while some rodent models exhibit a seizure phenotype that closely resembles those seen in humans, most do not completely recapitulate the complex pathophysiology of human epilepsies. Moreover, animal models for many genetic epilepsies do not exhibit spontaneous seizures. Despite extensive research using animal models, understanding of the cellular and molecular mechanisms surrounding monogenic epilepsies remains elusive.

To try to overcome some of these obstacles and expand the tools available to model epilepsies, human pluripotent stem cells (hPSCs), a term that encompasses both human embryonic stem cells (hESCs) and induced pluripotent stem cells (iPSCs), have become increasingly used. In 1998, the Thomson laboratory discovered that they could derive hESCs from blastocysts and that these cells could then be used to generate many different cell types in vitro (Thomson et al., 1998). The breadth of stem cell research was further enhanced in 2006 when Yamanaka and colleagues (Takahashi and Yamanaka, 2006) demonstrated that mouse somatic cells could be reprogrammed to generate iPSCs. This discovery was quickly followed by the generation of human iPSCs in 2007 (Meissner et al., 2007; Takahashi et al., 2007), and the use of human iPSCs in translational research, drug discovery, and therapeutic development has since become increasingly widespread. Skin or blood cells taken from a patient can be reprogrammed to iPSCs and used to model a genetic disease by generating the affected cell type. Germline genetic variants found in a given patient are carried on in their somatic cell-derived iPSCs, and in any subsequent progeny that are generated from these iPSCs. Since samples of the affected tissue cannot easily be taken from patients suffering from central nervous system (CNS) diseases, brain cells generated from patient iPSCs provide a useful tool to study and model the development of brain diseases, particularly neurodevelopmental disorders. In this chapter, we review the current techniques and applications of hPSC models for studying genetic epilepsies using both 2D and 3D neural culture approaches. Additionally, we highlight potential limitations and avenues for future research using hPSC model systems.

Generating iPSCs

The ability to generate patient-specific iPSC lines provides the opportunity to model a wide range of genetic epilepsies in an in vitro system. The forced expression of specific transcription factors reverts somatic cells to a pluripotent epigenetic state and silences lineage-specific genes. This process, called “reprogramming,” is typically carried out by introducing four transcription factors, Sox2, Oct3/4, Klf4, and c-Myc, into somatic cells (Takahashi et al., 2007; Takahashi and Yamanaka, 2006). These four factors are commonly referred to as the “Yamanaka factors.” Typically, reprogramming involves the use of dermal fibroblasts obtained by a skin biopsy. More recently, peripheral blood mononuclear cells derived from patient blood samples are increasingly used as a less invasive source of somatic cells for reprogramming, and kidney epithelial cells extracted from urine are another option (Okita et al., 2013; Zhou et al., 2011). These advances have broadened the source of iPSCs and have contributed to the ease and accessibility of iPSC research.

Gene Editing of iPSCs

Genome editing approaches have substantially advanced the use of hPSCs for modeling of brain disorders, including genetic epilepsies. Because the CRISPR/Cas9 endonuclease system facilitates the ability to make precise changes in the genome (Cong et al., 2013; Ran et al., 2013), this method offers several key applications for using hPSCs to model genetic epilepsies. First, the inherent variability between iPSC lines from different subjects due to genomic background affects may be abrogated by using CRISPR-based homology directed repair to correct epilepsy gene variants in patient iPSC lines and thereby generate isogenic control lines. Second, in cases where patient material is not available to generate iPSC lines, epilepsy-specific gene variants may be created in a control line to generate paired “virtual patient” and isogenic control lines. Third, CRISPR-based homology-directed repair may be used to introduce exogenous genomic sequences into a specific locus to tag genes of interest with fluorescent reporters for cell culture-based studies of intact or mutant protein localization or function (Roberts et al., 2017). Finally, CRISPR-mediated gene knockout (via the generation of out-of-frame insertions or deletions by non-homologous end joining) in a control line may be used to study genetic epilepsies caused by autosomal recessive variants or haploinsufficiency. Indeed, methods to make this latter approach more efficient by combining somatic cell reprogramming and CRISPR gene knockout have been developed and applied to generate iPSC models of genetic epilepsies (Tidball et al., 2017).

Methods of 2D Neuronal Differentiation

For studying epilepsies, the derivation of disease-relevant neural cell types from hPSCs offers unique opportunities to explore disease mechanisms and identify novel therapies. Most relevant to epilepsy are excitatory and inhibitory cortical neurons. Neuronal differentiation protocols for these cell types broadly fall into two main categories: small molecule directed differentiation and transcription factor induction. Below, we discuss these two methods applied to both excitatory and inhibitory cortical-like neurons. Similar protocols that generate hPSC-derived astrocytes and microglia have also been developed (Canals et al., 2018; Douvaras et al., 2017; Krencik and Zhang, 2011) and are not described further in the text (but see Fig. 42–1).

Figure 42–1.

Overview of schema for generating 2D in vitro cellular models using human iPSCs. Patient or healthy control somatic cells, such as fibroblasts or peripheral blood mononuclear cells (PBMCs), are reprogrammed into iPSCs using a cocktail containing the (more...)

Small molecule differentiation protocols used to produce cortical-like neurons involve the inhibition of the bone morphogenetic protein (BMP) and transforming growth factor-beta (TGF-β) signaling pathways. This method is known as dual SMAD inhibition (Chambers et al., 2009; Shi et al., 2012), as SMAD proteins are the signal transducing effectors for TGF-β/BMP signaling. Common protocols consist of monolayer or embryoid body formation with or without the use of small molecules. While monolayer protocols are less labor-intensive, embryoid body protocols provide a purer population of cortical neurons that are more mature and have longer neurites, although they do not exhibit any difference in electrophysiological properties compared to monolayer cultures (Chandrasekaran et al., 2017; Muratore et al., 2014). The cortical-like neurons exhibit electrophysiological activity after about 4 to 6 weeks from the start of differentiation. Modulation of the Sonic Hedgehog (SHH) pathway after development of a primitive neuroepithelium can bias cultures toward the generation of excitatory (with SHH pathway inhibitors) or inhibitory (using SHH activators) neurons. To derive interneurons, exposure to SHH pathway activators yields a homogeneous population of progenitors that express the medial ganglionic eminence (a key source of cortical interneuron progenitors) marker NKX2.1 by about 6 weeks (Liu et al., 2013a; Maroof et al., 2013; Nicholas et al., 2013). However, populations that are achieved through small-molecule patterning protocols for either excitatory or inhibitory cortical-like neurons typically exhibit heterogeneous stages of differentiation throughout the culture, and differentiation efficiency can often vary between cell lines. Additionally, prolonged culture is required for cells to reach maturity.

The forced expression of transcription factors in hPSCs has recently been used to overcome some of these challenges. Transcription factor induction rapidly generates a more homogeneous population of either cortical-like excitatory or inhibitory neurons with high efficiency and reproducibility (Lam et al., 2017; Pak et al., 2015; Sun et al., 2016a; Yang et al., 2017; Zhang et al., 2013). However, this rapid induction bypasses normal signaling pathways that occur during neural development and therefore may not be suitable for investigating certain neurodevelopmental disorders (Schafer et al., 2019). Cortical-like excitatory neurons are often generated by the forced expression of the transcription factor Neurogenin-2 (NGN2) alone or combined with NGN1 using a doxycycline-inducible expression system (Lam et al., 2017; Pak et al., 2015). These induced neurons show transcriptional expression profiles closer to that of neurons in superficial cortical layers, and they display short-term plasticity, synaptic function and modulation, and the ability to functionally integrate into the host after grafting. This is in contrast to the deeper layer cortical neurons that are typically generated using small-molecule patterning unless prolonged (3-month) culture durations are used (Shi et al., 2012).

Functional GABAergic interneurons with a high degree of synaptic maturity are generated by the transient expression of transcription factors ASCL1 and DLX2 with or without LHX6 and specific microRNAs (Sun et al., 2016a; Yang et al., 2017). The cells express subtype-specific GABAergic interneuron markers and functionally integrate into host circuits after in vivo transplantation. Excitatory and inhibitory neurons can be co-cultured together and provide a useful platform for studying diseases that affect neural circuitry. An ongoing focus in the field is the challenge of generating parvalbumin (PV)-expressing fast-spiking interneurons, which are thought to be critically involved in the pathophysiology of many epilepsies and other neuropsychiatric disorders (Hu et al., 2017; Jiang et al., 2016). PV interneurons are difficult to achieve in culture due to the prolonged maturity that is required to generate these cells. More recently, brain organoid models (see below) have emerged as a powerful alternative to 2D modeling as they allow for a wider variety of cell types to be generated and cultured for extended periods of time.

Physiological Assays for hPSC-Derived Neurons

As epilepsy is a disease characterized by functional changes in neuronal and neural circuit activity, the ability to assess the physiological impact of ictogenic or epileptogenic changes in hPSC-derived cultures is critically important. A number of approaches spanning the gamut from traditional electrophysiological recordings to imaging based assays have been employed to achieve this end (Fig. 42–2).

Figure 42–2.

Approaches to measure physiological activity in 2D neuronal and brain organoid cultures. A–C. Available techniques and their advantages and disadvantages. These approaches range from more traditional electrophysiology such as single electrode-based (more...)

Classical electrophysiological techniques such as patch-clamp recordings have been successfully used to identify cell intrinsic changes in membrane currents and synaptic activity that may contribute to an epilepsy phenotype (Liu et al., 2013b; Tidball et al., 2020). This approach has the advantage of providing granular and cell-type specific data, but it is also highly labor intensive and usually limited to recording from one or, at best, a few neurons simultaneously. Extracellular recordings permit simultaneous data gathering from larger populations of neurons, including both single-unit activities (i.e., extracellular action potentials) of individual neurons as well as lower frequency local field potentials (LFPs) generated by networks of interacting cells. The LFP can include contributions from neurons and non-neuronal cells such as astrocytes (Buzsaki et al., 2012; Obien et al., 2014). In the case of 2D hPSC-derived cultures, this is typically achieved by utilizing tissue culture plates embedded with multielectrode arrays (MEAs), as monolayer cultures lack sufficient depth to be compatible with more traditional recording tools such as glass electrodes. The MEA format also allows for additional advantages such as the potential for repeated measurements over time as the recording apparatus typically couples with the array at an external interface, thus preserving cellular sterility, the ability to test drugs in a multi-well format, and the ability to both stimulate and record. To achieve this, hPSC-derived neurons are typically plated directly on the MEA. Using this approach Odawara et al. were able to track the electrophysiological dynamics of hPSC-derived neurons for over 1 year, and demonstrated hypersynchronous bursts in response to pro-convulsant drugs and rescue of this with antiseizure medications (Odawara et al., 2016). Using a variation of this approach, another group generated hPSC-derived neuronal aggregates, and over the course of 3 weeks in culture, used a nine-electrode MEA apparatus to describe patterns of network connectivity and development, such as the onset of increased synchrony (Izsak et al., 2019).

Optical microscopic approaches offer another powerful set of methods to study physiological and network activity in hPSC-derived cultures. The most well-established of these is through fluorescence-based calcium indicators of neuronal electrical activity. These indicators are predicated on the principle that neuronal activity, both action potential generation and synaptic transmission that activates neurotransmitter release, results in increases in intracellular calcium stores (Grienberger and Konnerth, 2012). This change in calcium concentration can be detected by indicators that change conformation in response to calcium binding, thus resulting in a change in emitted fluorescence following excitation. Historically, the first reliable indicators were chemical calcium indicators (Grienberger and Konnerth, 2012). More recently, improvements in genetically encoded calcium indicators (GECIs) based on chimeric fluorescent proteins have become more widely used. GECIs can be specifically targeted to select neuronal populations, are amenable to chronic recordings, and have excellent signal to noise ratios (Dana et al., 2019). The development and improvement of laser scanning microscopy, including confocal microscopy and two-photon imaging techniques, has been an equally important set of technical innovations that has resulted in widespread use of these indicators primarily for in vitro imaging of rodent neuronal cultures and in vivo rodent brain imaging. These techniques have been adapted for characterizing the calcium dynamics of epilepsy associated hPSC-derived 2D neuronal cultures (Avazzadeh et al., 2019; Krey et al., 2013; Marchetto et al., 2010).

The aforementioned technical developments have popularized the use of calcium indicators; however, major limitations of this approach include that the calcium signal is only a proxy for the electrical activity of interest, and calcium indicators provide limited or no information on hyperpolarizing input or subthreshold inputs (Knopfel and Song, 2019). While improved deconvolution algorithms, coupled with newer GECIs, have been shown to estimate activity with a relatively high degree of fidelity to simultaneously recorded spontaneous or induced neuronal spiking, the other concerns with calcium indicator imaging are inherent limitations (Grewe et al., 2010; Pnevmatikakis et al., 2016; Vogelstein et al., 2010). This has motivated the development of optical voltage indicators, which directly measure neuronal electrical activity. Optical voltage imaging is challenging for a number of reasons, including, but not limited to, the need to target a small (outer) region of the plasma membrane and yet generate sufficient photons to discern activity changes, the requirement for millisecond fast imaging speeds, and the ability to integrate reporters into the plasma membrane without causing cellular stress or damage (Bando et al., 2019). Although organic voltage-sensitive dyes have been in existence for several decades, the more recent development of genetically encoded voltage indicators based on similar protein engineering principles as GECIs has resulted in improvements in targeting specificity, signal-to-noise ratio, and cell toxicity that has allowed for improved neuronal and neural circuit imaging (Piatkevich et al., 2019; Storace et al., 2015). While voltage indicators have yet to be applied to interrogate activity in epilepsy-associated, hPSC-derived neuronal cultures, this emerging technology has the potential to provide novel insights into circuit pathology in epilepsy by combining the temporal fidelity of traditional electrophysiologic recording techniques with the spatial information conferred by calcium indicator imaging.

In order to gain more detailed information about the contribution of specific cell types to circuit activity, virtually all of the optical and electrophysiological assays described above can be combined with optogenetic or chemogenetic tools to specifically manipulate neuronal populations of interest. Optogenetics utilizes bacterial opsins that respond to stimulation by specific wavelengths of light by opening ion channels, thus converting photons directly into current (Deisseroth, 2015). The delivery of these engineered proteins to specific neuronal populations allows for temporally precise and cell specific excitation and inhibition of neuronal activity. Chemogenetics in this context refers to a parallel approach using either engineered G protein–coupled receptors or ion channels that respond only to a specific and otherwise inert compound to gain chemical, instead of light-based, control of cellular activity (Atasoy and Sternson, 2018; Magnus et al., 2019). Optogenetic and chemogenetic tools have been used to manipulate and interrogate circuit activity in ex vivo and in vivo rodent models of epilepsy (Shiri et al., 2017; Yekhlef et al., 2017; Zhou et al., 2019), including their use to activate or suppress grafted hPSC-derived interneurons (Cunningham et al., 2014; Upadhya et al., 2019). These tools also are well suited to a broad range of in vitro cellular applications, including hPSC-derived cultures.

Genetic Epilepsy Modeling Using 2D hPSC Cultures

Brain Organoid Models

While human stem cell–based 2D cultures have shown value in epilepsy modeling, these systems have significant limitations, including physiological, functional, and molecular differences from in vivo human tissue (Bershteyn and Kriegstein, 2013). In particular, it is challenging to achieve complex neural circuit formation in 2D cultures for a number of reasons, including limited neuronal (in particular interneuron) diversity, maximum in vitro culture times that are usually less than 3 months, and the lack of a layered 3D structure. Given the importance of neural circuit dysfunction in the pathogenesis of epilepsy, this is a significant shortcoming of 2D models. Brain organoid cultures consist of 3D structures containing diverse populations of neurons and glia. Organoids can also recapitulate some aspects of the layered cytoarchitecture of brain and exhibit complex neural circuit activities (Lancaster et al., 2013; Samarasinghe et al., 2021; Sun et al., 2019; Trujillo et al., 2019; Watanabe et al., 2017). Moreover, although organoids cannot achieve the level of cellular or structural complexity of in vivo brain, they nonetheless present with some unique advantages compared to commonly used rodent models. For example, hPSC-derived organoids contain an enlarged subventricular zone (SVZ), with an outer subventricular zone (oSVZ) not found in rodent brain. Importantly, human brain organoids also contain a sizable population of the primarily oSVZ-associated progenitor population known as outer radial glial cells (oRGs; also known as basal radial glial cells). The oRGs are thought to be responsible for the massive expansion of human cortex, and they are relatively abundant in gyrencephalic species such as humans and nonhuman primates but are largely absent in lisencephalic species such as mice (Hansen et al., 2010; Martinez-Cerdeno et al., 2012). Given the known impact of genetic variants on epilepsy, organoid models provide a unique platform to determine the effects of these variants on cortical development and expansion.

In order to generate brain organoids, hPSCs are aggregated in suspension using a number of different approaches such that they differentiate and self-organize into 3D brain-like structures. While the available protocols for organoid generation continue to expand in this highly dynamic field (and are reviewed in detail elsewhere; Di Lullo and Kriegstein, 2017), the general approaches can be divided into two broad categories. In one set of so-called intrinsic protocols, stem cells are allowed to differentiate in the absence of specific patterning factors. The resulting organoids typically contain a wide assortment of cell types, including forebrain cortex, hindbrain, and retinal cells (Lancaster et al., 2013). In the second category of approaches are protocols that utilize the timed application of patterning factors to direct differentiation into specific brain-like regions. For example, SMAD inhibition is used in multiple protocols to limit mesoderm and endoderm formation in favor of ectoderm (Kadoshima et al., 2013; Pasca et al., 2015; Qian et al., 2016). Often, SMAD inhibition is combined with other morphogens or small molecules to generate dorsal forebrain-like brain organoids that consist primarily of glutamatergic (excitatory) neurons (Bagley et al., 2017; Kadoshima et al., 2013; Pasca et al., 2015). These dorsal “cortical” organoids notably lack inhibitory interneurons. Similarly, SHH pathway agonists both in the presence and absence of WNT inhibition have been used to generate ventralized organoids with a large population of inhibitory interneurons (Bagley et al., 2017; Watanabe et al., 2017). In addition to dorsal and ventral forebrain organoids, this approach of directed differentiation has been used to generate a growing list of additional regional specificities, including hypothalamus, cerebellum, hippocampus, and others (Muguruma et al., 2015; Qian et al., 2016; Sakaguchi et al., 2015; reviewed in Di Lullo and Kriegstein, 2017).

Modeling Genetic Epilepsies and Their Effects on Cortical Network Function With Brain Organoids

A number of studies, primarily using directed differentiation of human forebrain cortex-like organoids, have utilized 3D culture approaches to determine the effects of epilepsy-associated genetic variants on various anatomical measures, including organoid cytoarchitecture and organization, cell expression patterns, and neuronal migration. This work has been reviewed elsewhere (Baldassari et al., 2020; Nieto-Estevez and Hsieh, 2020). Recent studies have additionally examined neural circuit activities generated by organoids. One of the first to do so utilized microelectrode arrays to document the generation of neural oscillations that increased in complexity with age (Trujillo et al., 2019). Measurements were performed in cortical organoids that were retained in culture from 2 to 10 months. In a more direct study of epilepsy, Sun and colleagues generated a cortical organoid model of Angelman syndrome resulting from a mutation in the ubiquitin ligase UBE3A gene (Sun et al., 2019). Angelman syndrome is associated with intellectual disability and lifelong seizures. Cortical organoids examined after 120–150 days in vitro demonstrated evidence of neuronal hyperexcitability by patch-clamp recording, and calcium indicator imaging revealed hypersynchronous discharges in the mutant organoids that were not observed in isogenic controls.

Both of these studies involved directed differentiation of forebrain cortex-like organoids, which would not be expected to contain a significant number of inhibitory interneurons. Interestingly, transcriptomic analyses by Trujillo and colleagues did reveal the spontaneous expression of an inhibitory interneuron population by 10 months in culture (Trujillo et al., 2019), which may partly explain the increased complexity of oscillatory activity observed by these authors at later time points. In contrast, and as expected, Sun et al. observed sparse inhibitory interneurons in their culture conditions (Sun et al., 2019). Given the importance of interneurons and excitatory-inhibitory connectivity in neural circuit activity in general and the pathophysiology of epilepsy in particular, recently developed approaches to integrate these populations within a single organoid are of particular relevance to organoid modeling of epilepsy. Birey and colleagues pioneered an innovative and elegant approach to achieve this by generating “assembloids” (also referred to as organoid “fusions”) in which glutamatergic neuron-rich dorsal and interneuron-rich ventral organoids are separately generated and then combined by placing the two structures adjacent to each other such that they physically fuse together (Birey et al., 2017). This protocol appears to recapitulate in an organoid the in vivo process of tangential migration of interneurons, whereby ventrally born interneurons and interneuron progenitors migrate and integrate into dorsal cortex. The assembloid approach has since been used to model circuit activity in an organoid model of RTT resulting from mutations in the MECP2 gene (Samarasinghe et al., 2021). In this study, MECP2 mutations were shown to result in hypersynchrony by calcium indicator imaging as well as both a loss of gamma oscillations and increased high-frequency activity (previously known as high-frequency oscillations [HFOs]) by extracellular recording of local field potentials. These authors also leveraged the assembloid approach to specifically identify ventral organoid derived interneurons, as opposed to dorsal excitatory neurons, as the primary drivers of the hyperexcitable phenotype.

Challenges and Future Directions

The use of hPSC-based epilepsy models presents a number of challenges. One issue is line-to-line variability of iPSC lines both within and between subjects. This variability requires the use of several lines each from multiple patients to show the consistency of any given finding. The use of isogenic controls generated by CRISPR gene editing also helps to address potential concerns that findings are due to genomic variability rather than the gene variant being studied. Another issue is cellular maturation. Because iPSC-derived models can only be grown in 2D culture conditions for up to a few months, they often represent a more immature state of the target cell, an issue common to many different tissues derived from hPSCs. This limitation is at least partially alleviated by brain organoid models, which can be grown in culture for extended periods. Both 2D and 3D hPSC cultures also are not yet able to generate the complete repertoire of human brain region-specific cell types or the entire repertoire of neuronal subtypes (discussed more below), which may cause difficulties in modeling certain human brain disorders. iPSC-derived neural cultures also do not recapitulate the complex anatomical connectivity that is present in vivo (See Figure 42–3).

Figure 42–3.

Advantages and disadvantages of hPSC-derived 2D and 3D models of brain disorders. Patient-derived, genome-edited, and isogenic control hPSCs may be differentiated into 2D monolayer cultures or 3D brain organoids (Lam et al., 2017). In hPSC-derived 2D (more...)

Brain organoid models share some of these challenges and have some important limitations that are unique to this approach, some of which can be addressed with new protocols or considered experimental planning. These challenges can be divided into three major categories: (1) limitations related to cellular diversity, (2) limitations in cellular maturity, and (3) potentially high levels of variability in organoid generation. Limitations in cellular diversity include a lack of specific non-neuronal cell types such as immune cells or vascular endothelial cells/components in most organoid preparations, as well concerns about limited expression of certain neuronal subtypes, including specific classes of interneurons such as fast spiking PV-expressing cells. In response to the lack of non-ectoderm-derived cell types, various approaches are being developed, including independent generation and application of particular cells types such as microglia (Bodnar et al., 2021; Xu et al., 2021), as well as entirely novel methods like “organoid on a chip” models in which microfluidic devices are used to both mimic vascular activity and the paracrine/autocrine influence of endothelial cells (Cho et al., 2021). The potential lack of neuronal diversity may be addressed by other innovations such as the dorsal-ventral assembloid approach, which appears to enhance interneuron diversity (Birey et al., 2017; Samarasinghe et al., 2021).

The fidelity of brain organoids to many aspects of human brain development, including their temporal fidelity to in vivo development, can pose a challenge when attempting to model diseases that typically manifest in advanced age. As such, organoid models are clearly well suited to disease processes that manifest early in development. This does not preclude successfully modeling neurological diseases that occur later in life since preclinical manifestations may be present in cellular composition or circuit activity well in advance of phenotypic expression. Approaches to induce “premature ageing” such as seeding an organoid with pathologic tau or α-synuclein fibrils or expressing genes associated with age-related neurodegeneration are additional approaches that can make organoids a more tractable approach for modeling neurological diseases that manifest later in life (Bowles et al., 2021).

A final concern with organoid-based models is variability in organoid preparations both within and between laboratories. Part of this variability stems from the diversity of protocols for organoid generation. As the field matures, this issue is being addressed by specific multi-investigator cross-institutional efforts to compare multiple organoid approaches and select for consistency and reliability, by increasing awareness of the key factors in stem cell and organoid maintenance and culture that are critical to consistency, and more organically by the emergence of dominant protocols that are becoming commercialized and increasingly used by many laboratories. Within laboratory batch-to-batch variability in organoid preparation is also a concern. This can be addressed by careful batch testing and bulk purchase of critical media components such as knockout serum and N-2 supplement. Careful quality control by immunohistochemistry or PCR for appropriate differentiation of organoids as well complementary accounting of batch-to-batch variability in physiological measures, as recently published (Samarasinghe et al., 2021), will provide further assurance of the reliability of organoid modeling. Even with these caveats, patient hPSC-derived 2D and 3D cultures have provided extensive insight into cellular mechanisms of genetic epilepsies, and their unique properties make them extremely useful for disease modeling, regenerative therapies, and drug/toxicity screening.

Challenges notwithstanding, the rapidly advancing field of stem cell biology suggests that future developments will expand the use of hPSC models in epilepsy research. Although at present hPSC models are predominantly used for genetic epilepsies, it is conceivable that this approach will enable the human cellular modeling of acquired epilepsies. For example, the ability to direct hPSC-derived 2D or brain organoid cultures into hippocampus-like tissue (Sakaguchi et al., 2015; Sarkar et al., 2018) should offer a paradigm for studying temporal lobe epilepsy. Moreover, hPSC-derived neurons are starting to be used as an in vitro model of traumatic brain injury (Shi et al., 2021), and this approach may prove useful in the future to study posttraumatic epilepsy mechanisms or for therapy development.

Another use of brain organoid methods currently under development is as a human tissue model for screening teratogenic effects of current and potential future antiseizure medications. Brain organoids, along with future protocols to generate spinal cord organoids, should aid in our understanding of the mechanisms underlying neural tube formation and, when used as a screening assay, decrease the incidence of neural tube defects from antiseizure medications and other drugs. Another future prospect for 3D cultures that should impact epilepsy research is the development of improved region-specific organoid patterning combined with assembloids. In addition to the existing excitatory/inhibitory cortical organoid fusions described above, it is likely that thalamic-cortical (Xiang et al., 2019) and hippocampal-subpallial fusion organoids will soon be applied to study epilepsy. Lastly, progress in the incorporation into brain organoids of cell types normally found in the brain but not present using current brain organoids protocols, including microglia, endothelial cells, and meningeal cells, should improve the fidelity of modeling epilepsy with human cells. The rapid pace of progress in hPSC-based disease modeling suggests that this field will contribute to both mechanistic understanding of and precision therapy for the epilepsies.

Conclusions

hPSC models provide a renewable source of human cells that are being increasingly applied to the study of genetic epilepsies. Many hPSC-based models have already provided important advances in the mechanistic understanding of a number of DEEs. In addition to insight into ictogenic substrates, these stem cell approaches may be applied to study neurodevelopmental comorbidities and complications of epilepsy, including SUDEP. Moreover, both 2D and 3D hPSC models of genetic epilepsy have great potential for guiding precision epilepsy therapy. All of these applications, especially when used in combination with genetically modified animal models, offer the prospect of improvements in the care of persons with epilepsy.

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