Advances in Cell Culture Technologies

## **Chapter 1**

## Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture

*Marcus Ground, Karen Callon, Rob Walker, Paget Milsom and Jillian Cornish*

## **Abstract**

Valvular interstitial cells (VICs) are the primary cellular component of the heart valve. Their function is to maintain the structure of the valve leaflets as they endure some three billion beats in the course of a human lifespan. Valvular pathology is becoming ever more prevalent in our ageing world, and there has never been a greater need for understanding of the pathological processes that underpin these diseases. Despite this, our knowledge of VIC pathology is limited. The scientific enquiry of valve disease necessitates stable populations of VICs in the laboratory. Such populations are commonly isolated from porcine and human tissue. This is achieved by digesting valve tissue from healthy or diseased sources. Understanding of the many VIC phenotypes, and the biochemical cues that govern the transition between phenotypes is essential for experimental integrity. Here we present an overview of VIC physiology, and a tried-and-true method for their isolation and culture. We make mention of several biochemical cues that the researcher may use in their culture media to ensure high quality and stable VIC populations.

**Keywords:** valvular interstitial cell, heart valve, cell culture, myofibroblasts, aortic stenosis

## **1. Introduction**

The four valves of the mammalian heart are delicate, complex structures that ensure unidirectional flow of blood through the heart's four chambers. Two atrioventricular valves, the tricuspid and mitral, sit between the atria and ventricles. During systole, these valves are closed to prevent the backflow of blood into the atria. The semilunar valves—so named for their half-moon shaped leaflets—sit between the ventricles are their outflows: the pulmonary trunk and the aorta. During systole, these valves are open, allowing blood to exit the heart. During diastole, they slam shut, preventing the blood from flowing backwards. The valves themselves comprise two main structural features: the valve leaflets (also called 'cusps') and the fibrocartilaginous ring onto which they are mounted. The leaflets are beautifully thin sheets of tissue organised into three distinct layers each with a definite function. The upstream

layer, the 'ventricularis' of the semilunar valves and the 'atrialis' of the atrioventricular valves is rich in radially oriented elastin, allowing the leaflet to hinge open and closed. The middle layer, the 'spongiosa', is rich in glycosaminoglycans, forming the bulk of the matrix and structurally linking the superficial layers. Finally, the downstream surface, the 'fibrosa', is rich in circumferentially oriented collagen that confers strength against the mechanical load of the blood pressure when the valve is closed [1, 2].

Two key cell types are responsible for maintaining the structure and function of the heart valve leaflet. Valvular endothelial cells (VECs) form a confluent monolayer on the surface of the valve. These cells are phenotypically distinct from endothelial cells elsewhere in the vasculature.

The second key cell type, and the object of this chapter, is the valvular interstitial cell (VIC). These are effectively fibroblasts that populate the extracellular matrix of the valve leaflets.

Functionally, VICs are responsible for maintaining the matrix of heart valve over the course of growth and development. This is no mean feat—the heart valves must endure some three billion beats over their lifetime.

## **2. Valvular interstitial phenotypes and their physiology**

There are five recognised phenotypes of valvular interstitial cell: embryonic, quiescent, progenitor, activated, and osteoblastic.

#### **2.1 Embryonic VICs**

Embryonic VICs are responsible for valvulogenesis—the formation of the valves from the endocardial cushion during fetal development. The precise detail of how this occurs in not well understood, but several key features of these embryonic VICs are known. Firstly, they are highly proliferative, a feature which distinguishes them from other VIC phenotypes [3]. Second, is that these cells are endothelial in origin, having undergone the endothelial-to-mesenchymal transition coordinated by a complex sequence of cell signallers. Notch and TGF family ligands (such as BMP-2 and VEGF) induce TGF-β and Wnt/β-catenin signalling which supresses endothelial-type gene expression in the endothelial cells, and instead encourages the cell to become embryonic VICs, a mesenchymal cell type [4–6]. Once this transition is complete, the maturing valve and its embryonic VICs 'striate' the matrix into the three aforementioned layers. Precisely how this occurs is unknown, but recent evidence suggests that the differentiation is aided by the haemodynamic forces of the fetal circulation [7].

#### **2.2 Progenitor VICs**

The VIC population is not a static one. Rather, a subpopulation of VICs present in the valve tissue in adulthood express surface markers of progenitor cells. Studies on healthy porcine valves show that approximately 5% of VICs express ABCG2, NG2, or SSEA-4 [8]. The origin of these progenitor cells in not entirely clear. Evidence suggests that some of these progenitor cells migrate to the valve from the blood stream—studies in mice have shown that bone marrow progenitor cells marked with green fluorescent protein appear in the valve, and assume the role of qVICs by secreting collagen 1 [9]. Other researchers suggest that another source of pVICs are VICs already residing within the valve, and are 'activated' in response to injury [3].

### **2.3 Quiescent VICs and activated VICs**

Quiescent VICs (qVICs) maintain the matrix throughout adult life [3]. They are neither very metabolically nor mitotically active. Instead, their role in maintaining the valve matrix is by sensing mechanical forces, and responding by differentiating into activated VICs (aVICs). The activated VIC is a myofibroblastic phenotype—characterised by the synthesis of ECM proteins and α-smooth muscle actin (a contractile protein) [10]. The differentiation from qVIC to aVIC is the key feature of mechano-regulated growth of this tissue from fetus to adult. As the heart grows, the pressures across the valve leaflet increase, introducing undue stress to the ECM. qVICs resident in the ECM can sense this stress, and differentiate into aVICs that then secrete matrix proteins in order to strengthen the matrix, until such a time when the matrix is able to support the increased pressure. At this time the 'stress' stimulus is lost and the aVIC then dedifferentiates back into a qVIC. It should be no surprise that the fetal heart valve contains a high proportion of aVICs, and that the healthy adult valve contains nearly no aVICs [11].

This mechano-regulation of heart valve growth and development relies on the ability of the qVIC's mechano-sensing ability. This is achieved through three surface protein classes: integrins, cadherins, and mechano-sensitive ion channels [12]. Integrins bind the VIC to the matrix, forming a continuation of the cytoskeleton intracellularly with the ECM extracellularly. Cadherins are transmembrane proteins that bind other cells—in the case of VICs, cadherin 2 is upregulated in qVICs, and cadherin 11 in aVICs [12]. Mechano-sensitive ion channels, particularly calcium channels, also enable mechano-sensation by altering ion concentration within the cell in response to stress-induced deformation [13]. Collectively, these three mechanosensing modalities induce a transcriptional change within a qVIC in response to stress, which causes aVIC differentiation, upregulation of ECM component synthesis, and the production of α-SMA [10].

### **2.4 Osteoblastic VICs**

Osteoblastic VICs (obVICs) are, as the name suggests, bone-producing cells that arise from VICs in the context of heart valve calcification. Up to 13% of the cellular content of end-stage diseased valve leaflets are positive for osteoblast markers RUNX2 and ALP, and histology of diseased leaflets shows evidence of lamellar bone [14]. Exactly where the obVICs come from is not certain. Some authors suggest that obVICs arise exclusively from qVICs in the context of age-related valve disease [10], while others suggest that obVICs come from a restricted range of susceptible VICs or perhaps pVICs [14].

## **3. Valvular interstitial cells in health and disease**

### **3.1 From qVIC to aVIC and back again: dynamic reversibility of VIC activation**

The key feature of VIC physiology that underpins normal growth and development, and the pathogenesis of heart valve disease, is the reversibility of the transition between qVIC and aVIC [15]. This reversibility (depicted in **Figure 1**), or the failure thereof, is perhaps best illustrated by two examples: growth and development by fetal heart valves, and the development of calcific aortic valve disease. Each involves substantial activation of VICs and the subsequent production of matrix.

**Figure 1.**

*Shows the 'dynamic reversibility' of VICs as they transition between their quiescent and activated states, as well as the key protein expression markers associated with each phenotype.*

The fetal heart valve is an immature structure—not yet required to bear the forces of systolic blood pressure while the fetal circulation, with its shunts, remains active. Once a baby is born, however, those shunts close, and the semilunar valves must immediately assume their role in preventing blood from flowing backwards into the ventricles from their respective outflows. The transition from immature fetal valve to fully competent valve requires several histological changes. First, the immature, loosely organised ECM network of the early valve must become more structured and load-bearing. Secondly, the resident VICs must proliferate in order to adequately populate the valve matrix, and will need to change their phenotype to meet the increased mechanical needs of the growing valve [11].

The immature valve matrix has a lower proportion of elastin than the adult valve, and a much greater proportion of glycosaminoglycan [11]. Interestingly, the proportion of collagen—the primary ECM component of the valve by weight—appears to be fairly stable from the first trimester through to adulthood. What does change markedly, however, is the degree of 'organisation', assessed by various authors using methods such as picrosirius red staining viewed under polarised light [11], or by quantifying the degree of hydroxyproline cross-linking [16]. In the first instance, collagen fibre thickness and alignment is significantly increased from fetus to child, and in the second, hydroxyproline crosslinking is significantly elevated. The net effect is substantial increase in the strength of the valve matrix.

How do the resident VICs achieve this? The answer lies in the presence of aVICs. The cell population of the fetal valve is up to 30% activated VIC, as determined by α-SMA staining [11]. These cells contribute to the changes in valve ECM makeup by simultaneous upregulation of matrix proteins (collagen, for example) and matrix metalloproteases (MMPs) [17]. The combination of upregulated ECM structural proteins and the enzymes that break them down is 'remodelling'—i.e., the overall proportion of collagen remains static, but the maturation into strong, organised bundles proceeds.

In the fetal valve, the laydown of collagen in the correct orientation is driven by the mechano-sensing ability of the resident VICs. But as we age, there are situations where the dynamic reversibility of qVIC and aVIC phenotypes is upset, and the resulting products of VIC activation lead to disease. The prescient example is calcific aortic valve disease (CAVD)—the most common valvular pathology of the developed world, with prevalence rates of up to 15% in those over 75 years old [18]. In the early phase of this disease, aVIC activation and the accumulation of structural *Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

ECM components mimics the changes seen in fetal valves, only in this circumstance, the interruption of the 'dynamic reversibility' of aVICs is lost, and the valves become stiff and non-compliant [19, 20]. Valves affected by CAVD are characterised by the overproduction of collagen [21] and the dysregulation of MMP expression [22].

The 'dynamic reversibility' of VIC phenotypes and the mechanical properties of the valve matrix are a true 'chicken and egg' problem. VIC activation invariably leads to stiffening and matrix remodelling (either physiological in the case of the fetal valve, or pathological in the case of CAVD), which, in turn, results in a stiffer matrix, which then serves a signal for qVICs to activate [23].

#### **3.2 VIC calcification: murky origins**

The origin if the calcification in heart valve disease is not perfectly understood. What is known is that VICs can calcify via two distinct mechanisms: calcific nodule formation and obVIC differentiation (depicted in **Figure 2**) [10]. The relative contributions of these two mechanisms to heart valve disease, or if the two mechanisms are independent of one another is not known.

Current evidence suggests that calcific nodules arise from aVICs, and are in fact calcified apoptotic bodies left after the cell dies. This type of calcification is termed 'dystrophic' to distinguish it from the matrix calcification caused by obVICs [24]. Where the obVICs arise from, is not clear. Some have suggested that obVICs arise from the qVIC population independently, and in response to changes in matrix mechanics or the accumulation of calcium [25], while others suggest that obVICs arise from aVICs in a manner that depends 'activating' signalling [26].

Support for the idea that the obVIC and aVIC calcification are distinct pathways is seen in the apparent changes in α-SMA expression, which is markedly reduced in presence of osteogenic medium and markedly increased in the presence of TGF-β.

#### **Figure 2.**

*Shows the two phenotypes of VIC calcification: the dystrophic calcific nodules that occur after aVIC apoptosis, and the ossification of obVICs. What is unclear is if the obVIC phenotype arises directly from qVICs or aVICs.*

Conversely, qVICs cultured in osteogenic medium will demonstrate upregulation in key osteoblast markers: ALP and OCN [25]. To further confuse the true origin of obV-ICs, the same pattern of gene expression occurs when valvular endothelial cells are cultured in osteogenic media [27].

### **3.3 VIC signalling molecules**

The various signalling molecules that induced change in VICs has been extensively reviewed elsewhere [10]. There are, however, three molecules that commonly feature in culture medium preparations, and the physiology that underpins their use is included here.

#### *3.3.1 TGF-β*

Perhaps the most well characterised stimulus for VIC activation is transforming growth factor β (TGF-β) [28–34]. TGF-β, or more specifically the isoform TGF-β1 is one member of a much larger 'TGF-β superfamily', which includes 30 species such as bone morphogenic proteins (BMPs), growth and differentiation factors, and various other peptides key to growth and development [35]. In adulthood, TGF-β1 is involved in a myriad of cell functions, from cell fate determination, cell migration, response to injury, and ECM regulation [34, 36, 37]. In the context of VICs, the key function of TGF-β is its ability to regulate activation of VICs and maintain homeostasis of the valve ECM by induction of the aVIC phenotype. As well as structural proteins, ECM contains large amounts of 'latent' signalling molecules—signallers bound up with latency-associated peptides. These peptide complexes are hidden from cells, and so do not enact their functions until they are cleaved from the matrix during proteolysis following injury [36, 38]. TGF-β, as a promoter of ECM production, serves as a 'switch' in heart valves: when the valve is unable to cope with a tension force, microscopic injury releases TGF-β from the matrix, which in turn signals to cells to secrete more ECM, until such time that the matrix can withhold the tension and the TGF-β switch is turned off. It is through delicate control of TGF-β stimulated ECM secretion that a valve grows and remodels throughout the changing haemodynamic environment from fetal development to adulthood [6, 39].

TGF-β acts on cells via several biochemical pathways. The most well characterised pathway with respect to VIC physiology is the 'canonical TGF-β' pathway, or Smad pathway [34, 38, 40]. TGF-β forms a homodimer in the extracellular space and binds two type 2 TGF-β receptors (Tgfbr2), which then recruit two type 1 TGF-β receptors (Tgfbr1) forming an activated receptor-ligand complex [39]. The activated Tgfbr1 is then able to phosphorylate intracellular messenger proteins Smad2 and Smad3, which form a complex with Smad4 and move to the nucleus as transcription factors to activate the associated genes [39, 41]. It is this process that is responsible for the activation of qVICs to myofibroblasts and for the development of dystrophic nodules [3].

Non-Smad pathways initiated by TGF-β are termed 'non canonical'. These include the ERK/MAPK pathway [41], which has been shown to regulate key aspects of aVIC expression in a manner independent of the canonical TGF-β pathway [42].

#### *3.3.2 FGFs*

The fibroblast growth factor (FGF) family comprises 23 distinct proteins, 18 of which are ligands to various FGF receptors, effecting a wide range of cellular

#### *Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

functions: from migration to proliferation to cell survival [43, 44]. Of interest to the VIC researcher is FGF-2, also called basic FGF (bFGF), so named for its discovery in the alkaline fraction of pituitary extracts [45]. FGF-2 has been implicated in numerous biological processes, from cell migration and fate determination to pathological signalling in cancer metastasis [46]. Most interesting is that while this molecule is present and active throughout the body, a distinct physiological role has not been identified. Homozygous FGF-2 knockout mice survive perfectly well and give birth to healthy young.

FGF-2's signalling pathway first involves heparin binding [46]. Two heparin-FGF2 complexes then bind two FGF receptors (FGFR) to form a stable dimer, which then activates intracellular tyrosine kinase domains and effects changes in gene expression through a number of signalling cascades, among them MAPK and AKT pathways [43].

FGF-2 has been shown to be particularly important in regulating the quiescent VIC phenotype [34, 47–49]. The mechanism of FGF-2's effect appears to be multifaceted: both repressing the canonical TGF-β pathway by blocking Smad nuclear translocation [48], and by activating of the Akt/mTOR pathway [34, 49]. The downstream effects of this activation have been shown to oppose the effect of TGF-β. FGF-2 appears to promote the qVIC phenotype: VICs treated with FGF-2 have a spindleshaped morphology, and express more vimentin and less α-SMA [17].

#### *3.3.3 Nitric oxide*

The function of nitric oxide (NO) in the vasculature is understood to be a local control mechanism for maintaining blood pressure and perfusion. First, NO is produced in endothelial cells by endothelial nitric oxide synthase (eNOS) in response to stretch or biochemical stimuli [50]. Then, NO is able to rapidly diffuse into the surrounding tissue by virtue of its small size, where it binds guanylyl cyclase (a cytoplasmic receptor) [51] and leads to a myriad of downstream effects including relaxation of the vascular smooth muscle [50]. With respect to heart valves, NO production by VECs and its subsequent action of VICs is an important paracrine regulator of valve cell function [3, 10]. More importantly, dysregulation of NO signalling is implicated in the development of heart valve disease [52–55]. In the initial stages of CAVD, eNOS is upregulated on the aortic-side endothelium [56], where it contributes to pathological neovascularisation of the valve tissue [57]. The early eNOS response is a protective mechanism to temper the proliferation of VICs, but as overexpression of eNOS continues, resistance to NO develops—a phenomenon termed 'NOS decoupling' [58].

With respect to VIC activation, exogenous NO inhibits the formation of calcific nodules in porcine VIC cultures under TGF-β stimulation [59]. Culturing VICs in media supplemented with DETA-NONOate—a biologically inert molecule that spontaneously releases two NO molecules in a predictable manner [60]—limits the formation of new nodules.

This effect is not purely induced by exogenous NO, rather it can be replicated by co-culturing VICs with NO-producing VECs [61]. Under these conditions, VECs can inhibit the formation of aVIC-related calcific nodules in a manner that is reversed by addition of a NO scavenger. This suggests that of the multitude of paracrine signals that travel between the endothelium and the interstitium, NO is paramount in maintaining the valve in its quiescent state.

## **4. Sources of valvular interstitial cells**

There are a number of VIC sources for a researcher's consideration. No one option is without issue. Perhaps the easiest source of VICs is from animals. Pigs are by far the most common source of VICs for research [61–81], as they are very easily obtainable from meat processing facilities. As a biproduct of industry, obtaining and using porcine VICs requires little or no ethical oversight [82]. Another notable benefit is the anatomical similarity between the pig and human heart. This is of particular importance as the mechanical and structural forces regulate VIC physiology and gene expression [23, 68], and so choosing a structurally similar heart approximates a physiologically similar VIC. In a similar vein, pigs housed for research can be induced into diseased states for investigation of a particular pathology. For example, pigs fed on high-cholesterol diets experience atherosclerotic lesions, though no porcine model of calcific valve disease has been trialled to date [52, 83]. It should be noted that VICs derived from meat industry pigs can be considered healthy qVICs by definition their valves are not experiencing rapid growth, nor has age-related disease begun to influence their phenotype. This fact is either a boon or an issue for the cell culturist. Those interested in studying the development of valve disease might be eager to use a healthy population of qVICs, while those searching for therapeutic options for persistent aVIC or obVIC presence will likely have to choose another source. In our experience, porcine VICs are easily obtained and fairly forgiving when it comes to cell culture. Cultures derived from digested porcine valves are adequately proliferative and less sensitive to aVIC formation when cultured on hard plastic.

Sheep may represent a promising animal alternative for obtaining VICs liable to calcify. Sheep exhibit an elevated calcium metabolism compared to humans, and this has led to their use as an experimental model of heart valve calcification [84]. Several authors have made use ovine VICs for experimentation [85, 86].

Human VICs are the best option for studying heart valve disease, as none of the animal models approximate our valvular pathologies perfectly. The most ideal source is a young, healthy adult human heart valve. Cells from this source, in theory, are almost entirely qVICs. However, there are a number of constraints that make sourcing healthy human tissue difficult.

First is availability—unlike other cell types, VICs are not sourced from a healthy tissue that is routinely removed from people undergoing surgery (as it the case in, for example, adipose tissue removed during breast cancer surgery), and suitable heart valve tissue from younger donors is contained within a heart that is likely more suitable for heart transplantation. To complicate matters further, in the instance where a young donor heart is not suitable for transplant, the heart valve may be harvested as a 'homograft' (a heart valve transplant).

The second constraint is time. While VICs are relatively robust in the absence of nutrient supply [10], good laboratory practice dictates that cells should be isolated and incubated as quickly as possible. Very rarely are healthy heart valves available within temporal or spatial proximity to a laboratory. Despite these constraints, various researchers have made use of healthy human VICs, typically from donor hearts not suitable for either transplantation or homograft use [47, 87].

A much more commonplace source of human VICs is those isolated from patients undergoing valve replacement surgery. Valve replacement surgeries are very common—our experience at Auckland City Hospital saw an average of 5 valves available per week. The obvious problem with this source is the inherent disease state of the cells within the valve. Though our protocol above recommends removal of the more

*Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

calcified portion of the valve, the pathology is not localised at the calcification, and altered stiffness characteristics across the whole leaflet ensures that all cells present will have some degree of pathological programming [53].

The diseased state of these cells may actually be useful for researchers concerned with the study of aortic calcification—the most common valvular pathology in the developed world. Many examples of diseased valves used for this purpose are available in the literature [47, 87–102].

The key consideration to address with diseased human VICs is ethics. Patients must consent to their tissue being harvested and used in research, and protection of the tissue and any data generated must be ensured by the researcher. In addition, many cultures have unique requirements for the disposal or return of human tissue [103].

## **5. Isolation protocol**

This isolation protocol was developed by our laboratory for the isolation of human VICs from patients undergoing valve replacement surgery [104]. An abbreviated flow chart is included in **Figure 3**.

**Figure 3.** *Shows and abbreviated flow chart of steps 7–18.*

## **5.1 Consumables**


## **5.2 Hardware**


*Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*


## **5.3 Method**

Prior to tissue collection:


In the operating room:

7.Make sure to bring the warmed aliquot of PBS for tissue collection. When the surgeon removes the valves, ensure the scrub nurse or assisting surgeon offers the container in a sterile manner, and that the tissue is added to the container and sealed shut all while in the sterile field. Return to the laboratory as quickly as possible

In the laboratory:


## **6. Quality control considerations**

Regardless of cell source, the goal for VIC culture is a population of phenotypically stable cells (i.e. few aVICs) with as little endothelial contamination as possible.

### *Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

To assess phenotypic stability, we encourage researchers to plate out a sample of their VIC culture and assess it for features of myofibroblast activation in aVICs. Typically, we >5000 cells from our digestate and plated these in a 24-well plate, and left to grow for up to 6 days. We then assessed for the presence of aVICs first by visual inspection on phase microscopy, and then by α-SMA staining. aVICs differ from pVICs in their morphology. On phase microscopy, qVICs are slender, spindle-shaped cells with a central body surrounded by two or three thin processes. Their area rarely exceeds 5000μm2 , and their aspect ratio (the ratio between their major and minor axes) should be greater than 3 [17]. aVICs on the other hand are much larger, with areas often exceeding 10,000μm<sup>2</sup> . Their shape is much more circular, with a large cell body and loss of the thin processes. Similar cell area measurements were found by Porras and colleagues, who used FIB media to prevent (or reverse) aVIC formation in porcine cultures [72].

We suggest staining for α-SMA and vimentin to assess the degree of aVIC presence (an example of which can be seen in **Figure 4**). aVICs have dense bands of α-SMA running across the cell, terminating in adhesion complexes that link the intracellular force generated by this protein to the ECM [105]. True qVICs will have very little α-SMA staining, if at all.

Many researchers will quantify aVIC numbers by using α-SMA positivity, either by fluorescent intensity measurement or thresholding numbers of positive cells [47, 48]. In our experience, there is little utility in attempting to quantify the degree of VIC activation using α-SMA staining or phase microscopy, as all VICs will show some α-SMA expression and variation in morphology. More intensive methods for quantifying α-SMA include western blotting and real time PCR [47, 106]. These are comparative methods (i.e., they require two groups), and so are of little use in the 'quality control' stage. Experimentally, we have found that expression of ACTA2 (α-SMA's gene) and a number of other aVIC-associated genes correlate with the

#### **Figure 4.**

*Shows two cultures of VICs: one rich in qVICs on the left, and another rich in aVICs. Cells are stained for vimentin (green) and α-SMA (red). Cells were cultured for four days on plastic. Scale bars represent 100 μm.*

#### **Figure 5.**

*Shows a quality control procedure performed on a VIC culture (left). The yellow arrow indicates VEC contamination. This was performed alongside a VEC culture as a positive control. This culture was grown from 'Digest 1' aspirate. Both cultures were grown for four days on plastic. Scale bars represent 100 μm.*

presence of 'aVIC-looking' cells on phase microscopy and α-SMA staining. We suggest that researchers subjectively assess the appearance of their VIC cultures on day 6 post-isolation.

The second main feature of VIC quality control is endothelial contamination (**Figure 5**). Despite the initial digest and scraping we suggest in the protocol, it is impossible to remove all VECs. Endothelial contamination can run as high as 10% [10]. In our experience, VIC cultures generated by the means outlined above will produce cultures with less than 1% von Willebrand factor (vWF) positivity. To assess VEC contamination, we suggest first saving and plating the digestate from digest 1 in a 6-well plate or smaller (see step 11) for use as a positive control. In the same plate, we suggest reserving a small volume of the VIC suspension from digest 2, and culturing both for up to 6 days. We then suggest staining the two wells for an endothelial marker. The choice of marker is not especially important: VECs express vWF, CD31 (PECAM) and CD34, and are negative for α-SMA [74, 107].

## **7. Culture conditions**

The majority of the literature has VICs cultured on flat plastic in Dublecco's modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS). We have had good results with this formulation. There are, however, a number of other culture media that should be considered. Alternatives to DMEM include M199 medium [29, 108] and DMEM-F12 [73, 109].

Good laboratory practice stipulates that isolated cells should be cultured in such a way that preserves their *in vivo* phenotype as faithfully as possible. And so, it befalls the VIC researcher to consider the inherent aVIC promoting properties of chosen media. The supplementation of media with 10% FBS is standard practice for the culture of any fibroblast [110]. However, careful thought should be put into the inherent content of peptides that stimulate VIC activation. TGF-β, for example, is present in FBS in concentrations ranging from 10 to 20 ng/mL. TGF-β is a strong promoter

*Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

of VIC activation, and several authors have mitigated its effect by using a 2% FBS medium supplemented with insulin as a mitotic agent. This so-called 'FIB' media, first used by Latif and colleagues [47, 72], is also supplemented with 10 ng/mL FGF-2—an agent shown to reduce VIC activation in favour of the quiescent phenotype. We have demonstrated that VICs cultured in FIB media show significantly downregulated expression of aVIC-associated genes (α-SMA in particular), and a morphology more typical of qVICs (greater aspect ratio, and reduced total cell area) compared to VICs cultured in the standard DMEM preparation [17]. The inherent ability of FIB medium to suppress aVIC formation can be potentiated further by culturing VICs on collagencoated plates [72].

Induction of the obVIC phenotype requires osteogenic medium. This invariably contains ascorbic acid, dexamethasone, and β-glycerophosphate [28, 109, 111], supplemented in some instances with TGF-β or BMP-2 [89].

Seeding densities vary widely in the literature, and will differ depending on the surface on which the VICs are grown. On hard plastic or glass, we have found VICs to be rather sensitive to contact inhibition and myofibroblastic activation. We suggest that VICs are seeded at a density of 10,000 cells/cm2 for experiments running for less than five days. At this density, cell growth reaches a plateau on day 4 [104].

## **8. Conclusion**

VICs are the specialised fibroblasts that populate the human valve and maintain its structure and function throughout our lives. The increasing incidence of valvular pathology means that there is no better time to gain understanding about the physiological and pathological processes that affect these cells. Despite this, our understanding of VIC health and disease is in its infancy. Culture of VICs is an important part of the scientific exploration of heart valve disease. Several key methodological considerations need to be addressed for the jobbing scientist:


## **Acknowledgements**

This work was supported by the Green Lane Research and Educational Fund and the Health Research Council of New Zealand.

## **Author details**

Marcus Ground1,2\*, Karen Callon2 , Rob Walker1 , Paget Milsom3 and Jillian Cornish<sup>2</sup>

1 University of Otago, Dunedin, New Zealand

2 University of Auckland, Auckland, New Zealand

3 Auckland City Hospital, Auckland, New Zealand

\*Address all correspondence to: marcus.ground@auckland.ac.nz

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

*Perspective Chapter: Valvular Interstitial Cells – Physiology, Isolation, and Culture DOI: http://dx.doi.org/10.5772/intechopen.112649*

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## **Chapter 2**

## Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent Stem Cells

*Qiang Li*

## **Abstract**

Human pluripotent stem cells (hPSCs), which include human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs), hold immense potential for various biomedical research in both academic and clinical applications. This chapter provides a comprehensive review of culturing techniques for hPSCs, covering two-dimensional (2D) adherent culture, three-dimensional (3D) suspension culture, and the utilization of hydrogel scaffolds in 3D hPSC culture. Furthermore, it explores the application of advanced 3D cell manufacturing techniques to facilitate the production of large quantities of high-quality hPSCs, catering to the needs of advanced biomedical applications. By addressing these topics, this chapter aims to present a comprehensive overview of diverse cultivation methods and their wideranging applications in hPSC research, encompassing fundamental studies and advanced biomedical investigations.

**Keywords:** hPSCs, 2D & 3D cell culture, hydrogel, scaffold, cell manufacturing

## **1. Introduction**

Human pluripotent stem cells (hPSCs) include human embryonic stem cells (hESCs) [1] and human induced pluripotent stem cells (hiPSCs) [2, 3]. These cells are characterized by unique features such as indefinite self-renewal and the remarkable capacity to differentiate into all the cell types in the body. Therefore, the hPSCs represent invaluable cell sources for fundamental stem cell research and a wide range of biomedical applications, including cell therapies, tissue engineering, drug discovery, and disease modeling [4–8]. In addition to hPSCs, tissue-derived adult stem cells can serve as valuable resources for stem cell therapies. These multipotent stem cells, including the stem cells isolated from fetal tissues, mesenchymal stem cells (MSCs), and hematopoietic stem cells (HSCs), exhibit the capacity to differentiate into restricted cell types within their respective lineages [9]. The two-dimensional (2D) adherent culture of hPSCs is a commonly employed in research laboratories for fundamental stem cell studies. The 2D adherent cell culture methods can only produce

the cells in limited quantities. However, all biomedical applications require significantly larger quantities of cells [10]. For example, the treatment with one patient of myocardial infarction typically necessitates approximately 109 cardiomyocytes. Similarly, the treatment of one diabetic patient requires around 109 β cells. In the case of blood transfusions, approximately 1012 red blood cells are needed. Moreover, the engineering of a human-size liver and the screening of a library with one million compounds at once typically require approximately 1010 cells [11, 12]. Researchers have explored suspension culture methods [13–19] to scale up the production of hPSCs and their derivatives. Furthermore, alternative methods have been developed to enhance the scalability of hPSC culture, including cell microencapsulation within hydrogels such as alginate [20–23] and thermoreversible hydrogels [24, 25]. These methods utilize a supportive matrix that closely mimics the natural extracellular environment, effectively preventing the formation of excessive cell agglomerations and reducing exposure to shear forces generated in the dynamic suspension culture. As a result, they exhibit high efficiency in expanding hPSCs [21, 24]. To facilitate the production of hPSCs on a large scale, implementing three-dimensional (3D) cell manufacturing processes is crucial. Extensive research has been dedicated to implementing 3D cell manufacturing processes for large-scale production [26–30].

## **2. Two-dimensional (2D) adherent and three-dimensional (3D) suspension culture of hPSCs**

The hPSC culture media comprises vital nutrients, growth factors, and supplements that promote cell viability, proliferation, and maintenance of pluripotency. Initially, hPSC culture media contained serum and conditioned media [1, 31]. Currently, xeno-free and fully chemically defined media, such as mTeSR [32, 33] and Essential 8 (E8) [34], are commonly used for the cultivation and propagation of hPSCs. These media formulations eliminate the reliance on animal-derived components, offering a well-defined and controlled environment that supports the optimal growth and maintenance of hPSCs. For example, the E8 medium comprises eight essential components: DMEM/F12, insulin, FGF, L-ascorbic acid, selenium, TGFβ, transferrin, and NaHCO3. DMEM/F12 serves as the base medium, providing a nutrient-rich foundation. Insulin and FGF2 play crucial roles in promoting the survival and proliferation of hPSCs. L-ascorbic acid acts as a stimulant for cell proliferation. Selenium is vital for the sustained expansion of hPSCs. TGFβ is incorporated to enhance pluripotent marker expression and ensure consistent long-term culture stability. When combined with a ROCK inhibitor (RI, Y27632) [35], transferrin improves initial survival and supports high cloning efficiency. NaHCO3 is used for pH adjustment of the medium. These xeno-free and fully defined media efficiently support the expansion of hPSCs, enabling the transition of these cells from basic laboratory research to clinical applications.

The commercial hPSC cell lines, including hESCs and hiPSCs, can be purchased from WiCell Research Institute, the American Type Culture Collection (ATCC), or the European Collection of Authenticated Cell Cultures (ECACC). The equipment and supplies for hPSC maintenance include cell culture incubator, biosafety hood, 4°C refrigerator, −20°C freezer, −80°C freezer, liquid nitrogen dewar, inverted microscope, water bath, centrifuge, cell-freezing container, cryovials, pipette set with various tips, and different sizes of tubes (e.g., 1.5, 15, and 50 mL) along with standard cell culture plates (e.g., 6-well cell culture plates or 24-well cell

*Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent... DOI: http://dx.doi.org/10.5772/intechopen.113860*

culture plates) and low-attachment cell culture plates (e.g., low-attachment 6-well cell culture plates or low-attachment V bottom or U bottom 96-well cell culture plates). Additionally, the following materials and reagents are required to culture hPSCs: Essential 8™ medium (Gibco, or STEMCELL Technologies), Matrigel matrix (Thermo Fisher Scientific, Corning Life Sciences, or VWR), Rock inhibitor (Tocris Bioscience, Selleck Chemicals, or MilliporeSigma), Accutase (STEMCELL Technologies, MilliporeSigma, or Invitrogen), EDTA (0.5 M), pH 8.0, RNase-free (Thermo Fisher Scientific, VWR, or Cayman Chemical), DPBS (no calcium, no magnesium, Gibco, MilliporeSigma, or VWR), Dimethyl sulfoxide (Sigma-Aldrich, Thermo Fisher Scientific, or VWR), Penicillin-streptomycin solution (Corning Life Sciences, MilliporeSigma, or VWR), and 70% ethanol.

#### **2.1 2D cell culture methods**

The 2D adherent culture of hPSCs in cell culture plates or Petri dishes is a commonly employed method in laboratories for fundamental stem cell studies such as cell expansion and differentiation (**Figure 1**). The culture substrate plays a significant role in supporting hPSC attachment and proliferation, maintaining pluripotency. hPSCs were initially cultured on a layer of feeder cells, which refer to mitotically inactivated mouse embryonic fibroblast (MEF) or human fibroblast feeders that support hPSC growth and maintenance [1, 37]. Due to the significant variability and irreproducibility associated with the feeder-dependent methods, a feeder-independent culture protocol was developed [33], in which the hPSCs were cultured on extracellular matrix (ECM) rather than the feeder cells. Matrigel, an Engelbreth–Holm–Swarm mouse sarcoma tissue extract, has emerged as one of the most extensively utilized extracellular components for feeder-free culture of hPSCs in laboratories [5, 31, 33, 36]. Matrigel is a partially chemically defined and xenogeneic substrate encompassing collagens, laminin, and various other chemical compounds [38]. Therefore,

#### **Figure 1.**

*2D cell culture of human pluripotent stem cells [33, 36]. (a–c) the 2D cell culture is prepared for passaging. The colony and cell morphology when observed through 2× (a), 4× (b), and 10× (c) objectives, respectively. The center of the colonies appears denser and brighter and cells at the center of the colony may appear smaller. Scale bar: 100 μm [33]. (d-f) Following plating, the cells quickly adhere to the plate within a few minutes (d), spread within 2 hours (e), and subsequently grow as colonies within 24 hours (f). Scale bars: 50 μm [36].*

xenogenic-free, chemically defined ECM, such as laminin [39, 40], vitronectin [41], and synthetic surfaces [42], was developed to facilitate hPSC expansion and differentiation on the 2D adherent substrates. hPSCs cultured in 2D systems provide valuable insights into their molecular characteristics and differentiation potentials.

The 2D adherent cultures are widely used in laboratories for hPSC expansion and differentiation. The procedures involve several steps to ensure successful cell seeding, cell growth, and maintenance. These steps include (1) thawing frozen cells, (2) seeding cells on ECM-coated culture plates, (3) cell maintenance, (4) cell passaging, and (5) cell freezing for storage. Throughout these processes, it is important to exercise caution to minimize the risk of cell contamination and stress. Adhering to aseptic techniques, maintaining proper culture conditions, and strictly following established protocols will help to preserve the quality of hPSCs, especially for maintaining their pluripotency for long-term culture. Detailed protocols for culturing hPSCs on a 2D surface are provided below, offering step-by-step guidance and best practices.

*E8 medium preparation:* The E8 basal medium should be stored in a refrigerator at 4°C. Aliquot the E8 supplements into 1-mL tubes and store them in a freezer at − 20°C or − 80°C. To prepare the E8 full medium, mix 48.5 mL of the E8 basal medium with 1 mL of the E8 supplement and 0.5 mL of the penicillin-streptomycin solution.

*Matrigel preparation:* Aliquot the Matrigel into small volumes (e.g., 200 or 500 μL) and store them in a − 80°C freezer for a long time. The Matrigel was left in a 4°C refrigerator overnight to dissolve, resulting in a viscous liquid. To aliquot the Matrigel, place the Matrigel and all other required tubes in an ice bag in the biosafety hood in case the Matrigel forms the hydrogel during the procedures. To prepare the Matrigel coating medium, dilute the Matrigel with cold DMEM/F12 basal medium to a final concentration of 100 μg/mL. To coat the cell culture plate, add 1 mL of the Matrigel coating solution into one well of the 6-well plate. Place the cell culture plate at room temperature in the biosafety hood for at least 1 hour or place it in the incubator at 37°C for at least 30 minutes.

#### *2.1.1 Thawing frozen cells and seeding on the 2D surface*


*Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent... DOI: http://dx.doi.org/10.5772/intechopen.113860*

	- 1.Before passage, add RI to each well at a final concentration of 10 μM. Then, put the cells back in an incubator for culture for at least 1 hour. Note: RI treatment can significantly improve cell viability after passage.
	- 2.Remove the cell culture medium and add 1 mL of DPBS (without Ca2+ or Mg2+) to each well. Gently shake the plate to ensure proper distribution, then carefully remove the DPBS.
	- 3.Add 1 mL of 0.5 mM DETA solution to each well. Allow the cells to remain undisturbed at room temperature for approximately 5–7 minutes.
	- 4.Remove the EDTA solution from each well. Using a 1-mL pipette, gently pipette the cells up and down with 1 mL of E8 full medium supplemented with 10 μM RI.
	- 5.Transfer 200 μL cell solution to a new Matrigel-coated well and add 2 mL fresh E8 full medium supplemented with 10 μM RI.
	- 6.Place the cells in the incubator. To ensure even distribution of the cell clusters within the wells, gently shake the cell culture plate back and forth several times.
	- 7.On the second day, replace the medium with 2 mL of E8 full medium. Continue culturing the cells for an additional 3–4 days. Observe the cell morphologies using an inverted microscope and allow the cells to reach approximately 80% confluency before the following passage. Change the medium daily.
	- 1.Prepare the cell freeze medium by combining 10% DMSO with 90% E8 full medium supplemented with 10 μM RI. Note: Add the DMSO into the E8 medium slowly, drop by drop.
	- 2.The EDTA-based procedure described above was used to dissociate the cells into cell clusters. Note: Freezing cell clusters instead of single cells can enhance the efficiency of cell recovery.

In summary, 2D cell culture methods are widely used in laboratories for basic stem cell research. For hPSC maintenance, it is essential to exercise caution to minimize the risk of cell contamination and cell stress. Preserving the high quality and pluripotency of hPSCs requires strict adherence to aseptic techniques, meticulous maintenance of culture conditions, and rigorous adherence to established protocols. Despite their extensive use, 2D culture approaches have inherent limitations. These systems fail to fully replicate the intricate 3D microenvironment *in vivo*, which hampers accurate modeling of cell-cell interactions, morphogenesis, and organ development. As a result, researchers have developed 3D suspension culture methods to address these questions.

## **2.2 3D suspension culture methods**

In laboratories, 3D suspension culture methods have also emerged as valuable tools for studying hPSCs' expansion and differentiation in a more physiologically relevant environment (**Figure 2**). This approach allows cells to interact with neighboring cells and the extracellular matrix, promoting cell-cell interaction and tissue-like organization. The 3D culture enables the formation of more complex structures, such as embryoid bodies (EBs) or organoids, which are self-organized 3D tissues with multiple cell types. Organoids derived from hPSCs have demonstrated remarkable cellular organization and functionality, offering great potential for cell therapies, disease modeling, and drug screening. The 3D suspension culture is widely used for hPSC and hPSC-derived organoid cultures [8, 43–48]. In this system, the cells can be cultured in low-attachment cell culture plates or spinner flasks. The detailed protocols for culturing hPSCs on 3D suspension are provided below, offering step-by-step guidance and best practices.

## *2.2.1 2D to 3D suspension passage cells*


*Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent... DOI: http://dx.doi.org/10.5772/intechopen.113860*

#### **Figure 2.**

*3D suspension culture of human pluripotent stem cells [19]. (a–e) Single cells were seeded in the 3D suspension culture on day 0 (a); the formation and growth of cell spheroids from day 1 to day 4 (b–e). (f) Undesirable cell spheroid agglomeration formed in the center of the dish. Scale bars: 500 μm.*

	- 1.Calculate the cell concentration and dilute the cells in E8 full medium supplemented with RI. Seed around 3000–6000 cells per 200 μl medium per well in the low-attachment 96-well plate.
	- 1.Transfer 1×105 cells into a single well of the low-attachment 6-well plate. Add 2 mL of fresh E8 full medium supplemented with 10 μM RI.
	- 2.Place the cell plate onto a shaker inside the incubator for suspension culture. The cells will form small cell spheroids within 24 hours.
	- 3.Passage the cells on days 5–7. The final concentration of the cells in the 3D suspension culture is about 1–2 million cells per mL.

## *2.2.4 3D to 3D suspension passage cells*


Biomedical applications require large quantities of cells of high quality. To scale up the production of hPSCs, researchers have explored suspension culture methods using trimethyl ammonium-coated polystyrene microcarriers [14] or Matrigel-coated microcarriers [13]. However, it has been observed that the cell expansion rate tends to decrease gradually over long-term culture on microcarriers, and there can be challenges associated with detaching the cells from these microcarriers [13, 14]. To address these challenges, researchers have developed 3D suspension culture techniques for hPSCs without using microcarriers [15–19]. hPSCs have been successfully cultured as floating aggregates in static conditions using low-attachment cell culture plates or in suspension culture with regular agitation of the culture vessel, such as

*Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent... DOI: http://dx.doi.org/10.5772/intechopen.113860*

spinner flasks. Currently, the 3D suspension culture of hPSCs and hPSC-derived organoids in spinner flasks is also commonly applied in laboratories.

#### **2.3 3D culture with hydrogel scaffolds**

Hydrogel scaffolds like alginate [20–23], or thermoreversible hydrogels [24, 25] have been developed to culture hPSCs in a more physiologically relevant microenvironment. These hydrogel scaffolds can be used as physical barriers to isolate cells from agglomeration and hydrodynamic stresses generated from the dynamic suspension. These methods enhance the scalability of hPSC culture and exhibit high efficiency in expanding hPSCs [21, 24]**.** For example, a thermoreversible hydrogel-based cell culture system has been developed to expand and differentiate multiple cell lines of hPSCs [24, 25]. In this system, single cells were seeded within this thermoreversible PNIPAAm-PEG hydrogel matrix (**Figure 3**). As the hydrogel is very soft, the single cells can clonally grow into uniform spheroids within 3–5 days by deforming the scaffolds to create spaces. The hydrogel eliminating the cell agglomeration and hydrodynamic stresses could significantly improve the culture efficiency. The hydrogel scaffold enabled serial expansion of hPSCs with good cell viability, growth rate, yield, and purity. Moreover, after expansion, the hydrogel can be easily dissolved into liquid by adding cold DPBS to the well. Then, the cell spheroids can be easily harvested and centrifuged for the following passage procedures. This method provides an efficient, defined, scalable, and good manufacturing practice-compatible approach for hPSC production and differentiation [24].

Additionally, alginate hydrogels offer tunable mechanical properties and can encapsulate hPSCs to form 3D structures. Alginate hydrogel tube cell culture system

#### **Figure 3.**

*3D culture of hPSCs in thermoreversible PNIPAAm-PEG hydrogels [24]. Phase images depict cell morphologies following culture in PNIPAAm-PEG hydrogel. Single-cell seeding densities were 2.5×105 , 1.0×106 , or 2.5×106 cells per mL, respectively, and cultures were maintained for 4 days in either mTeSR or E8 medium with 1-day or 4-day RI. Scale bar: 250 μm.*

(AlgTubes) [21–23] provides cells with a culture microenvironment that aligns with their physiological needs. In this system, hPSCs are seeded in microscale alginate hydrogel tubes (**Figure 4**). Within 24 hours, these hPSCs can form small cell clusters, enhancing cell viability post-seeding. The diameter of the tubes can be adjusted to approximately 200–400 μm, allowing nutrients and metabolic waste to diffuse freely through the hydrogel tube walls. The design of AlgTubes represents a paradigm shift in cell culture efficiency, as evidenced by improvements in various aspects such as growth rate, viability, yield, and genetic and phenotype stability. Furthermore, AlgTubes can be easily adapted for scalable expansion in bioreactors, showcasing the high potential for producing hPSCs and their derivatives with exceptional quality and quantity, making them ideal for various biomedical applications.

## **2.4 3D cell manufacturing for advanced biomedical applications**

HPSCs hold great promise for a range of cutting-edge biomedical applications, all of which necessitate substantial quantities of high-quality cells [11, 12]. (1) hPSCs can differentiate into diverse cell types, such as cardiomyocytes and neurons, with applications in regenerative medicine and organ transplantation. For instance, treatment of myocardial infarction in a single patient often demands roughly 10<sup>9</sup> cardiomyocytes. (2) hPSCs can be derived from individual patients, paving the way

#### **Figure 4.**

*3D culture of hPSCs in Algtubes [21]. Phase images illustrate the cell morphologies during culture in Algtubes. Cells were seeded at densities of 1×106 , 2×106 , 5×106 , or 10×106 cells per mL and were cultured for 9 days. Within 24 hours, individual cells aggregated to form small clusters. These clusters subsequently grew into spheroids by day 5, which further evolved into fibrous cell masses by day 9.*

*Perspective Chapter: Two-Dimensional and Three-Dimensional Culture of Human Pluripotent... DOI: http://dx.doi.org/10.5772/intechopen.113860*

for personalized disease modeling and high-throughput drug screening. Notably, screening a library containing one million compounds at once typically demands around 1010 cells. (3) hPSCs offer the potential to produce substantial quantities of blood cells, including platelets. For example, approximately 1012 red blood cells are required for blood transfusions.

Advancing the mass production of hPSC-based therapeutics toward a scalable and cost-efficient process will facilitate widespread adoption in all biomedical applications. The field of 3D cell manufacturing has opened exciting possibilities for large-scale production of hPSCs and their derivatives. Developing the 3D cell manufacturing processes requires incorporating approaches that allow for the precise control of culture conditions, such as nutrient and oxygen supply, pH, and mechanical forces, as well as precise monitor the cell conditions, including cell viability, cell states, and cell function [26–29]. Extensive research has been dedicated to implementing 3D cell manufacturing processes using bioreactors for large-scale production of hPSCs [27, 30, 49, 50]. For instance, bioreactors such as the Cellspin Integra Biosciences spinner flask [51], the NDS Technologies Bioreactor [52], and the Able Bioreactor [53] have successfully generated clinically relevant cell numbers. While the agitation of culture in bioreactors can improve mass transport and reduce cell agglomeration, it can also introduce hydrodynamic stresses, such as shear stresses, which may negatively impact hPSC expansion and differentiation. The variations in the stirred-tank bioreactor are influenced by numerous factors, including the bioreactor design, such as impeller geometry and size, as well as vessel geometry and size. Therefore, there is a critical need for automated 3D cell manufacturing in advanced biomedical applications, enabling the large-scale production of hPSCs in accordance with current good manufacturing practices (cGMP) and strict safety assessment criteria. In summary, advancements in this field continuously broaden our comprehension of hPSCs and their wide-ranging potential in addressing diverse human diseases and industrial demands.

## **3. Conclusion**

This chapter has provided a comprehensive overview of the different culturing methods used in stem cell research and their biomedical applications. The 2D cell culture methods have been extensively employed in laboratories for fundamental stem cell research. Detail protocols for hPSC culture in 2D adherent substrates were summarized. Furthermore, 3D suspension culture methods have demonstrated their value in basic stem cell research and advanced biomedical studies. The utilization of hydrogel scaffolds, including alginate and thermoreversible hydrogels, has enabled the development of 3D culture systems for hPSCs expansion and differentiation. Additionally, the application of 3D cell manufacturing techniques has shown promise in producing large quantities of high-quality hPSCs for advanced biomedical applications. This chapter has offered readers a comprehensive understanding of the diverse culturing methods and their roles in stem cell research, ranging from fundamental studies to cutting-edge biomedical advancements. This knowledge serves as a foundation for readers interested in delving deeper into stem cell biology and biomedical applications, empowering them to make valuable contributions to the field.

## **4. Troubleshooting**

Problem 1: Matrigel forms hydrogel during handling.

Solution 1: Thaw the frozen Matrigel at 4°C overnight. Ensure all handling steps, including aliquoting and preparing the Matrigel coating solution, are performed while keeping the Matrigel on ice.

Problem 2: In 2D culture, cell colonies cannot be detached from the plate. Solution 2: Appropriately extend EDTA treatment time.

Problem 3: In 2D culture, there is a low cell survival rate after cell plating. Solution 3: Ensure RI is added before cell passage and continue including RI in the cell culture medium within 24 hours after passage.

Problem 4: In 3D suspension culture, lots of large cell aggregations form. Solution 4: Appropriately increase agitation rate.

## **Appendices and nomenclature**


## **Author details**

Qiang Li John A. Paulson School of Engineering and Applied Sciences, Harvard University, Boston, MA, USA

\*Address all correspondence to: qiangli@seas.harvard.edu

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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## **Chapter 3**

## Perspective Chapter: Investigating Cancer Tumor Microenvironment *In Vitro* – Co-Culture Studies on Adipocytes and Cancer Cells

*Ozge Rencuzogullari, Pelin Ozfiliz-Kilbas, Enes Bal and Burcu Ayhan-Sahin*

## **Abstract**

The tumor microenvironment increases the growth and invasion of cancer cells, makes classical chemotherapy applications inadequate, and is associated with a poor cancer prognosis. Recent studies reveal that cancer stroma supports tumor growth and metastasis and develops resistance to chemotherapy. *In vitro* co-culture techniques are widely used to study cross-talk between tumor microenvironment cells such as adipocytes, endothelial cells, fibroblasts, macrophages, and cancer cells. Co-culture techniques are classified into two main categories: indirect and direct methods. Transwell (indirect) co-culture of mature adipocytes with cancer cells has shown cancer cell viability, growth, proliferation, invasion, and metastases. This chapter covers the general methods of co-culture studies and will emphasize the results obtained on the co-culture of adipocytes and cancer cells.

**Keywords:** tumor microenvironment (TME), cell culture, co-culture, adipocytes, organoids

## **1. Introduction**

Cancer is a multidisciplinary disease that occurs when the balance between cell proliferation and death is disrupted in the direction of proliferation. After excessive proliferation, tumor mass invades vessels and metastasizes to adjacent tissues [1]. In other words, a tumor colony is a group of cells that have gained a higher proliferation rate and can invade other tissues and metastasis [2]. Due to its high metastasis capacity, although it changes according to the type, the mortality rate is high in many cases [3]. Cancer is named according to the tissue or organ it originates from. Indeed, this terminology is insufficient most of the time, and each cancer type is also subdivided in compliance with molecular marker, histological staining status, and cell type. About 200 cancer subtypes have been identified by the National Cancer Institute (NCI), but the exact number of subtypes is still a mystery. Subtyping in cancer cases is essential because the subtype affects the therapy model [4]. The accumulation

of genetic and environmental risk factors triggers the initiation of carcinogenesis. Although there are cancer-type-specific markers, KRAS, BRAF, PIK3CA, epithelial growth factor receptor (EGFR), p53, and c-myc are the most common mutated genes in various cancers [5].

Surgery supported by chemotherapy is still the primary treatment method for solid tumors. Still, the adverse effects of chemotherapy reduce the life quality of patients because of the toxicity of drugs to healthy cells and the tumor cells. Radiation therapy and immunotherapy can also be used for treatment. The main drawback of developing new therapies with minimal adverse effects is the intra- and peri-tumor heterogeneity [3]. The tumor contains highly proliferating cancerous cells and is surrounded by different types of cells and tissues, defined as a tumor microenvironment (TME). Recent studies concerning tumor biology showed that TME has a significant role in drug resistance, tumor development, and malignancy [6]. Intra-tumor heterogeneity also affects the classification and subtyping of tumors. In some cases, intratumor solid heterogeneity complicates the type and prevents identifying the case, whether a known subtype or a new one [4]. Therefore, it is essential to define the tumor microenvironment to assess the subtype, malignancy, and metastasis profile of a tumor and study in cultures for new therapeutic approaches.

This chapter summarizes the important concepts of TME and potential applications in cell culture. As TME is quite complex, this chapter focuses on the studies of TME in adipocytes and cancer cells. This chapter summarizes the components of TME and the developing cell culture applications that show great potential to model various cancers. We also summarize the co-culture applications in cancer research. This chapter also provides further insights into novel cell-culture-based TME studies.

## **2. Cellular and non-cellular components of tumor microenvironment (TME)**

TME, defined as the environment of a tumor that tumor cells interact with, is composed of cellular and noncellular compartments [7]. The cellular compartment contains mainly fibroblasts, endothelial cells, adipocytes, stem cells, stellate cells, and the innate and adaptive immune cells such as B lymphocytes, T lymphocytes, macrophages, dendritic cells, natural killer (NK) cells, neutrophils, and myeloid-derived suppressor cells (MDSC) (**Figure 1**) [8]. These cells are laid on a scaffold formed by an extracellular matrix (ECM), which includes proteins, proteoglycans, glycoproteins, and chemicals secreted by cellular compartments such as growth factors, cytokines, and chemokines [9]. Tumor cells interact with other cellular components, and the extracellular matrix provides a structure to hold all these elements together. Interactions between cellular immune cells and non-cellular components account for the tumor heterogeneity and complexity of TME and limitations for targeted therapies in patients [10]. As a result of this interaction, the cellular environment also shows some genetic and morphological changes. It causes conditions supporting the tumor's development, contributing to metastasis and drug resistance [11]. Due to its contribution to tumor development, metastasis, and drug resistance, the tumor microenvironment is an important topic that has been popular recently by researchers and offers a new perspective on cancer treatment [8].

It was first discovered in the 1890s when tumor and tumor microenvironment interacted, emerging with the theory "the seeds and the soil" of English surgeon Stephen Paget [12]. This theory that metastasis depends on communication between *Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies… DOI: http://dx.doi.org/10.5772/intechopen.113859*

**Figure 1.** *Illustration of cellular and non-cellular of the tumor microenvironment.*

certain cancer cells as the "seeds" and specific tissue microenvironments as the "soil" still retains its thought [13]. In further studies, it has become important to investigate the role of TME-associated cellular responses in carcinogenesis. Dysregulated immune responses are the main characteristics of inflammatory tissue microenvironment. TME exhibits an increase in inflammatory regulators such as tumor-infiltrating cells, T cells, macrophages, dendritic cells, and natural killer cells that are the cause of chronic inflammation to promote tumor progression, carcinogenesis, and metastasis [14]. A suppressed immune system, defined as immunosuppression, is one of the potential adverse effects of several cancers.

#### **2.1 Cellular components of tumor microenvironment**

#### *2.1.1 Fibroblasts*

Cells of connective tissue that are neither epithelial nor endothelial nor immune cells are referred to as fibroblasts [15]. Fibroblasts are substantial cells involved in extracellular matrix formation by synthesizing collagen, laminin, and fibronectin and thus contribute significantly to wound healing [16]. Resident fibroblasts, resident endothelial cells, and epithelial cells can be activated in tumorigenesis and differentiate into cancer-associated fibroblasts (CAFs) in response to growth factors and cytokines. Cancer-associated fibroblasts are the primary cell type in the stromal part of the tumor microenvironment [17]. These differentiated fibroblasts contribute to inflammation, angiogenesis, TME remodeling, epithelial-mesenchymal transition (EMT), metastasis and anti-drug resistance *via* producing matrix metalloproteinases (MMPs), growth factors (TGF-β, IGF, HGF, VEGF), cytokines and chemokines (IL-6, CCL7, CXCL2) and inhibiting natural killer cells, and cytotoxic T lymphocytes [9, 16, 18, 19]. Therefore, they promote tumor development and progression and are shown to be related to poor prognosis in pancreatic and breast cancers [17].

#### *2.1.2 Endothelial cells*

Endothelial cells are the chief cells that line the blood vessels, forming a cellular layer between the blood and the tissue, which organizes the interchange of materials from each side. Endothelial cells differentiate into tumor endothelial cells (TECs) in tumors that organize and play critical roles in immune system regulation, tumor cell proliferation, invasion, and metastasis by expression of inhibitory receptors for immune cells, vascular cell adhesion molecule 1 (VCAM1), and Notch1 signaling [10, 17].

#### *2.1.3 Adipocytes*

Adipocytes convert glucose and fatty acid to triglycerides in a mechanism called lipogenesis and, in this way, provide the energy demand of other cells [20]. In the tumor microenvironment, cancer-associated adipocytes (CAAs) are likewise energy storage cells [21] by supplying lipids to tumorigenic cells [10]. Apart from lipid and energy supply, they release various metabolites (fatty acids), hormones (leptin, adiponectin), proteins (collagen, VEGF, and matrix metalloproteinases), inflammatory factors (CCL2), and cytokines (IL-8), which are referred to as adipokines [9, 22, 23]. Through these adipokines, they regulate macrophage differentiation [9], immune cell homeostasis [23], immune evasion and tumor progression [9, 22], ECM remodeling [22], metastasis [24], and angiogenesis *via* increasing vascularization [25]. Many cancer types, such as breast, ovarian, colon, and prostate cancer, are primarily known for high adipocyte concentration; therefore, it is important to understand the interactions of the cells in TME with adipocytes to improve the therapeutics [26].

#### *2.1.4 B lymphocytes*

Although B lymphocytes' primary function is antibody secretion upon antigen recognition, in the tumor microenvironment, they activate mast cells and promote the production of cytokines when they interact with the antigens of tumor cells [27]. B lymphocytes have mainly antitumor functions, whereas when specific chemokines (CXCL13) and cytokines (IL-10) are present, they promote tumorigenesis and induce angiogenesis [9]. Thus, the role and function of B lymphocytes in the tumor microenvironment differ in accordance with the chemical secretions in the environment and tumor type, which affects the tumor's prognosis status [28].

#### *2.1.5 T lymphocytes*

T lymphocytes constitute one of the immune system's main groups of adaptive immune response. T lymphocytes have many subgroups, of which CD8<sup>+</sup> cytotoxic T lymphocytes (Tc) are responsible for the detection of tumor antigens and for destroying the tumor cells [18]. In the tumor microenvironment, as well as Tc lymphocytes, CD4<sup>+</sup> T helper lymphocytes (Th) also play essential promoting roles as both stimulation of Tc lymphocytes and B lymphocytes *via* secreting various cytokines [9]. On the contrary, another subgroup of T helper lymphocytes, Th17 lymphocytes, exerts double-sided effects. Th17 lymphocytes trigger inflammation and angiogenesis on the one hand and have anti-tumorigenic effects on the other [29].

The common immunosuppressor T cells, regulatory T cells (Tregs), are typically identified near tumors and suppress anticancer immune responses [30]. In normal cells, Tregs sustain self-tolerance mechanisms by preventing the proliferation of T lymphocytes and induction of immunity against self-antigens [31]. Studies indicated that the FOXP3+ CD4+ CD25+ subtype of Treg cells is instrumental in controlling immune responses in the tumor microenvironment. Tumor cells and macrophages in the tumor microenvironment secrete CCL22 chemokine, whose receptor, CCR4, was expressed by Treg cells and causes the accumulation of Treg cells in TME [30]. Tregs contribute to immunosuppression by secreting immune inhibitory cytokines such as transforming growth factor (TGF-β) and interleukin 10 (IL-10), which inhibit the anti-tumor activity of killer cells [27].

### *2.1.6 Macrophages*

Macrophages are the phagocytic cells in the innate immune system [32]. However, when recruited to the tumor microenvironment with certain cytokines or differentiated from circulating monocytes, they can acquire different phenotypes and are referred to as tumor-associated macrophages (TAMs) after that [17]. These different phenotypes are known as M1 anti-tumor macrophages and M2 pro-tumor macrophages, which are determined by the polarization status of the TAMs [9]. TAMs, particularly the M2 phenotype, are the cells that are the most infiltrated immune system cells in TME [33]. Besides, since they secrete extracellular matrix components, they participate in ECM remodeling [19].

## *2.1.7 Dendritic cells*

Dendritic cells are the antigen-presenting cells in the immune system that detect foreign antigens, present them to T lymphocytes and induce adaptive immune response [34]. In the tumor microenvironment, different subtypes of dendritic cells can be infiltrated. Classical dendritic cells and plasmacytoid dendritic cells are responsible for anti-tumorigenic response. On the other hand, tumor cells secrete CCL-2, CXCL1, and CXCL5, thus preventing dendritic cell maturation [9]. Further, dendritic cells promote Treg cell differentiation by secreting cytokines [10]. In this manner, dendritic cells' function depends on the chemicals secreted in TME [22].

#### *2.1.8 Natural killer (NK) cells*

Natural killer (NK) cells are the primary defense cells of the immune system against tumor cells [35]. When they encounter tumor cells in the bloodstream, natural killer cells essentially destroy them, whereas they cannot function effectively in the tumor microenvironment [22]. Although the dendritic cells recruit natural killer cells to the tumor microenvironment through various chemokines such as IFN-γ, they cannot correctly mature [9].

#### *2.1.9 Neutrophils*

Neutrophils, widespread in circulation [35], are the cells of the innate immune system responsible for the phagocytosis of dead cells or pathogens and for presenting antigens to adaptive immune system cells [9, 36]. In the case of tumorigenesis, neutrophils are recruited to the tumor site. In the early stages of tumorigenesis, these neutrophils are differentiated into the anti-tumor N1 subtype under the influence of IFN-γ [37, 38]. N1 subtype neutrophils release cytokines and reactive oxygen species, thus inducing apoptotic death of tumor cells [22]. In later stages of tumorigenesis, cancer-associated fibroblasts (CAFs) secrete TGF-β that promotes the differentiation of N2 subtypes and suppression of N1 subtype [28]. These N2 neutrophils secrete cytokines as IL-1β, growth factors as vascular endothelial growth factor (VEGF), and matrix metalloproteinases as MMP-9, in this way, stimulate inflammation, angiogenesis, matrix remodeling, and invasion [22, 28, 37].

### *2.1.10 Myeloid-derived suppressor cells (MDSCs)*

MDSCs secrete cytokines and growth factors; in this way, they suppress immune cells and induce tumor cell dissemination and renewal of tumor cells [27]. They originate from myeloid progenitor cells in the bone marrow and then migrate to the tumor microenvironment to support tumorigenesis, angiogenesis, and metastasis [9]. In TME, they also differentiate into tumor-associated macrophages (TAMs), which act together responsible for resistance to anti-angiogenic treatments [17].

#### **2.2 Non-cellular components of tumor microenvironment**

Non-cellular components of the tumor microenvironment involve mainly the extracellular matrix (ECM), which is different from the matrix of normal tissue in that it enables the neoplastic cells to metastasize. Collagen and fibronectin are the main components of ECM in solid tumors and ensure the endurance of the matrix [17]. Factors secreted from cellular components of the tumor microenvironment, such as cytokines, chemokines, proteases, growth factors, polysaccharides, and integrins, are also significant members, and these components provide the communication and interaction of cells with each other in TME [39, 40]. The importance of non-cellular components of the tumor microenvironment is that they organize not only the scaffold where the tumor mass resides but also regulate the polarization and the fate of the cellular components [41]. Non-cellular components of ECM also participate in drug resistance by providing a dense environment and disallowing the drugs to penetrate [17].

To summarize, the cancer tumor microenvironment is characterized by abundant stroma, extracellular matrix, stellate cells, cancer-associated fibroblasts, various immune cells, and cytokines are associated with a hypoxic environment, high

*Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies… DOI: http://dx.doi.org/10.5772/intechopen.113859*

vascularity and intense immunosuppression, proliferation, metastasis, and drug resistance of several cancers [36]. Various studies explained the direct interaction between tumor cells and immune cells, but the relationship between immune cells and TME needs to be clarified in the cell fate decision.

## **3. Cell culture applications studying TME**

To improve personalized therapy and the prognosis of cancer, extensive and varied investigations are performed from experimental to clinical applications. However, the complexity and heterogeneity of TME contribute to cancer development, progression, and drug resistance. Thus, developing cell culture applications is required to enlighten the interaction network of TME and cancer cells.

Since it was first introduced in 1907 for nerve fibers [42], cell culture techniques have been eased in cancer research to mimic the *in vivo* evolvement and reaction of tumors *in vitro* [43]. Classical two-dimensional (2D) cell cultures allow the cell to grow only in two dimensions; in other words, cells can only grow along the surface of the culture flask (**Figure 2**) [44]. These cultures may be conducted in adherent cultures and suspension cultures, of which the latter imitates the non-adhesive cells, such as hematologic malignancies [45], to resemble the natural environment in blood or lymph [43]. In adherent cultures, cells are attached to the plastic surface of flasks or dishes as a single layer and grow, interacting with the surface's neighboring cells and protein coating. These cultures provide a cost-effective, high-performance, and reproducible way to study *in vitro* [46]. For many years, these cultures have been used to investigate the cytotoxicity of drugs in

**Figure 2.** *Comparison of 2D vs. 3D cell culture.*

pharmacology studies, the migration and invasion characteristics of the cells in understanding the underlying mechanisms, the genome modifications in enlightening the genetic basis of cancer cells, and so on [47]. However, studying just one cell type in a 2D cell culture is nowhere near the interactions between cellular and non-cellular components of the tumor microenvironment [48]. Therefore, new techniques and applications must be developed to mimic the *in vivo* tumor microenvironment better.

#### **3.1** *In vitro* **co-culture applications for TME and cancer cell interactions**

The co-culture method is the direct or indirect interaction of multiple cell types in an identical culture environment [49]. Through the interaction of various cell types, co-culture systems regulate the properties of individual cells. Cellular mechanisms, including paracrine signaling, cell-to-cell communication, and modeling epithelialmesenchymal transition (EMT), are better estimated by co-culturing numerous cell lines into a single three-dimensional (3D) model [50].

Two types of cells are defined as "target cells" and "assisting cells" responsible for the co-culture environment. Target cells act to build up the engineered tissue and are responsible for the mechanical and metabolical functions of the tissue. Target cells are guided by assisting cells to exhibit various desired characteristics such as proliferation, differentiation, matrix production, or secretion [49].

Co-culture aims to form tissue-like characters of cells *in vitro* through cell–cell communication of multiple cell types in direct or indirect interaction. Co-cultures are used to explore three cell-to-cell communication patterns: cell–cell adhesion, including adherence, gap and tight junctions, cell-ECM adhesion, and paracrine signaling *via* soluble factors.

Co-culture techniques have been essential for researching the interaction between different cell types under diverse conditions, such as cancer cell growth, differentiation, proliferation, and metastasis. The complexity of the *in vivo* environment, where several cells interact and signal concurrently, is not fully mimicked by one cell type-based *in vitro* cell cultures due to inadequate prediction of signaling cascades. Therefore, several co-culture models have been improved to mimic TME, which are less expensive than *in vivo* animal models.

#### *3.1.1 Direct co-cultures*

Direct co-cultures allow two or more different cells to make physical connections with each other while mixing in the 2D or 3D cell culture environment. Mixed populations of monolayer cells on slides or in dishes are standard 2D cell culture methods [51].

Due to their simplicity and ease of control, 2D culture systems are acceptable as the easiest way to investigate cellular interactions, such as examining adhesion molecules, cytokine production, and juxtracrine signals. However, the composition of native tissues can be modeled using 3D culture systems, which is accomplished by culturing multiple cells in materials such as fibrin, agarose, collagen, or alginate [49].

#### *3.1.2 Indirect co-cultures*

Additionally, conditioned medium is widely employed in indirect co-cultures. Multiple cell types are separated through good inserts like Transwell or Boyden chamber permeable membranes or pre-conditioned media. Utilizing permeable

*Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies… DOI: http://dx.doi.org/10.5772/intechopen.113859*

membrane inserts allows secreted soluble factors to pass through the membrane, only enabling signaling *via* the cell secretome. Alternatively, utilizing previously created conditioned media from one of the co-culture cell types are used [52].

Indirect 2D co-culture systems allow a better explanation of specific cellular functions and features that occur specifically to cell types. However, in indirect 3D coculture systems, multiple cell types are separated using hydrogel or collagen matrix, which allows better mimicking of native tissue structures and eliminates the requirement for direct cell-to-cell contact between cell types [49].

Transwell plates [53, 54], microfluidic platforms [55–58], or solid supports like Petri dishes [59, 60], 3D scaffolds [61], hydrogels [62], or microarrays [63] are often utilized choices for co-culturing populations that are somewhat separated from one another (**Figure 3**). For co-cultures, many custom microfluidic devices have been

**Figure 3.** *Available technologies for co-culture [64].*

developed [65–69]. The possibility for high throughput in microfluidics is one of its key benefits. Microfluidic setups might not represent larger volumes as complexity rises [56], and this benefit may be undermined. Techniques used in monoculture modified for co-culture include cell movement experiments. Most traditional laboratory cultivation focuses on maintaining pure cultures of certain species [65, 70]. Although there have been recent developments that could be further utilized, such as microcarrier beads, the micro-Petri dish [71], diffusion chambers, dialysis reactors, and other techniques [70, 72], the cultivation of monocultures on agar plates and in liquid medium has always followed the same paradigm [71, 72].

Cell-to-cell direct co-cultures have some limitations and need more optimization steps. Therefore, indirect co-cultures are preferred as they provide reproducible results. However, indirect co-cultures cannot capture cell–cell adhesion, integrin, and notch-type signaling.

#### **3.2 Adipocytes-related cancer studies**

Since TME has become widely recognized as a significant contributor to tumor progression and cancer, it has gained importance to determine the roles of individual TME components in cancer. Adipocytes, endothelial cells, fibroblasts, macrophages, and muscle cells are cell types commonly studied in co-culture systems [73]. Adipocytes were subsequently recognized as a significant TME component in tumor development, angiogenesis, and metastasis through the secretion of soluble factors [74]. Adipocytes were once considered the body's energy reserves, but today, it is recognized that they have a role in various pathological processes, such as inflammation. In particular, they impact gene expression and cell motility in TME [75].

Mammalian epithelial cells are surrounded by an extracellular environment comprising adipocytes and other stromal cells, including fibroblasts, and endothelial and inflammatory cells. Adipocytes and adipocyte precursor cells are a substantial percentage of breast TME, which may play a significant role in the interaction between stromal and ductal epithelial cells and function in the secretion of growth factors, hormones, and adipokines [76].

According to the studies in cancer metastasis, it was determined that free fatty acids (FFAs) released from adipocytes through lipolysis may have been an energy source for cancer cells through mitochondrial fatty acid oxidation (FAO) [77]. In culture conditions, FFAs from adipocytes to cancer cells enhance FAO and support tumor cell proliferation, migration, and invasion. Elevating lipid load in adipocytes increases FFA transfer and is stored in tumor cells with lipid droplets, which is also associated with enhancing CAA-mediated proliferative effects in the TME [78].

In the first studies investigating the role of TME in breast cancer, the role of fibroblasts, macrophages, and other inflammatory cells was investigated. Still, the role of adipocytes was not recognized. Further studies in breast cancer have shown that the adipocyte-rich environment can promote cancer proliferation. It was shown that murine mammary carcinoma SP-1 cells injected subcutaneously into adiposerich tissues in mice promoted tumor growth and metastasis. Still, neither growth nor metastasis is observed in tissues with low levels of adipose. These findings were reinforced by the discovery that estrogen regulates the adipocyte-associated growth and proliferation of SP-1 cells [79, 80].

It has been determined by DNA microarray studies that adipokines secreted from adipocytes regulate cell survival, proliferation, invasion, and angiogenesis by inducing IGF2, FOS, JUN, cyclin D1, MMP1, ATF3, and NFkβ transcriptional target

#### *Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies… DOI: http://dx.doi.org/10.5772/intechopen.113859*

molecules. In addition, the induction of these genes was specific to factors released by adipocytes and cannot be observed in other breast stromal cells [74].

Previous studies showed that co-culture of differentiated murine preadipocytes with mammary tumor cells caused the delipidation of adipocytes with the increase of proinflammatory cytokines and matrix remodeling proteins while decreasing adipocyte differentiation markers, including resistin, adiponectin, and hormone-sensitive lipase (HSL). These adipocyte cells, triggered by tumor cells and changed phenotypically and functionally, are considered cancer-associated adipocytes (CAA) [81]. In addition, bidirectional contact between adipocytes and tumor cells changes the shape of adipocytes into adipocyte-derived fibroblasts, which are the parts of the CAAs and induce invasion, migration, and metastasis [82]. Studies have shown that interleukins produced by CAAs such as IL6, IL1β, TNFα, and MMP-11 are functionally responsible for supporting the invasion capacity of tumor cells and augmenting the proinflammatory TME properties. It was shown that pancreatic cancer cells linked to obesity release cytokines such as IL1β, which promotes infiltration of immune cells and leads to cancer progression and drug resistance [83]. Numerous pro-inflammatory cytokines, such as tumor necrosis factor alfa (TNF-α) and plasminogen activator inhibitor-1 (PAI-1), are produced by adipocytes and overexpressed in obese patients [84].

The methods by which stromal cells in TME alter tumor properties in different microenvironments are initiated to be clarified by several direct or indirect studies. Stromal cell-secreted factors such as matrix metalloproteases (MMPs) and growth factors regulate the cancer cell decision. As mentioned in previous studies, the interaction between adipocytes and cancer cells causes the change in the phenotype of adipocytes into CAA and delipidation with the overexpression of inflammatory cytokines but increases in the adipocyte differentiation markers [81]. Secretion of inflammatory cytokines such as IL-6, IL1β, or leptin promotes cancer development and metastasis in several cancer cells.

Even if the interaction between adipocytes and cancer cells is known, these cell– cell interactions are complicated, and it is also known that they cannot occur by the effect of a single secreted molecule or a signal. Therefore, molecular mechanisms of heterotypic signaling in bidirectional crosstalk between adipocytes and cancer cells are frequently investigated with *in vitro* and *in vivo* studies to enlighten carcinogenesis and metastasis [76]. Adipocytes and breast cancer cells are immediately near one another due to invasive breast tumors that penetrate the basement membrane and infiltrate fibrous tissue barriers, enabling paracrine interaction between two cell types [85].

To understand how adipocytes affected the motility of epithelial and cancer cells and the role of adipocytes on the cancer-related gene expressions on co-culture systems in TME, adipocyte-conditioned media treated MCF-10A mammary epithelial cells and invasive MCF-10CA1 carcinoma cells were used. The first step of the study was the differentiation of primary human breast preadipocytes into lipid-accumulating mature adipocytes. Preadipocytes were cultured in a differentiation medium for seven days to do this experiment. After differentiation, adipocytes were maintained in DMEM-F12 containing 1% (penicillin–streptomycin fungizone) PSF, 10% (fetal bovine serum) FBS, and five ng/ml insulin for a week. To compare the role of TME in normal and cancer cell motility, cells were treated with adipocyte-conditioned media. For the formation of conditioned medium (CM), preadipocytes were serum-starved in DMEM: F12 containing 1%PSF for 24 h. Meanwhile, MCF-10A and MCF-10CA1 cells were serum-starved overnight and treated with CM from preadipocytes and mature adipocytes for 24 h. It was found that mature adipocyte-conditioned media

increases cancer cell motility and migration. In addition, mature adipocyte-CM increases phosphorylated-Akt levels in MCF-10CA cells. However, angiogenesis (fibroblast growth factor receptor-2, interferon-a, and platelet-derived growth factor β-peptide) and matrix metalloproteinases (MMP-1, −2, and − 9) genes were downregulated in adipocyte CM-treated MCF-10A cells, in adipocyte-CM treated MCF-10ACA1 cancer cells, PAI-1, TGF-B-1 expression upregulated but MMP1 and angiopoietin-1 were downregulated. These data confirmed the positive effect of adipocytes on cell survival, proliferation, migration, and tumorigenesis in breast cancer (**Figure 4**) [86].

The dysregulation of adipokine synthesis and secretion is a significant risk factor for obesity, and Type 2 diabetes mellitus (T2DM) results from obesity. Given that T2DM is caused by insulin resistance, insulin promotes adipokine production, leading to cancer development [87]. Considering T2DM is related to cancer, metformin is used as an oral antidiabetic drug explaining the role of adipocytes in inflammationassociated cancer development. To find out how adipokine production is associated with obesity and T2DM, the molecular mechanism in insulin resistance and breast cancer development needs to be clarified. Similar to other studies, preadipocytes were differentiated for 14 days, and after differentiation, adipocyte cells were propagated in low glucose (5 mM) conditions for 48 h. Then, MCF-7 cells were treated with

#### **Figure 4.**

*A workflow example in adipocytes-related cancer studies.*

#### *Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies… DOI: http://dx.doi.org/10.5772/intechopen.113859*

preadipocyte and adipocyte-CM for 72 h. It was found that adipocyte-derived CM significantly induced MCF-7 cell proliferation that preadipocyte-derived CM did not. In addition, IL-6 expression was increased in differentiated mature adipocytes, which are downregulated by metformin. Findings suggested that conditioned media from adipocytes increase cell proliferation and pro-inflammatory adipokine expression, which are decreased by metformin treatment (**Figure 4**) [88].

Utilizing direct or indirect 2D co-culture, in which tumor and stromal cells are cultured together on a flat, has begun to be inadequate for explaining the complex intercellular interactions between heterogeneous cancer, immune and stromal cells, and between cells and ECM [89]. However, compared to 2D culture, 3D culture is the most commonly effective method for constructing tissue architecture lacking monolayer cultures and better recapitulating the effect of TME. The 3D cell culture enables the investigation of cell function, gene expression, and paracrine signaling effects.

Organoids are one of the improved *in vitro* tissue engineering and 3D-cell culture applications that represent the structure and function of the organ *in vivo*, such as a population of self-organizing stem cells developing into specific complex organs [90]. Organoids can be produced through cell signaling pathways regulating self-organization and differentiation of embryonic stem cells (ESCs), induced pluripotent stem cells (iPSCs), and adult stem cells (ASCs) for the establishment of complex organs. For successful organoid formation and growth, improved cell culture conditions activate cellular signaling pathways triggered by intrinsic factors and extrinsic mediators, such as extracellular matrix (ECM), to maintain stem cell function [91]. Since the cancer organoids have the same characteristics and genetic background as the original tumors, co-culture cancer organoid models have been sufficiently developed to mimic the heterogeneous environment *in vitro* to investigate the complex interaction network of cellular and non-cellular components in TME. However, maintaining different cancer organoids requires cancer-specific procedures and methods.

In recent years, many studies have been conducted on organoids for cancer research. In such a study, biopsies of metastatic lesions of colorectal cancer patients were collected, and patient-derived organoids (PDOs) were generated in cultures. These PDOs were tested to consider the sensitivity of patients to chemotherapy and observed that PDOs can be used to foresee which patients bearing metastatic colorectal cancer are insensitive to irinotecan-based chemotherapy [92]. In another study, organoid models were formed from human breast epithelial samples and used to understand the molecular mechanisms underlying breast cancer oncogenesis. Four breast cancer-associated tumor suppressor genes were knocked out by CRISPR-Cas9 in mammary progenitor cells, and observed that at least three tumor suppressor genes must be blocked for breast tumorigenesis [93]. Wang et al. carried out a study with PDOs from hepatocellular carcinoma (HCC) patients to demonstrate the relationship between the presence of CD44 surface marker and resistance to sorafenib treatment in HCC patients. They also indicated that blocking Hedgehog signaling can reverse this sorafenib resistance [94]. Another study used PDOs generated from colorectal cancer patients for genomic and transcriptomic analysis to assess multi-drug response. It is indicated that these PDOs can be used to predict the response of the patient to drug treatment and develop personalized therapies [95].

The method is frequently initiated by mechanical or enzymatic digestion of tumor tissues obtained from the minimum necrotic margins into pieces about 1 mm in diameter. Following the breaking down of tumor samples, it is required to seed the cells with supplemented media onto a 3D Matrigel scaffold matrix, which consists of ECM proteins such as laminin, entactin, proteoglycan, and collagen IV and can

be improved by tumor-specific growth factors such as Wnt3A, transforming growth factor beta (TGF-B) receptor inhibitor, and Noggin of epidermal growth factor (EGF) [96]. Organoid cells are co-cultured with TME components through this process.

## **4. Conclusion**

Identifying the complex network between stromal cells and cancer cells in the TME is a potential hallmark to enlighten cancer progression and metastasis. This chapter summarizes the role of TME in various cancers by describing current co-culture cell culture applications.

The critical ideas of TME and its possible uses in cell culture are outlined in this chapter. This chapter concentrates on the research on TME in adipocytes and cancer cells since TME is highly complicated. The components of TME and emerging cell culture applications that have a great deal of promise to represent different malignancies are outlined in this chapter. We also provide a summary of co-culture's uses in cancer research. Additionally, this chapter offers more information on cutting-edge TME investigations based on cell culture.

## **Acknowledgements**

We acknowledge İrem Nur Ateş and Deniz Turan for their valuable contribution to constructing this chapter.

## **Conflict of interest**

The authors declare no conflict of interest.

## **Author details**

Ozge Rencuzogullari, Pelin Ozfiliz-Kilbas\*, Enes Bal and Burcu Ayhan-Sahin Department of Molecular Biology and Genetics, Istanbul Kultur University, Istanbul, Turkey

\*Address all correspondence to: p.ozfiliz@iku.edu.tr

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

*DOI: http://dx.doi.org/10.5772/intechopen.113859 Perspective Chapter: Investigating Cancer Tumor Microenvironment* In Vitro *– Co-Culture Studies…*

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## **Chapter 4**

## Perspective Chapter: *In Vitro* Contracting Cardiomyogenic Models from Whole Fish Embryos and Larvae – Method, Properties, and Applications

*Bianka Grunow and Valeria Di Leonardo*

## **Abstract**

Heart diseases remain a leading cause of mortality worldwide. The development of effective treatments and interventions relies on a deep understanding of cardiac biology. Traditional two-dimensional (2D) cell cultures and animal models fall short in replicating crucial physiological and pathological features of cardiac tissue. In response, 3D cardiac models have emerged, offering a more faithful replication of the native heart tissue's architecture and functionality in a controlled environment. Although technical hurdles limit the widespread adoption of *in vitro* 3D models, they hold promise for advancing cardiovascular research. This chapter provides a description of the development of 3D spontaneously contracting cardiac primary cultures derived from fish embryos and larvae, presenting an easily accessible model for diverse applications, including the investigation of viral heart infections, as well as biomedical, pharmacological, and cardiology research. In this chapter, we will highlight the importance of *in vitro* model systems for modern cardiac research. Additionally, we will provide an overview of the protocol and results concerning the creation of *in vitro* 3D heart-like cell aggregates using enzymatically digested whole fish embryos/larvae. These aggregates exhibit long-term stability and spontaneous contractions, making them promising candidates for high-throughput screening.

**Keywords:** cardiac model, 3D culture, heart, fish virology, pharmacology

## **1. Introduction**

Cell cultures represent a fundamental *in vitro* technology in several research fields [1–3]. Since these experimental systems have become part of everyday research, they have contributed to important discoveries, including key findings in cardiomyocyte mechanisms concerning the cardiovascular field [3]. Cardiomyocytes are the main components of heart tissue and have their origin in mesodermal stem cells [4]. A lot of work in cardiovascular research has been focused on the development of cardiomyogenic models to study various conditions, e.g., heart diseases and healing capacity [5].

*In vivo* models of animal species like rats and pigs are currently used as the gold standard to understand pathology mechanisms and explore novel treatments within the context of a whole organism [3, 6], as the cardiovascular apparatus is characterized by a high degree of complexity, which means that a number of cardiovascular diseases are usually multifactorial [7, 8]. However, these kinds of experimental *in vivo* models have some important limitations that make them alone insufficient in certain circumstances such as the discovery and development of new therapies [3, 6]. The main disadvantages of *in vivo* models are in fact the ethical concerns, the high costs, and the low throughput. As an alternative, *in vitro* systems are good platforms for high-throughput drug efficiency and toxicity tests and are widely used to study biological mechanisms in a faster, low-cost, and reproducible way [6]. Taken together, *in vivo* and *in vitro* systems allowed the study of complex biological mechanisms and, during recent years, new emerging vertebrate models enriched the tools of researchers engaged in cardiovascular studies for human and veterinary medicine [9].

Since the 1990s, teleost fishes, in particular, have grown in popularity because of the introduction of zebrafish (*Danio rerio*) as a commonly used animal model in several research fields due to biological similarities with humans and generally good conservation and expression of important target genes. Specifically, zebrafish is a small freshwater cyprinid that is cheap and easy to grow and produces a high amount of external transparent embryos, making it an efficient model for multi-disciplinary studies in cardiac research, covering a range of topics including, but not limited to, development, genetic biology, and regeneration [10]. In addition to zebrafish research, in general, fish cell cultures of the mainly remunerative fish species are increasingly established due to their relevance in the important economic sector of aquaculture and fisheries, and the necessity to obtain more cost-effective model systems for research in this field as well [11].

For these reasons, multiple *in vitro* cardiomyogenic models from different mammalian and teleost species were developed for basic and applied research during the last decades [3, 9].

The technology used to produce cardiomyogenic models can vary according to the cell source and the use of 2D or 3D culture conditions. Among the variety of experimental techniques, this book chapter will focus on the cultivation methods for 3D autonomously beating cardiomyocytes (defined as "SCC—Spontaneously Contracting Cell aggregate", ZebraFish Heart AggregateZFHA, and SCPC—Salmon Cardiac Primary Cultures") using whole fish larvae from the pre- and post-hatch stage as the main cell source, including derivation outcomes from different species. In addition, the following sections explore the potential applications of 3D spontaneously beating cardiomyogenic fish cell cultures, with emphasis on basic research regarding their use to study resilience to climate change, and in applied research in pharmacology and fish virology.

#### **1.1 Overview of cardiomyogenic** *in vitro* **models**

The main approaches employed to generate *in vitro* heart models can be categorized based on the type of biological sample utilized. These include the utilization of whole isolated heart organs [12], sections of heart tissue [13], explant cultures [14], cultivation of induced pluripotent stem cells (IPS) [15], as well as the culture of embryonic stem cells, and embryoid bodies [16]. However, if these approaches are carried out using adult heart tissue from mammals, the regenerative capacity is not sufficient. In addition, obtaining biological samples from humans is challenging due to costs and ethical considerations.

#### *Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*

Furthermore, the general limited availability of differentiated cardiomyocytes in 3D models poses a significant constraint in cardiovascular research. A promising tool for research involves the use of induced pluripotent stem cells to generate fully developed cardiomyocytes. Nonetheless, this technology is still in its early stages and faces limitations in terms of differentiation efficiency, thereby reducing the model's reliability [17]. Over the past few years, a growing body of research has focused on utilizing cardiac scaffold-free spheroids for drug testing and toxicology purposes. Numerous studies have been published during this time, often employing a combination or co-culture of various cell types including rodent or human primary cardiomyocytes, human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs), fibroblasts, stem cells, and endothelial cells [17–23]. An inherent problem with previous *in vitro* cardiac test systems derived from various organisms such as mice, birds, amphibians, and fish is their limited activity. These test systems show either no contraction or only in response to electrical stimulation or only brief contractions, lasting from a few minutes to a few hours or days, which limits their use in the study of contractile activity of cardiac tissue [24]. Another disadvantage of conventional cell cultures is their lack of resemblance to organotypic structures because, in most cases, *in vitro* models are available as individual cells or as adherent cells organized in two-dimensional layers.

It is widely recognized that the physiological environment of cells is characterized by the presence of an extracellular matrix and complex interactions based on biochemical and mechanical stimuli [1, 3, 17]. Specifically, cardiomyocytes are naturally and constantly subjected to shear stress from blood flow, mechanical deformations, electrical impulses, and calcium transients, which are mostly absent in the artificial environment of classical two-dimensional (2D) cell cultures. For these reasons, there is a need to achieve experimental three-dimensional (3D) models with the ability to mimic the complex and dynamic microenvironment found in living tissues [17].

The advantages of 3D cell cultures over conventional 2D cell cultures are manifold. In 3D cultures, cells can interact with each other and their environment, allowing for the formation of cell-cell contacts, cell-matrix interactions, and the development of tissue-like structures. In particular, cells in 3D cultures can communicate more effectively through direct cell-cell contacts such as gap junctions. This enables the exchange of important molecular signals, growth factors, and cytokines, leading to improved cell differentiation, functionality, and responsiveness to external stimuli. This better mimics the conditions and cellular behavior *in vivo*, leading to more accurate and reliable experimental results.

Consequently, 3D cultures provide a valuable platform for the study of cell–cell interactions, tissue development, disease progression, and drug treatments [17, 22].

Since 2006, there has been a growing interest in utilizing fish cells for a more detailed study of cardiac physiology and human cardiac research [24–26]. Researchers have highlighted significant differences between murine and human cardiac electrophysiology, while noting the remarkable similarity between fish and human cardiac electrophysiology in terms of action potential properties and thus ion channels [24, 25]. Compared to the human models, 3D spontaneously beating cardiomyogenic cultures from fishes are not dependent on clinical samples, and several experimental analyses confirmed the presence of fully developed cardiomyocytes organized in adult tissue phenotype and owning many biological characteristics in common with human cardiomyocytes [27–34]. The establishment of *in vitro* model systems with fish cells is therefore important not only for basic research in fish biology and virology but also for research in human medicine and pharmacology.

## **2. Establishment and cultivation of 3D spontaneously beating heart aggregates from fish**

It is possible to obtain autonomously contracting 3D cardiomyogenic aggregates from both pre- and post-hatching whole fish larvae in a simple and efficient way (**Figure 1**). The main lab equipment and materials needed are in line with common primary cell culture isolation protocols and are summarized in **Tables 1** and **2**. At this point, it is significant to use larvae that are at least at eye point stage and still within the yolk sac phase so that they are not actively feeding. Larvae of several salmonid species like rainbow trout (*Oncorhynchus mykiss*), brown trout (*Salmo trutta*),

#### **Figure 1.**

*Illustration of the establishment of spontaneously contracting cell aggregates (SCCs)/ZebraFish heart aggregates (ZFHAs) and Salmon cardiac primary cultures (SCPCs).*

*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*


#### **Table 1.**

*Overview of the cell lab equipment required for the establishment of SCCs, ZFHA, and SCPCs.*


#### **Table 2.**

*Summary of the main solutions/reagents used for the establishment and cultivation of SCCs, ZFHA, and SCPCs divided per protocol step.*

maraena whitefish (*Coregonus maraena*), Atlantic salmon (*Salmo salar*), but also zebrafish (*Danio rerio*) or sturgeon species like *Acipenser oxyrinchus* were used to produce these cardiomyogenic *in vitro* systems [27–34].


## **2.1 Troubleshooting—contamination**

Contamination in primary fish cell cultures poses a significant challenge to maintaining the integrity of research experiments and ensuring the reliability of results/outcomes. Here, we outline a systematic and scientific approach to mitigating and preventing contamination, both bacterial and fungal. Before initiating cell cultures, clean and sterilize all equipment and glassware. Just work under the sterile bench and use gloves that were previously cleaned with a 70% ethanol solution. Additionally, the contamination can be avoided/prevented or reduced through careful washing steps before preparation.

Upon identification of contamination, take the following steps:


*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*


#### **2.2 Troubleshooting—absence of cardiomyogenic aggregates**

The isolation protocol described in Section 2 is generally easy to perform compared to common cell isolation practices, but good technical execution is nevertheless essential to ensure the presence of functionally active cardiomyogenic aggregates. The following are some tips to consider in order to maximize the yield of heart aggregates during the isolation process.


## **3. Characteristics**

## **3.1 General properties**

A first work in 2010 proved the possibility of obtaining SCCs from rainbow trout by applying the previously described protocol [27]. In rainbow trout, it was shown that a short time before hatching, the highest ratio of SCCs can be gained with up to five SCCs from one fish larva [29]. In zebrafish, here called zebrafish heart aggregate (ZFHA), larvae should be used 3-4 days post-fertilization (dpf) [31].

The 3D SCCs are able to mimic myocardial tissue organization after a few days in culture. After 7 days in culture, most of the SCCs are developed [29]. Analysis of SCCs, larval heart, and adult fish heart suggested that the embryonic progenitors of heart tissue are able to differentiate *in vitro* in fully developed adult-like cardiomyocytes while the heart from larval stage revealed immature and unorganized sarcomere structures within the cells, so that heart cells of larvae are not fully differentiated. These kinds of observations were verified by electron microscopic analysis for SCCs generated from salmonids [29, 32]. Additional confirmation of the theory was performed with zebrafish Tg (cmlc2:eGFP) larvae whose fluorescence indicates the expression of eGFP attached to cmcl2 in the heart [34]. This fluorescence signal was also present in the *in vitro* ZFHA indicating the further differentiation and propagation of heart progenitor cells [34].

This work, in particular, demonstrated the possibility of cultivating active cardiomyocytes from rainbow trout or other fish species in the long term over several months of constant beating frequency, **Figure 2** [29]. These important results show the possibility of obtaining highly specialized adult-like cardiac structures from whole larval cell cultures and the possibility of using fish heart aggregates as *in vitro* specification models in developmental biology.

## **3.2 Cell anatomy**

The cell anatomy of SCC, which includes various cell types of the heart, has been extensively characterized using advanced imaging techniques such as confocal microscopy and electron microscopy, providing detailed information about the ultrastructure and spatial organization of cells [27–34].

Transmission electron microscopy confirmed the presence of well-developed sarcomeres within the beating cell aggregates with a length ranging from 1.4 to 1.6 μm consistent with adult cardiac tissue structure [27, 29, 31]. Moreover, the reported presence of T-tubuli structures in the z-bands and near the sarcoplasmic reticulum indicates the importance of ion flux and action potential [29].

Moreover, immunohistochemical analysis within SCCs revealed the presence of well-organized sarcomeres and heart-specific proteins [27–31]. LC-MS mass spectrometry analysis confirmed the previously reported observations by showing that 80% of the isolated proteins from 3D SCCs/ZFHA were also expressed in the atrium

*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*

and/or ventricular tissue of adult animals of the same species [31]. Studies on SCCs generated from larvae of zebrafish, rainbow trout, and Atlantic salmon have demonstrated important aspects of cell morphology within SCCs [27–34]. In fact, it has been shown that *in vitro* heart aggregates exhibit a syncytium-like structure with a high number of mitochondria, cell contact, and myofilaments, reflecting their contractile capacity, active metabolism, and long-term stability [27–34].

Pacemaker cells are the primary contractile cells responsible for generating the long-lasting rhythmic contractions observed in SCCs. Two important proteins associated with pacemaker cell function are HCN4 (Hyperpolarization-activated cyclic nucleotidegated channel 4) and Connexin45 (Cx45). HCN4 is a key ion channel protein, which is responsible for the "funny" current (If), which plays a critical role in the generation of pacemaker potentials and the spontaneous depolarization phase of the action potential. Cx45 is a gap junction protein that facilitates electrical coupling between pacemaker cells and helps synchronize their activity. Both have been found in the SCCs [28].

Taken together, these results suggest the high reliability of SCCs in the *in vitro* reproduction of heart tissue development and organization, highlighting their potential use as 3D cardiac model systems.

#### **3.3 Cell physiology**

*In vitro* systems featuring functional ion channels serve as invaluable resources for scientific research. Ion channels, which are transmembrane proteins forming pores, facilitate the movement of electrically charged particles (ions) across cell membranes. They play a crucial role in regulating electrical signaling within cells. By employing *in vitro* systems that replicate ion channels, researchers gain the ability to investigate their functionality and regulation under carefully controlled conditions [35].

Electrophysiological techniques enable the recording and analysis of electrical signals generated by cells. Intracellular action potentials refer to the electrical impulses that occur within individual cells, while extracellular field potentials represent the summation of electrical activity from multiple nearby cells. These measurements provide valuable insights into cellular communication, ion channel function, and tissue-level electrical properties. Electrophysiological measurements of intracellular action potentials and extracellular field potentials thus provide valuable information about cellular electrophysiology. These techniques have multiple applications and contribute to our understanding of cardiac function in general, and more specifically arrhythmias, ion channelopathies, and drug effects on cardiac cells [36].

In the case of SCCs, several studies have demonstrated their electrophysiological properties. These studies have shown that SCCs can beat autonomously for up to 6 months without electrical stimulation. Electrophysiological analysis using sharp electrodes revealed that the cardiomyocyte characteristics of the SCC are very similar to human cardiomyocytes *in vivo* [28, 30]. The action potential exhibits a distinctive plateau phase and a rapid final repolarization phase. Initial tests have shown the reaction on several drugs (please see Section 4.2.1.) as well as on temperature. The contraction frequency of the SCCs correlated highly with temperature [29]. A steady and significant increase in their frequency with increasing temperature and vice versa was observed [29], as found in the *in vivo* condition in fish and mammals [37–39].

Overall, *in vitro* systems and electrophysiological measurements in fish models offer valuable insights into ion channel function, cardiac electrophysiology, and drug effects. These models contribute to our understanding of cardiac disorders and aid in the development of therapeutic interventions in human and veterinary medicine.

## **4. Applications**

In line with the described properties of Section 3, SCCs represent a potential investigative tool in numerous research fields and allow the study of different aspects of cardiac tissue, which are consistent with its complexity. The following is a nonexhaustive description of some of the main potential areas of application according to the experimentally proven properties of SCCs to date.

#### **4.1 Basic science: fish heart research**

### *4.1.1 Climate model*

*In vitro* cardiac models are valuable tools for studying the impact of climate change on cardiac health and function [40–43]. These models enable researchers to investigate the effects of various environmental factors associated with climate change on the heart in a controlled laboratory setting. This includes exploring the effects of temperature fluctuations and changes in oxygen levels on cardiac cells, allowing researchers to manipulate temperature and oxygen concentrations to simulate hypoxic or hyperoxic conditions to overall mimic different climate scenarios *in vitro*. In this way, the effects of temperature and oxygen changes on cardiomyocyte viability, contractility, electrical activity, and overall cardiac function can be assessed, allowing further investigation of the biological mechanisms underlying cardiomyocyte behavior under adverse environmental conditions [37–44]. Moreover, extensive research has demonstrated the influence of temperature on cardiac contractility across vertebrates [24, 37, 41, 42]. Therefore, the need to use *in vitro* heart model in the field of climate change is also of great importance to researchers working on fish health, aquaculture, and biodiversity conservation. Indeed, ectothermic animals experience temperature changes due to their environment that result in a significant remodeling of cardiac tissue [40, 42]. The changes in the heart usually primarily affect contractile activity and metabolism. Fish living in temperate climates are particularly susceptible to significant and long-term seasonal temperature changes [45–47]. Among these fish species, salmonids like rainbow trout can remain active between 4 and 24°C [42, 48]. Also, minnow fishes like zebrafish can experience a broad change in temperature of about 10°C considering fluctuations between winter and summer [49, 50]. For carp SCCs, an increase in culture temperature from 21 to 31°C caused a significant decrease in the contraction frequencies [51]. Due to their biology, fishes are provided with mechanisms to protect heart function from temperature changes. In the past, the comprehension of biological responses to lower temperatures has been the main target of experimental research. However, higher temperatures are becoming one of the main global concerns, making the biological responses to high temperatures another important field to investigate [42]. To date, several biological mechanisms in response to sharp temperature changes are already known and, in some cases, they are similar to human responses. For example, hypothermia leads to bradycardia and higher blood viscosity in both humans and fishes [41, 42]. However, some important differences between mammals and fishes can help to reach a deeper understanding of temperature change resilience strategies. For example, it has been shown that trout cardiac actin-myosin ATPase activity was more Ca2+ sensitive than that of rats within their respective physiological temperature [42, 52]. In addition, researchers reported that the Ca2+ quantity required by trout cardiac muscle to reach half-maximal tension was about one-tenth of that of rat cardiac tissue [42]. To conclude, fish *in vitro* heart

*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*

models, which can be generated from several fish species, could provide key findings into the cellular and molecular mechanisms underlying temperature-induced cardiac stress. A deeper understanding of these processes can contribute to improve health resilience in case of unfavorable environmental conditions in response to increasing global temperatures and extreme climate events.

### **4.2 Applied science**

#### *4.2.1 Fish cell culture in human research/pharmacology*

There is a growing interest in the use of fish cells as *in vitro* model for human cardiac research because of their physiological similarity [24–26]. The electrocardiogram (ECG) of the most widely used murine cardiac model exhibits some important electrophysiological similarities to humans, including a P wave (atrial depolarization), PQ interval (conduction of impulses from the atria through AV nodes, and the His-Purkinje system to the ventricles), and the QRS complex (ventricular depolarization). However, the action potential of mouse cardiomyocytes differs significantly from that of humans [26, 53, 54], with notable differences in cardiac ion channels [24–26, 55]. One notable drawback of murine cardiac models is the absence of an ERG channel homolog (Either-a-gogo-Related-Gene channel), which limits the modeling of repolarization disturbances and the applicability of pharmacological testing. Additionally, myosin light chains (MLC2a and MLC2v) exhibit distinct development patterns in mouse cardiomyocytes. The absence of surface marker protein SIRPA in mice leads to a 10-fold higher tolerance to drugs that may be harmful to humans [34, 56].

Despite these disadvantages, fish models, such as rainbow trout and zebrafish, have gained popularity in ion channel research, as the basic properties of many ion channels are similar in fish and humans. The ERG channel present in fish models is important in drug research, especially for studying drug-induced torsades tachycardia, a specific type of ventricular arrhythmia associated with the inhibition of the rapid delayed rectifier current (IKr) and the prolongation of the QT interval on the electrocardiogram [57].

To further advance pharmacological research, the developed *in vitro* cardiac model SCC offer significant advantages, including the potential for high-throughput screening of various factors like small molecules, nucleic acids, proteins, and lipids [34]. Zebrafish research, in particular, benefits from having a fully sequenced genome, enabling the study of cardiovascular mutants to better understand the progression of cardiovascular diseases. In addition, SCCs derived from trout as well as zebrafish, hold promise for studying ion channel disruptions and identifying suitable drugs for safety screening [28, 30, 31, 34]. SCCs can be maintained in cell culture for extended periods while retaining their functionality and exhibiting contraction rates similar to the human heart [28–30]. Moreover, the pharmacological responses of L-type calcium channels using Isoproterenol and KATP-channels using Rilmakalim have been successfully demonstrated [28].

The existence of a hERG channel in these 3D SCCs further enhances their significance. In [30], extracellular field potential recordings using a multielectrode array (MEA) showed a significant prolongation of field potential duration after administration of common hERG potassium channel blockers. The addition of terfenadine at 10 μm and dofetilide at 1 μm resulted in a 2-fold and 10-fold increase in field potential durations, respectively. This response of SCCs to these drugs highlights their suitability for *in vitro* IKr assays, as required by Guideline S7B [58].

#### *4.2.2 Fish virology*

Another application of cell cultures can be found in fish virology. Fish virology is a significant research field due to the substantial economic losses caused by fish viruses in aquaculture [59]. Outbreaks of fish diseases can have severe economic and social impacts. For instance, in Chile, outbreaks of infectious salmon anemia virus (ISAV) between 2007 and 2009 resulted in losses of approximately 2 billion US dollars and 15,000 jobs in the aquaculture industry [45, 60].

The Atlantic salmon is a species known to be vulnerable to several cardiac diseases, including cardiomyopathy syndrome (CMS) caused by piscine myocarditis virus, heart and skeletal muscle inflammation (HSMI) induced by piscine reovirus, infectious salmon anemia (ISA) caused by infectious salmon anemia virus (ISAV), and pancreas disease (PD) induced by salmonid alphavirus (also known as salmon pancreas disease virus or SPDV) [33, 61–63].

SPDV has been detected in salmonids such as Atlantic salmon (*S. salar*), rainbow trout (*O. mykiss*), and brown trout (*S. trutta*) for over three decades [61, 64], and more recently, it has also been found in some non-salmonid marine species like the dab (*Limanda limanda*) [65–67]. The relatively limited host range of SPDV compared to terrestrial alphaviruses suggests that temperature sensitivity likely plays a role in virus replication [68]. SPDV is associated with two diseases impacting the salmonid aquaculture industry [69, 70]. Pancreas disease (PD) in Atlantic salmon was first recognized in 1976 and described in 1984 [71]. A similar disease in rainbow trout, initially reported in France in 1994, is called as Sleeping Disease (SD) [72]. The mortality rates associated with these diseases vary, but even the survivors' experience significant growth reduction, leading to economic losses.

The impact of fish viruses can vary depending on the specific viral strain and the type of host cells involved. Different fish species and cell types may exhibit varying susceptibility and responses to viral infections, leading to diverse outcomes in terms of disease severity and transmission dynamics. Understanding these host-virus interactions at the cellular level is crucial for effective fish virology research and the development of targeted control measures. In this research area as well, there is an effort to transition to *in vitro* models in order to reduce the use of a high number of experimental animals [45].

The SCC *in vitro* model system has therefore already been tested for use in fish virology [32, 33, 73]. Since the main focus of research here was on Atlantic salmon, the model system was renamed SCPCs (Salmon Cardiac Primary Cultures) [32]. SCPCs have proven to be valuable tools in fish virology, enabling the study of pathogens such as Salmon Pancreas Disease Virus (SPDV) and Infectious Salmon Anemia virus (ISA virus) and their effects on cardiac cells. SPDV and ISA viruses are significant pathogens that can cause severe diseases and economic losses in salmonid aquaculture. SCPCs provide a controlled and specific environment to investigate the interactions between these viruses and cardiac cells. Researchers have utilized SCPCs to examine various aspects of viral infection, including viral replication dynamics, viral entry, cellular tropism, and the subsequent molecular and cellular responses of cardiac cells. In the case of SPDV, SCPCs have shed light on the pathogenesis of the virus in cardiac tissues, including its effects on cell viability, morphology, and function [32, 33].

Moreover, SCPCs offer the advantage of representing a more biologically relevant model compared to traditional cell lines, as they maintain the characteristics and functionality of primary cardiac cells [32]. This allows for a better understanding

*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*

of the virus-host interactions within the natural cardiac environment. Additionally, SCPCs may serve as a platform for testing potential antiviral interventions, such as antiviral compounds or vaccines, to assess their efficacy in controlling viral replication and minimizing virus-induced cardiac damage in the future [33].

In conclusion, SCPCs have emerged as valuable *in vitro* models for studying SPDV, ISA virus, and other fish viral diseases. They provide insights into the molecular and cellular events underlying viral pathogenesis in cardiac tissues, aiding in the development of effective control strategies and therapeutic interventions. The utilization of SCPCs could contribute to mitigating the impact of these viral pathogens on salmonid aquaculture and improving the management of related diseases [32, 33, 73].

## **Conclusion**

*In vitro* cardiomyogenic models using whole fish larvae and embryos provide a versatile platform for investigating cardiac biology and pathology. Their unique properties and applications make them valuable tools in cardiac research for both fish and humans. With further advances and refinements, these models hold great promise for unraveling the complexities of cardiac diseases and advancing therapeutic strategies, as well as examining adaptations in heart cell anatomy and physiology due to climate changes.

## **Acknowledgements**

We would like to thank Dr. George Franz for illustrating the cartoon.

## **Conflict of interest**

The authors declare no conflict of interest.

## **Funding**

The publication of this article was funded by the Open Access Fund of the FBN.

## **Notes/thanks/other declarations**

Special thanks go to Prof. Charli Kruse, who gave me the possibility to invent this model system. Further, I would like to thank Prof. C. Kruse and Dr. Daniel Rapoport from the University of Luebeck in Germany for their support in testing the SCCmodel system with regard to pharmacological application. Further, I would like to thank Prof. Holly Shiels from the University of Manchester, UK, for the possibility to adapt the SCCs establishment also on zebrafish larvae/embryos and establish the ZFHA. Moreover, I would like to thank Dr. Patricia Noguera from Marine Scotland in Aberdeen, who tested this model system for application in fish virology.

This model system has been patented and is registered under the following number: WO2011029584A1.

## **Acronyms and abbreviations**


## **Author details**

Bianka Grunow\* and Valeria Di Leonardo Research Institute for Farm Animal Biology, Fish Growth Physiology, Dummerstorf, Germany

\*Address all correspondence to: grunow@fbn-dummerstorf.de

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

*Perspective Chapter:* In Vitro *Contracting Cardiomyogenic Models from Whole Fish Embryos… DOI: http://dx.doi.org/10.5772/intechopen.113858*

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Section 2
