DEPARTMENT OF BIOLOGY
Master’s Degree in Molecular Biotechnology
Thesis
Optimization of the extracellular hepatic matrix reengineered for applications in regenerative medicine and
development in vitro models.
Supervisors: Candidate:
Prof.ssa Arti Ahluwalia Alessandra Bassotti
Prof. Alessandro Corti
C
ONTENTSFrequently used abbreviations ... 5
1. Abstract ... 7
2 Introduction ... 9
2.4.2.1 3D cell-cell interactions and co-culture systems .... 23
2.4.2.2 Precision cut liver slices ... 24
2.4.2.3 Cell sheet stacking ... 24
2.4.2.4 Hydrogel... 25
2.4.2.5 Decellularized 3D scaffolds ... 27
2.4.2.6 3D bioprinted liver tissues ... 29
2.4.2.7 Spheroid culture model ... 30
2.4.2.8 Spherical hydrogel microparticles... 31
2.4.2.9 Bioreactors ... 32
4 Materials and methods ...40
6 Conclusion ... 87 7 Bibliography ...90
FREQUENTLY USED ABBREVIATIONS
2D Two dimensional
3D Three dimensional
BSA Bovine Serum Albumin
CG Collagen-GAG
dECM Decellularized extracellular matrix dLM Decellularized liver matrix
DMMB Dimethyl methylene blue
ECM Extracellular matrix
EDTA Ethylenediaminetetraacetic acid
FBS Fetal Bovine Serum
FDA Food and drug administration
GAG Glycosaminoglycan
GAGs Glycosaminoglycans
HCl Hydrochloric acid
HCV Hepatitis C Virus
HEPES 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid
LDH Lactate Dehydrogenase
mM Millimolar
mTG Microbial Transglutaminase
MW Molecular weight
PBS Phosphate Buffer Saline
PEG Poly (ethylene glycol)
PEG-DA Poly (ethylene glycol) diacrylate
PLA Poly (lactic acid)
PLCL Poly (lactide-co-caprolactone) PLGA Poly (lactic-co-glycolic acid) PLLA Poly (L-lactic acid)
PS Phosphatidylserine
ROS Reactive Oxygen Species
SDS Sodium dodecyl sulfate
TGFβ Transforming Growth Factor beta TNFα Tumor Necrosis Factor alfa
A
BSTRACT The liver is the primary gland involved in the metabolism of xenobiotics, drugs and toxic compounds and metabolic waste products. It has a high regenerative capacity, due to the proliferation and differentiation of liver cells that restore the organ function. However, when the liver damage is severe and prolonged, the inflammatory response and the repair mechanisms that are activated lead to a progressive fibrotic degeneration.Fibrosis is a dynamic process that involves the progressive accumulation of extracellular matrix in an attempt to repair the damage caused by different kinds of pathological conditions.
Studies focusing on the physio-pathological mechanisms associated with liver damage, degeneration and fibrosis are of great interest, and could play a key role in the identification of the major risk factors involved in these processes. In this perspective, substantial efforts have been made to develop suitable in vitro and in vivo models mimicking liver fibrosis. Several in vitro models are based on two-dimensional cultures or co-cultures of different hepatic cell types; however, a more faithful reproduction of the in vivo conditions can be obtained with three-dimensional models (3D) of cell cultures. Indeed, the 3D models allow to exploit the interactions of cells with the extracellular matrix, as well as with other cells. In recent years, methods have been developed for 3D cultures based on the use of scaffolds of smart materials, both of natural (e.g. collagen, extracellular matrix decellularized and digested ...) and synthetic origin (e.g. polyethylene glycol,
extracellular matrix, capable of supporting cell growth and mimicking porosity, permeability and mechanical stability of the in vivo conditions.
The aim of the present Thesis was to set up a suitable model for in vitro 3D cultures by using a natural hydrogel obtained from a decellularized porcine liver extracellular matrix (ddECM) preserving the structure and the composition of the native microenvironment and to test its applicability to mimic the matrix stiffening associated to liver fibrosis. Suitable experimental conditions were optimized to obtain 3D cultures of human hepatoma HepG2 cells encapsulated into ddECM gels. Cell viability and proliferation were assessed by ATP assay, LDH assay and Trypan blue exclusion test supported by a qualitative imaging with Hoechst 33342 and propidium iodide. The matrix stiffening was induced by using the microbial transglutaminase (mTG) an enzyme that catalyzes the formation of covalent cross-links between glutamine and lysine in proteins. The effects of such treatment on micromechanical properties of hydrogels and on encapsulated cells were then evaluated. Finally, the last part of this study was focused on the possible use of the optimized ddECM in the production of small-sized, alginate-free spherical hydrogel microcapsules with a new microencapsulation device (Spherical Hydrogel Generator) recently developed at the Research Center “E. Piaggio” (Pisa).
The results obtained indicate that ddECM-derived hydrogel can be used to prepare suitable 3D in vitro models mimicking in vivo ECM composition and potentially allowing to improve the study of patho-physiological processes such as the ECM stiffening associated to the healthy-to-fibrotic transition.
INTRODUCTION
BACKGROUND
The liver is the largest internal organ of the body, accounting for 2% of the weight of an adult (∼1.5 kg), and it is responsible of several functions such as metabolism of sugar, proteins, amino acids and lipids, detoxification of exogenous chemicals, production of bile acids and storage of various essential compounds, such as vitamins and iron. Liver failure has potentially fatal consequences, representing the twelfth most frequent cause of death in the US (Kasuya J. and Tanishita K., 2012) with over 1 million deaths per year worldwide (Gonzalez F. and Keeffe T.,2011).
This organ is known for its regenerative capacity, in fact upon an acute injury it may restore its original structure and mass even when large parts are destroyed or removed.
A long-term damage of liver – as in the case of chronic liver diseases - is accompanied by an impairment of its regenerative capacity, with extensive areas of necrosis and apoptosis of hepatocytes. Chronic damage induces severe inflammatory and reparative processes which in turn may lead to the replacement of large part of liver parenchyma with a scar tissue, a process known as fibrosis. Fibrosis
can lead to portal hypertension (the scar tissue distorts blood flow through the liver) or evolve to cirrhosis in which the scarring results in disruption of normal hepatic architecture and liver dysfunction. Despite their high incidence and destructive outcomes, the only
transplantation, which have the disadvantage to be restricted by the number of compatible donors (Banares R. et al., 2013). On the other hand, the poor availability of therapeutic drugs could be due to the lack of robust and representative in vitro models of human liver
fibrosis (Jung L. et al., 2013).
To date, in vivo models are still the most popular preclinical assessment model, even though the predictive value for the human physiological response in terms of both efficacy and toxicity is sometimes poor (Yanguas S.C. et al., 2016). Furthermore, ethical concerns stimulate the replacement of the animal models according to the 3R principle – “Reduction, Refinement and Replacement” (Van Grunsven L. A., 2017).
As claimed by Forbes et al., (2015) cell-based therapies, engineered tissue transplantation or bio-artificial liver system are some of the methods that should be developed since they result more predictive and efficient than in vivo models. In particular, liver tissue engineering is considered a potentially valuable new therapeutic technique for liver diseases.
GENERAL LIVER STRUCTURE AND IN VIVO MICROENVIRONMENT
The liver is part of the hepatobiliary system and consists of parenchymal cells, named hepatocytes, organized in cord-like structures by non-parenchymal cells (NPCs).
Hepatocytes and NPCs form a hexagonal unit know as lobule which is the functional unit of the liver (Van Grunsven L. A., 2017).
Figure 2.1Schematic representation of the hexagonal-shaped liver lobules, with the central vein and portal triad. Amplification of the lobule segment, showing the stratified parenchyma and the countercurrent flows of the bile and blood circulation. Adapted from Treyer and Müsch (2013).
Hepatocytes are responsible of most of the hepatic functions such as the synthesis of glutamine, bile acid, cholesterol, lipids, urea, albumin and blood clotting factors; furthermore, these cells carry out the metabolism of xenobiotic by cytochrome P450 enzymes (Huang B. et al.,2006). These cells present a unique cytoarchitecture closely linked to their specific functions with a cuboidal profile and a particular stacked disposition along the liver lobule that result in an exclusive polarization. These plates of hepatocytes define the basic functional unit of the liver, known as the acinus.
Although hepatocytes are the most abundant cell population of the liver, it is now widely accepted that the NPCs play an important role in supporting hepatocyte functionality and proliferation (Monga et al.,2010). NPCs consist of five major non-parenchymal cell types: liver sinusoidal endothelial cell (LSEC), Kupffer cells, stellate cells, biliary epithelial cells, and natural killer cells that will be described below.
Liver sinusoidal endothelial cell (LSECs) are characterized by fenestration and the lack of basement membrane. This kind of cell represent more than a barrier between blood and hepatocytes, they actively participate in hepatic clearance, in fact LSECs act as scavengers thanks to the high receptor-mediated endocytic activity, which remove physiological or foreign macromolecules from the blood (Braet F. and Wisse E., 2002).
In addition, LSECs are implicated in inflammatory reaction mechanisms including detection of pathogen associated molecular patterns (PAMPs), secretion of cytokines and chemokines, and involvement in adhesion and diapedesis of leukocytes (Braeuning A. et al., 2013).
Kupffer cells are the resident liver macrophages, with high endocytic and phagocytic activity. These cells are able to release specific cytokines that are mediators of the local and systemic inflammation (Dixon L. et al., 2013). The immune response is further mediated by the natural killer cells (NKs), which are intrahepatic leucocytes.
Hepatic stellate cells (HSCs) are perisinusoidal cells located between the endothelium and the liver parenchyma, in the space of Disse. These cells are responsible for the storage of vitamin A, control the production and homeostasis of extra-cellular matrix (ECM), regulate contractility of the sinusoids and secrete cytokines. HSCs can also acquire a myofibroblast phenotype, which impairs ECM regulation and leads to fibrosis (Puche J. et al., 2013). This transition is regulated by the interaction of HSCs with several cell
types that led to the activation of specific pathways involved in wound healing reaction (Trautwein C, Friedman SL et al.,2015.)
Cholangiocytes or biliary epithelial cells(BECs) connect the biliary tract in the liver and are involved in the secretion of bile via the canalicular network as well as mucin secretion and reparative function in disease conditions.
The ability of these different cells types to cooperatively interact with each other demonstrates the complexity of the liver’s structure and functionality. Therefore, the study and development of in vitro model that mimic, as closely as possible, the in vivo environment is extremely difficult and requires the ability to overcome a multitude of obstacles, as for instance, the alterations of the Extracellular Matrix (ECM) that led to severe pathologies such as fibrosis or cirrhosis.
The ECM composition, topography and biomechanical properties change completely during the onset of inflammation. It is mainly located in the portal tracts, central veins and in the sinusoidal walls of the space of Disse and it is well known that her protein like type I collagen and fibronectin (in the liver parenchyma) and collagen type III, IV and laminin (in the portal and central regions) (Godoy P. et al.,2013), activate intracellular signaling cascades and gene regulation in hepatocytes that led to the modification of the phenotype by means of the bond to their cell-surface receptors (e.g. integrins).
LIVER FIBROSIS AND CELLS INVOLVED
In general, the etiopathogenic mechanism of liver fibrosis can be considered as a chronic wound healing process characterized by an increased deposition of connective tissue caused by a chronic liver injury associated with infections, metabolic disorders, alcohol abuse or autoimmune diseases (Friedman SL., 2003). The cell types mainly involved in this process are the HSCs. Indeed, HSCs represent in normal condition approximately 5-8% of the total cell population, but upon liver injury, and especially during chronic liver damage, a dynamic and programmed modification of HSCs phenotype occurs.
In this paragraph the cell populations and the signaling factors involved in liver fibrosis will be described.
Figure 2.2 Phenotypic features of hepatic stellate cell activation during liver injury and resolution. Adapted from Friedman L (2000).
The activation of HSCs can be separated in two distinct stages, i.e. initiation and perpetuation phase (Forbes S. et al,.2004).
Activation of HSCs is caused by reactive oxygen species (ROS), inflammatory cytokines and growth factors that originates from injured hepatocytes, sinusoidal endothelial cells and Kupffer cells. This stimulus induces morphological changes in HSCs shape, the loss of vitamin A content and the expression of cell surface receptors for growth factors and cytokines (Trautwein C. et al.,2015).
Hepatocytes are the main source of lipid peroxides and apoptotic bodies in injured liver, that are absorbed by HSCs. This induces the expression of type I collagen and the increase of ROS production causing intracellular signaling cascades that determine the synthesis of monocyte chemo-attractant protein-1 (MCP-1) and transforming growth factor-beta (TGFβ).
According to what suggested by Trautwein (2015), other actors that facilitates HSC initiation are endothelial cells with secretion of fibronectin and Kupffer cells with release of TGFβ and ROS. Autocrine and paracrine signals contribute to transient HSCs activation that involve initial inflammatory reaction and collagen deposition in the liver (Crespo S. et al.,2016).
During the subsequent perpetuation phase, HSCs acquire a myofibroblastic phenotype and become more proliferative and contractile, leading to enhanced production of ECM proteins, angiogenesis regulation and to the amplification of the immune response. The proliferative stage associated with the activation of HSCs is governed by PDGF but there are some evidences that
include also TGFα and the epidermal growth factor (Svegliati Baroni G. et al.,2005).
The extent of fibrosis and inflammation depends on the type and duration of the liver injury. When the injury is acute, the regeneration process is sufficient to replacement the necrotic and apoptotic cells, removing scar tissue, and resolving the inflammation. However, when injury is chronic, this procedure is insufficient and liver tissue is gradually replaced by extracellular matrix. As this process progresses, morbidity and mortality due to disease-related complications will increase unless the causal factor is removed.
IN VIVO AND IN VITRO MODELS OF LIVER FIBROSIS
2.4.1 ANIMAL MODELS OF LIVER FIBROSIS
Although intense research during the last twenty years has led to considerable improvements in the knowledge of liver fibrosis pathogenesis, effective antifibrotic therapies are still lacking. In this perspective, a better understanding of the mechanisms implicated in the initiation, progression and resolution of fibrosis is required. Animal models were demonstrated to be essential to improve our knowledge about all the processes involved in fibrogenesis and to identify potential therapeutic targets of antifibrotic therapies (Delire B. et al.,2015). Indeed, their use offers several advantages such as (i) the possibility to collect multiple samples at different time-points and to perform sequential studies, (ii) a shorter time for disease development, (iii) the ability to control and reduce variables that cannot be closely followed in humans and (iv) the ability to study
the implication of specific genes/signaling pathways by the use of genetically modified animals (Starkel P. and Leclercq I.A., 2011.) Moreover, as compared to in vitro systems, animal models allow to study both the intra-organ interactions (i.e. cell-cell and cell-matrix) - and the complex crosstalk of the liver with the entire body, including immune, vascular, metabolic and endocrine systems. Unluckily, animal models cannot help to explain all the mechanisms involved in human diseases (including liver fibrosis) since large differences exist between humans and animals as regard pathogenicity, timing and immuno-inflammatory reactions induced by pathogens (Delire B. et al.,2015). However, several in vivo animal models were developed and specifically designed to try to mimic the different conditions involved in the onset of human liver fibrosis. Among these models, the four most commonly used are briefly discussed here.
1. Chemical-based animal models. Chemical-based animal models are widely used because of their high reproducibility, ease of use and appropriate reflection of the mechanisms involved in human liver fibrosis (Crespo S. et al.,2016). Several toxic compounds are commonly used to induce hepatotoxicity, including carbon tetrachloride (CCl4), dimethylnitrosamine (DMN), thioacetamide (TAA) and ethanol (Ensminger W. et al.,2004). These agents were administered by intraperitoneal injection to produce liver fibrosis on a relatively short-term basis. On the contrary, when administered orally or via inhalation, fibrosis damage is limited and takes more time to develop.
2. Surgery-based model. This technique is based on the occlusion of the bile duct (bile duct ligation, BDL) in order to mimic a hepatic fibrosis induced by cholestasis.
This type of obstruction induces an increase in biliary pressure which is associated with an augmented secretion of pro-inflammatory cytokines by biliary epithelial cells and is followed by the onset of a mild inflammatory response.
In general, early mortality in rodents may happen after BDL due to bile leakage, rupture of biliary cysts or gall bladder.
The mortality rate after BDL in rats is about 20% and the peaks are reached in mice after ten days. BDL can be particularly used for short-term studies of liver fibrosis associated with cholestatic injury (Delire B. et al.,2015).
3. Infection-based model. Another animal model is based on the infection with some species of Schistosoma, a parasite of humans responsible for schistosomiasis – which is able to induce – among the other effects - liver fibrosis.
Schistosoma mansoni infection is voluntarily established in mice for the high likeness to human infection and the high reproducibility. Nevertheless, different mouse strains can show great variations in hepatic fibrosis levels. Alternatively, animals can be infected by percutaneous administration of thirty-five cercarias through the tail or by intravenous administration of 10,000 viable eggs (Cheever et al.,2002). This represent the main cause of the development of granulomas associated with liver fibrosis. Collectively, the role of the inflammatory cytokines in these infection models contributes to
the activation of the HSCs and thus to the progression of liver fibrosis.
4. Diet-based model. This model is based on the administration of a methionine or methionine/choline deficient diet in experimental animals in order to induce a condition that is pathologically similar to the human metabolic steatohepatitis or fatty liver disease. A number of specific diets can be used to induce progression of Non-Alcoholic Fatty Liver Disease (NAFLD) to Non-Non-Alcoholic Steatohepatitis (NASH) in experimental animals (Anstee and Goldin 2006). Nevertheless, these diet-based models fail to mimic the typical characteristics of the human pathology, thus restricting interspecies extrapolation of results (Anstee and Goldin 2006).
5. Other models. Different transgenic animal models - in which the expression of specific genes involved in fibrogenesis was up- or down-regulated - were also developed. Indeed, genetically modified animals have become powerful research models in the past decades. In particular, they allow to gain insight into the involvement of specific proteins and signaling pathways in the generation of liver fibrosis, thus facilitating the identification of potential targets for new drugs. Nevertheless, genetic models rarely develop liver fibrosis and frequently need a second stimulus for disease induction. This indicates interaction between the environment and the genotype to manifest the disease
In conclusion animal models can be suitable for studying some features associated to the development of liver fibrosis and to test
differences constitute their major limitation and must be taken into account when animal models are used.
2.4.2 IN VITRO MODELS OF LIVER FIBROSIS
The development of suitable in vitro models mimicking the functionality of in vivo systems is highly important within several scientific fields. To date, pharmaceutical companies use in vitro models in the process of drug development to achieve preliminary results and to evaluate whether to move to in vivo studies (Ware BR. et al.,2017). The main advantage of these models is the possibility to employ human cells or tissues that may produce more relevant results for the comprehension of the mechanisms involved in human diseases.
Conventional 2D cell culture is a good - but not optimal - surrogate to study liver fibrosis. These models should at least, ensure the presence of a 3D structure and the sufficient physiological and pathophysiological expression of a variety of ECM components. Indeed, there are increasing evidences of the different effect of 2D and 3D ECM on key biological features of fibroblasts and myofibroblasts, including proliferation, migration, contraction, matrix deposition and degradation. (Trautwein C. et al., 2015.)
For in vitro studies of liver fibrosis, the most commonly hepatic cell types used, can be divided into three main categories, i.e. primary cultures, cell lines and hepatocyte-like cells derived from pluripotent stem cells. One of the most largely used cell line is represented by Hep G2 a perpetual cell line derived from the liver
tissue of a 15-year-old Caucasian American male with a well-differentiated hepatocellular carcinoma. These cells are a suitable in vitro model system for general cellular and biochemical investigations. They secrete a variety of major plasma proteins, e.g. albumin, transferrin, plasminogen, alpha 2-macroglobulin (Théard D. et al., 2007).
HepG2 cells display a controllable formation of apical and basolateral cell surface domains that resemble the bile canalicular (BC) and sinusoidal domains, respectively, in vivo.
Because of their high degree of morphological and functional differentiation in vitro, HepG2 cells are a suitable model also employed in trials with bio-artificial liver devices for the study of human liver diseases (Fanelli A., 2016).
Limitations of in vitro culture systems are related to a fast decline in hepatocyte-specific functions, de-differentiation and loss of hepatocyte polarity after short term culture.
Three dimensional models could be a solution, since it is possible with this technique to try to reproduce the conditions present within a tissue, e.g. by improving the structural cells support and providing a flexible physical environment to allow remodeling in response to tissue dynamic processes such as wound healing. The only 3D model that would reflect the most part of characteristics of the hepatic ECM is the liver ECM itself. The liver ECM is known to provide bioactive signals for the cells and acts as a reservoir for growth factors that locally enhance cellular functions (Mazza G. et al., 2017). It is becoming clear that recreate the liver microenvironment with its cell-matrix interactions, cell-cell adhesion and cellular
cell culture systems are valid to investigate cells behavior and for drug screening but have limitations in maintaining specific cell properties as observed in a 3D microenvironment.
In recent years, rising developments within the field of biomaterial and tissue engineering have simultaneously created and improved various kind forms of 3D in vitro models that can be utilized to better study cellular systems (Van Grunsven LA. et al 2017). Figure 2.3 shows some 3D models that reflect the many aspects of the hepatic ECM.
Figure 2.3Examples of 3D models reflecting the many aspects of the hepatic ECM (Van Grunsven LA. et al 2017).
2.4.2.1 3D cell-cell interactions and co-culture systems
The lack of a physiologic cellular context when studying cell activation in vitro can be addressed by co-culturing different liver cell types (Bhatia SN. et al.,1999)
Experiments with hepatocytes have demonstrated that 3D co-cultures with HSCs favors several hepatocyte functions such as engraftment, proliferation and differentiation, due to the secretion of soluble factors produced by HSCs. These mixed cultures typically maintain functionality over extended periods of time (Loreal O. et al.,2012).
Recently, it has been observed in co-culture systems of hepatocytes and HSC cell lines that the cell-to-cell proximity is of high importance to initiate the fibrotic process induced by fatty acids. By contrast, co-cultures based on primary HSCs and Kupffer cells reflect the role of immune cells in the regulation of fibrotic responses, while co-cultures consisting of HSCs and endothelial cells have shown the importance of HSCs in angiogenesis (Giraudi PJ. et al.,2014).
Anyway, current co-culture models cannot accurately mimic the in vivo liver cellular and structure composition. Although cell culture models contributed significantly to the understanding of stellate cell biology, they still fall far from recapitulating intercellular and cell-extracellular matrix interactions in the process of hepatic stellate cell activation and fibrogenesis in vivo (Mazza G. andAl-Akkad W., 2017).
2.4.2.2 Precision cut liver slices
A potential 3D model for the study of cell activation and liver fibrosis is based on the Precision-Cut Liver Slices (PCLS) technique. Unlike current in vitro models, PCLS could enable the study of these processes in vitro in a multicellular system in which cell and cell-extracellular matrix interactions are maintained.
PCLS are liver explants with a normal thickness ranging from 100 to 250 μm and a diameter of 5 mm, which allows oxygen and nutrients diffusion. PCLS can be incubated in cell culture plates, which in turn may be incorporated in dynamic organ culture systems (Fisher A. and Vickers R., 2013).
A major drawback of PCLS is that they show limited cell viability and their use has to be restricted to a period of time ranging between 24 hours and 1 week (Mazza G. and Al-Akkad W., 2017).
PCLS have been obtained from various species (e.g. rat, mouse and human) and have been extensively used for drug metabolism and toxicity studies. However, the use of PCLS in the study of hepatic stellate cell activation and liver fibrosis is still limited (Van de Bovenkamp M and Groothuis GM., 2007).
2.4.2.3 Cell sheet stacking
Cell sheet engineering is a unique scaffold-free tissue-engineering approach that uses poly(N-isopropylacrylamide) (PNIPAAm)-grafted, temperature-responsive culture dishes (TRCDs). The temperature-responsive culture surface can be
into a standard tissue culture polystyrene dish (Mazza G. and Al-Akkad W., 2017).
This material becomes hydrophilic at room temperature (20°C) and cells cannot attach; when heated at 37°C the shell becomes hydrophobic and cells can attach and proliferate on the surface. Interestingly, cells expanded on the surface can be collected without enzymatic treatment, i.e. by simply reducing the temperature of the culture plate. Hepatic cell sheets have been produced by using both parenchymal and non-parenchymal cells and this system improved hepatic functions such as albumin and urea synthesis as compared to 2D monolayer cultures. Nevertheless, the absence of vascularization - and therefore oxygen deficiency - limits the use of this system to short period of time.
2.4.2.4 Hydrogel
The tissue engineering evolution and the continuous biomaterials development have resulted in a wide range of techniques and materials with different physical and biological properties optimized for the study of distinct cell types (Geckil H.et al.,2010). Hydrogels are defined as highly hydrated polymer materials (>30% water by weight), which maintain structural integrity by physical and chemical crosslinks between polymer chains that partially resemble the physical characteristics of native ECM (Figure 2.4) (Saldin L. et al.,2017).
Figure 2.4Effect of water on polymer materials (Saldin LT. et al.,2017).
The most common artificial matrices employed for engineering biological tissue are synthetic polymers, such as polylactide-co-glycolide (PLG), polyethylene glycol (PEG), polycaprolactone (PLA), and naturally-derived hydrogels like alginates, celluloses and polyethylene (Frenguelli L., 2016).
The use of biomaterials for hepatocytes culture has gained relevance in the past years due to their capacity to stimulate specific cellular functions, direct cell-cell interactions and support the 3D architecture (Kim K. et al.,2015).
The major advantage of synthetic biomaterials is the easiest modulation of physio-chemical and biological properties as compared to natural-derived biomaterials. Nevertheless, synthetic biomaterials are more prone to poor biocompatibility and show degradability for tissue engineering and cell-based therapies than natural derived biomaterials.
For these reasons 3D scaffolds can be developed from biological ECM-derived material, the so-called “biomatrix”. The liver matrix
is a perfect example of a natural type of biomaterial. It’s consist of the native ECM recovered from decellularized livers, which can be used as a scaffold and re-populated. Obviously, the liver matrix is depleted of cellular components but ideally the original ECM composition remains intact (Uygun K. and Yarmush T., 2013).
2.4.2.5 Decellularized 3D scaffolds
Over the past few decades, several studies have demonstrated the suitability of using decellularized human or animal tissues by employing naturally occurring ECM scaffolds for tissue engineering (Baptista P.M. et al.,2011). Organ liver engineering is based on three fundamental concepts: (I) native liver ECM represents an ideal and required substrate for liver study; (II) three-dimensional acellular liver scaffolds should retain the three-dimensional macro-microstructure of liver; (III) liver regeneration can be promoted when the re-seeded three-dimensional acellular liver grafts are placed in the appropriate three-dimensional microenvironment (Mazza G. et al.,2017).
Liver ECM is the ideal microenvironment in which hepatocytes can maintain their phenotype and functionality.
To reproduce the human pathophysiological 3D environment and high ECM protein compositions, decellularization protocols need to be tissue-specific, disease-specific and species-specific (Mazza G. et al.,2017).
As summarized in Figure 2.5, several approaches were described to obtain a decellularized ECM.
Figure 2.5. Methods of tissue decellularization (Uygun et al.,2010).
The first method illustrated was based on the whole organ perfusion, with detergents through its blood vessels. In addition to perfusion, it’s possible to expose the whole organ to a combination of pressure gradient, immersion and agitation (Uygun et al.,2010). Other methods described in literature include washing and saponification of liver to remove blood, debris and to hydrolyze plasma membranes (Mattei G. et al.,2014)
The first study focused on liver decellularization was performed on rat liver, with rat hepatocytes that were maintained viable and with high expression levels of albumin, urea and CYP450 genes till 10 days after re-cellularization (Uygun et al.,2010). As reported by Faulk and colleagues (Faulk DM. et al.,2015), substrates derived from porcine, bovine and human liver were compared to study
hepatocytes survival, polarity and their major liver-specific function in vitro. These studies showed that in a porcine liver ECM, the structural and functional proteins scaffold provide an optimum physical substratum for the attachmentand spatial organization of cells. Furthermore, other studies have demonstrated the preservation of bioactive factors, such as bFGF, PDGF and VEGF, that improve the cell viability in the decellularized tissue (Bhatia S. Balis et al., 2014).
2.4.2.6 3D bioprinted liver tissues
In addition to the decellularized 3D scaffolds, biofabrication technologies, such as bioprinting, have emerged as tissue engineering approaches for building organs as well as smaller organoids or tissue construct (Murphy S.V. et al.,2014).
Technically, three main types of printing are commonly used: laser guided, droplet based and extrusion bioprinting. Each technique has its advantages and disadvantages with respect to cell handling, cell printing accuracy and structural possibilities. The laser-based methods are highly accurate, but offer low throughput, are averse to cell survival and not yet commercially available. Instead, the droplet-based and extrusion-based systems are commercially available, can reach a higher throughput and a higher cell viability, but are not very accurate (Van Grunsven L.A. et al.,2017).
This relatively new ground of tissue engineering has mainly focused on biomaterial development, different printing methods and short-term performance of mostly hepatocyte cell lines such as HepG2.
fixed and cultured up to 8 weeks in gelatin/chitosan hydrogels using an extrusion based bioprinting system.
Although these techniques are in improvement, to date, they remain too expensive for the complexity of the equipment required and for the quantity of cells needed. In addition, these models suffer in terms of reproducibility and are not suitable for high throughput applications.
2.4.2.7 Spheroid culture model
Spheroid cultures, unlike bioprinting, require fewer cells, represent an inexpensive model, are much less complex in comparison to the scaffold-based models and, finally, they can be used for high throughput applications (Gunness P. et al.,2013)
One of the first liver spheroid models developed and studied was obtained by Landry and colleagues (Landry et al.,1985). They isolated and seeded rat liver cells onto a non-adherent plastic substratum which stimulated cells to form aggregates in 1-2 days. The use of spheroids has really improved in vitro cell culture technique both preserving its simplicity and increasing its applicability. The general spheroid model concept is to seed single cells into a well or a vessel and through gravity to allow the cells to sediment to the bottom where they can aggregate to form a spheroid.
Figure 2.6A Scanning Electron Microscope (SEM) Image of a 3D Spheroid on a Nano culture plate.(from JSR Life Science)
This model improves cellular communication - as compared to 2D in vitro models - thus increasing cell viability and supporting the maintenance of a specific phenotype. Cells in spheroids may thus maintain their functionality for longer time, as compared to 2D cultures: indeed, it was observed that primary human hepatocytes (PHH) spheroids cultures remain phenotypically stable for at least 5 weeks (Van Grunsven L.A. et al.,2017).
Spheroid culture model is thus simple to obtain and more cost-efficient, but does not allow the formation of complex structures
2.4.2.8 Spherical hydrogel microparticles
As described above, the 3D architecture of hepatic cells spheroids provides maximization of cell-cell interactions maintaining the in vivo structural arrangement. On the other hand, within the 3D spheroids structure there is a nutrient and oxygen gradient formation that could cause cell suffering. On the contrary, in
which provides a 3D microenvironment with characteristic dimensions small enough to ensure an adequate supply of oxygen and other nutrients in the media as well waste removal.
Microencapsulation confers protection against stress and prevents the constant washout of extracellular matrix and soluble factors (Haghgooie R. et al.,2010).
From this perspective, microencapsulation is an emerging method that allows the use of a variety of soft biomaterials or biomaterial composites to confine cells in—usually spherical and preferably micron sized-gel constructs (Tirella A. et al.,2014). Classically, microencapsulation uses alginate, a well-known natural co-polymer derived from seaweed.
In addition to hydrogels, (ECM)-based microparticles have received much attention as that reconstruct a suitable microenvironment for encapsulated cells.
Various methods for the ECM-based microparticles preparation have been proposed, including microfluidic devices, electro spraying techniques, membrane emulsification techniques and liquid droplet formation.Using these techniques, a large amount of homogeneous ECM microparticles could be quickly prepared (Yoshida S. et al.,2017).
2.4.2.9 Bioreactors
In recent years, to support biological processes for multiple applications - such as production of biopharmaceuticals or tissue engineering - Bioreactors (BRs) were developed (Singh H. et al.,2008). The key feature of these systems is the high level of
bioprocess control, which is achieved by on-line monitorization and automated environmental culture parameters regulation, such as temperature, pH, partial pressure of oxygen (pO2), nutrient and
metabolite concentration.
In the past few years the use of bioreactors has become very popular since it represents a dynamic cell culture system that allows efficient metabolite exchange and may also provide an indirect physical stimulus through percolative or interstitial-like flow (Van Grunsven L. A. 2017).
In conclusion, great progresses have been made in the field of in vitro liver cultures and 3D liver cultures. The future will tell whether bioprinted liver tissue, spheroids, decellularized livers and PCLS cultures will replace single-layer primary cultures and in vivo rodent models such as the most widely used liver fibrosis.
Clearly, the future that is emerging is an interdisciplinary effort, that require integration of molecular cell biology, immunology, biophysics, biomaterials, device design, and pharmacokinetic modeling for a clinical translational medicine.
MECHANICAL FORCES AND FIBROSIS
As described by Wells (Wells RG., 2013), mechanical forces are essential to the development, progression and (potentially) regression of tissue fibrosis. Although often ignored in studies and models of fibrosis, mechanical signals are similar to chemical signals in their range of effects and are likely to be equally important. Understanding the role of mechanics in fibrosis is one key to better
diseases and for developing new therapies (Wells RG., 2013). Each tissue has an optimum stiffness level, and this can change in response to biochemical and physical stimuli, e.g. during the development or in pathological conditions (Saneyasu T. et al.,2016). Multiple studies have shown that liver become stiffer in fibrosis, with elastic modulus values ranging from approximately less 400-600 Pa for normal tissue to approximately 2-22 kPa for fibrotic tissue (Georges PC. et al.,2007). To test the hypothesis that matrix stiffness might affect cell activation, different studies focused on covalently crosslinked scaffolds in order to modify their stability and their mechanical properties. Three different approaches are commonly used to obtain crosslinked scaffolds, i.e. physical methods, chemical reagents or enzymes.
2.5.1 CROSSLINKING METHODS
In the last few years, mild-crosslinking methods have been successfully developed to form artificial matrices. In physically crosslinked gels, e.g. by UV light, interactions between polymers chains in amphiphilic block and graft copolymers are established by ionic and/or hydrophobic interactions or crystallization (Teixeira LS. et al.,2012).
In chemically crosslinked gels, covalent bonds are formed between polymer chains and gels can be generated by using a radical-induced polymerization, a chemical reaction, a high energy radiation exposure or an enzymatic reaction.
The physical crosslinking has the advantages of reversibility and absence of chemical (potentially harmful to the cells) reactions, but
its stability in vivo might be severely affected by interactions with bodily functions, both physiological and mechanical. In contrast, chemical crosslinking allows the formation of gels with controllable mechanical strength and superior physiological stability (Davis N. et al.,2010).
More recently enzymatically crosslinked hydrogels have sparked great interest due to the mildness of the associated reactions. The enzymes and the methods used are briefly described below.
Tyrosinases, also known as phenoloxidase and monophenol monooxygenase, catalyzes macromolecular network formation in the absence of co-factors. Tyrosinase is a copper-containing enzyme that catalyzes the oxidation of phenols, such as in tyrosine residues and dopamine, into activated quinones, in the presence of O2 (Chen
T. et al.,2003). The formation of gelatin or chitosan gel takes place in few minutes, but they can be formed only in the presence of chitosan and are unstable and mechanically weak. More than for in vitro studies, these gels can be used as glue to accelerate healing of wounds (Wu L. et al.,2011).
Lysyl oxidase is a key component in the formation and repair of the native extracellular matrix. This ubiquitous enzyme oxidizes primary amines of lysines to aldehydes. These covalent bonds that are formed stabilized the fibers that form collagen and elastin. Consequently, this enzyme is involved in the repair of many connective tissue including skeleton, cardiovascular and respiratory tract (Kothapalli C. et al.,2009). The enzyme can be used as a cross
matrix. Hydrogels become more robust thanks to continuous lysyl oxidase activity. This enzyme is abundantly present in serum, therefore the crosslinking of polymers containing lysine may occur spontaneously without the addition of an exogenous source of the enzyme (Toledano S. et al.,2009). Lysyl oxidase can be used to increase the cells extracellular matrix production incorporated in the hydrogel. In addition, it can also improve the intrinsic mechanical properties of engineered tissue by the formation of covalent bonds between the rich lysine polymers of the hydrogel and the primary amines of proteins in native tissues (Bakota E. et al.,2009).
Phosphopantetheinyl transferase is a small enzyme that presents a general mechanism to form synthetic hydrogel. The general mechanism of transferase catalysis to form synthetic hydrogels occurs by transfer of a phosphopantethein prostetic group of coenzyme A-functionalized PEG macromers to a serine residue of engineered carrier proteins. The hybrid hydrogels are formed by mixing the precursors of 8-arm-PEG-coenzymeA, at 37°C, neutral pH in the presence of Mg2+in about 15 minutes. This type of reaction is very attractive for cell biology and tissue engineering studies but is yet to be explored (Mosiewicz K. et al.,2010).
Phosphatase, thermolysin, β-lactamase and phosphatase/kinase are enzyme that can change the amphiphilicity of small peptide derivatives, e.g. by phosphorylation mediated by a kinase or dephosphorylation mediated by a phosphatase. This modification may then activate the self-assembly and non-covalent interaction of
the amphiphilic peptides in nanofiber, with consequent hydrogels formation (Yang Z. et al.,2008). The phosphatase catalyzes the removal of phosphate groups from a substrate which becomes hydrophobic. In an aqueous environment, these substrates can self-assemble to form a 3D nanofiber networks, through non-covalent interactions, which allows the gel formation. The thermolysin catalyze the formation of bond between peptides through reverse hydrolysis. This enzyme therefore reduces the solubility of one of the peptides that can self-assemble into a hydrogel via hydrophobic interactions. The β-lactamase, produced in some bacteria, catalyzes the breakage of rings and formation of a linear molecule of 4 carbon atoms, present in the structure of lactams. When these molecules are linearized, they can self-assemble to form hydrogels (Schnepp Z. et al.,2006)
Peroxidases are a wide family of enzymes that typically catalyze the decomposition of hydrogen peroxide and organic peroxides according to the following reaction: ROOR’+2e- -> ROH+R’OH. Most of the peroxidases use hydrogen peroxide as substrate. This family is commonly used for its low cytotoxicity in vivo and for the rate of the reaction catalyzed (Lee F. et al.,2009).
Transglutaminases constitute a broad enzymes family that catalyze post-translational protein modifications by inducing the formation of isopeptide bonds, the covalent conjugation of polyamines, the lipids esterification and the glutamine deamination. They are an alternative to chemical cross-linking, as they catalyze
protein, mainly a lysine, and γ -carboxamide group of a glutamine of another protein or peptide. Once formed, these bonds are highly resistant to proteolytic degradation. The gels that are formed through these cross links can be used to include cells, since they are highly biocompatible, and they have excellent transport properties that facilitate studies of drug administration (Jin R. et al.,2011). Recently, the microbial transglutaminase (mTG), derived from a variant of Streptoverticillium mobaraense has been found to be very useful for the formation of cross-links in hydrogels. The mTG is a calcium independent enzyme that catalyzes the formation of covalent cross-links between glutamine and lysine in proteins. In addition to the preparation of hydrogels, this enzyme has many other application, e.g. in the food industry. The cross-link formation not only depends on the presence of free residue of glutamine and lysine, but also on the tertiary structure of proteins (Teixeira L.S. et al.,2012).
Figure 2.8 Transglutaminase reaction, formation of covalent bonds between a free amino group of a lysine of a protein and γ -carboxamide group of a glutamine of another protein or peptide. (Teixeira LS. et al.,2012).
AIM OF THE THESIS
The aim of the present Thesis was to set up a suitable model for in vitro 3D cultures by using a natural hydrogel obtained from a decellularized porcine liver extracellular matrix (ECM) and to test its applicability for mimicking the matrix stiffening associated to liver fibrosis.
A protocol for decellularization and digestion of liver ECM (ddECM) developed in our laboratories was thus optimized to obtain suitable hydrogels with high cytocompatibility properties. The human hepatoma HepG2 cell line was chosen for the inclusions in ddECM hydrogels and different parameters such as vitality, morphology and proliferation were evaluated.
The matrix stiffening was induced by using the microbial transglutaminase (mTG) an enzyme that catalyzes the formation of covalent cross-links between glutamine and lysine in proteins. The effects of such treatment on micromechanical properties of hydrogels and on encapsulated cells were then evaluated.
Finally, the last part of this study was focused on the possible use of the optimized ddECM in the production of small-sized, alginate-free spherical hydrogel microcapsules with a new microencapsulation device (Spherical Hydrogel Generator) recently developed in Research Center “E. Piaggio”.
MATERIALS AND METHODS
P
REPARATION OF DECELLULARIZED LIVER MATRIX4.1.1 HEPATIC TISSUE HARVESTING
Porcine livers were obtained from healthy, 1-year-old freshly slaughtered animals at a local slaughterhouse (Pontedera – Pisa). Pig liver consists of five lobes (right lateral, right medial, left medial, left lateral and caudate lobe) wrapped in a tough fibrous capsule, i.e. Glisson’s capsule. Individual lobes, except for the small caudate one, were sectioned from collected livers. Liver lobes were then frozen at -20°C to induce cell breakage and to facilitate the following steps of decellularization.
This initial freezing is essential to break the plasma membranes. At the time of processing the liver was partially thawed at room temperature in a water bath, this because liver texture would make impossible the obtaining of the discs and so to avoid the denaturation of proteins, thawing must not occur through heat sources. Samples were then cut with a hollow metal cylinder of 14 mm diameter and the obtained cylinders were further cut in order to obtain 3 mm thick discs. The capsule of Glisson was then removed, and the disks were frozen until the time of their use.
Frozen livers were then punched with a tool to obtain 14 mm diameter cylinders, which were subsequently cut in 3 mm thick liver discs.
Figure 4.1 Porcine livers were obtained from healthy, 1-year-old freshly slaughtered animals
4.1.2 LIVER DECELLULARIZATION
Liver decellularization was performed as described by Mattei and collaborators (Mattei et al., 2014) with minor modifications (Table 1). Two different concentrations of the non-ionic detergent Triton-X 100 were used to obtain liver decellularization.
Briefly, liver slices (n. 20-25) were firstly washed in 1% w/v phosphate buffered saline (PBS) for 24 hours to remove blood residues. Subsequently samples were incubated in a 1% v/v Triton-X 100 solution prepared in PBS for 24 hours followed by a further incubation in a 0,1% v/v Triton-X 100/PBS solution for 12 hrs. Finally, samples were washed for 48 hours with PBS in order to remove any trace of contaminating detergent.
All the incubations (volume: 200 ml) were performed at 4°C on an orbital shaker at 200 rpm, changing the solutions approximately every 12 hours. Final samples were then stored at 4°C until further use.
Time(h) Day1 Day2 Day3 Day4 Day5 8:00 PBS1X Triton1% Triton0.1% PBS 1X PBS 1X 18:00 PBS1X Triton1% PBS 1X PBS 1X PBS 1X
Table 4.1Schematic decellularization process.
4.1.3 PREPARATION OF FINAL DECELLULARIZED AND DIGESTED ECM
Samples obtained after the decellularization step were first homogenized with a Ika Ultra-Turrax disperser tool, then lyophilized in a BenchTop Pro-Freeze Dryer VirTis SP Scientific for 12 hours and finally homogenized toa thin powder (mesh < 20). The final powder was then further processed with an acidic solution of porcine pepsin (Sigma) to achieve sample digestion. A mixture of 40 mg/ml of decellularized ECM with 4 mg/ml of pepsin powder was thus prepared in 0.1 M HCl and incubated for 36 hours under constant agitation. After 24 hours, a gelation test was performed of digested ECM at 37°C for 1 hour. Subsequently, 0.1 M NaOH and 10X PBS solutions were added to stop the enzymatic reaction and to obtain a final 30 mg/ml decellularized and digested matrix (ddECM) in PBS 1X with a pH of 7.4.
The remaining ddECM was then exposed to a source of Ultraviolet-C light (250-280 nm) for 15 minutes., transferred into a sterile vial and stored at -20°C until use.
CELL CULTURE AND SCAFFOLD INCLUSION
The HepG2, human cells line (Interlab Cell Line Collection, IRRCS AOU San Martino, Genova (GE), derived from hepatocellular carcinoma was used. Cells were cultured in Minimum Essential Medium (MEM) supplemented with 10% Fetal Bovine Serum (FBS, Sigma), 2mM L-Glutamine (Sigma), 1% non-essential amino acids (Sigma) and then were maintained in a humidified environment at 37°C with 5% CO2.
For cell inclusion in ddECM gels, 30 mg/ml matrix was routinely diluted to a final concentration of 10 or 15 mg/ml in complete culture medium. The cold (4°C) mixture was then used to resuspend aliquots of trypsinized cells to a final concentration of 30000 cells/gel of ddECM. Aliquots of 50 μl of cell suspensions were then transferred into a 96-wells microplate and incubated at 37°C/5% CO2 to achieve complete gelification. This ddECM volume was chosen to obtain 1 mm thick gels, since it seems to allow a complete diffusion of oxygen and nutrients (Pouliot R. et al 2016). After gelification, aliquots of 150 μl of complete culture medium were added to each well.
M
TG
EXTERNAL SOLUTIONTransglutaminase constitute a broad family of enzymes that catalyze the formation of cross-links between residues of glutamine and lysine in proteins. The modification of collagen with crosslinks is able to increase tissue stiffness. Among the various transglutaminases, the microbial transglutaminase (mTG) has been
Transglutaminases are enzymes that are generally considered to be of low toxic potential for HepG2 cells, as also confirmed by in vitro determinations on HepG2 cells (Balestri W. 2016). To induce gel stiffening, mTG was thus added to culture media after gelation at a concentration corresponding to 100 U of enzyme per gram of ddECM, as previously suggested (McDermot M. 2003).
To obtain the desired amount of mTG to be placed in 150 µl of medium solution, the following relation was used:
𝑄𝑚𝑇𝐺[𝑚𝑔 𝑚𝐿] = 𝑞(𝐸𝐶𝑀)[𝑔] ∗𝑥 [ 𝑈 𝑔] ∗ 1 0.1[ 1 𝑚𝐿] ∗ 1 100[ 𝑔 𝑈]
Where x stands for 0 or 100 (unit of mTG for gel).
The gels were stored in the incubator again at 37 °C. The mechanical tests were then performed 24 hours and 10 days later.
M
ECHANICAL TESTSFor the validation/comparison phase, the tests were done in collaboration with Research Centre “E. Piaggio”, anyway the procedure is here summarized. Compressive mechanical tests were performed using twin column ProLine Z005 testing machine (Zwick
Roell), while for the subsequent phase of viscoelastic
characterization, the tests were performed using Instron and a new bioreactor MechanoCulture TR (MCTR). ProLine Z005 was
equipped with a 10N load cell at room temperature.
The 𝜀̇ method developed by Mattei and collaborators (Mattei et al. 2015) at the Research Center “E. Piaggio” consists in the application
of a deformation rate 𝑑𝜀
𝑑𝑡 = 𝜀̇ constant at t = 0 obtaining the relative Young’s modulus. In this test 5 different strain rates were used, 0.01 - 0.005 - 0.001 - 0.0005 (1 / s).
The ddECM solutions were poured, into cylindrical molds (8 mm in height and 13 mm in diameter) and stored at 37 ° C for about 120 min. The samples were then removed gently from the molds and stored in Petri dishes and treated with mTG. At 24 hours and 10 days the samples were analyzed.
A
LAMAR BLUE METABOLIC ASSAYAlamar blue® metabolic assay (Invitrogen, UK) was used to determine the effects of decellularized ECM on cell viability and proliferation. The active ingredient of Alamar Blue® (resazurin) is a nontoxic, cell permeable compound that is blue in color and virtually not fluorescent. Upon entering cells, resazurin is reduced by the mitochondrial enzymes to resorufin, which is fluorescent. Viable cells continuously reduce resazurin to resorufin, thereby generating a quantitative measure of viability (Nociari et al., 1998). The metabolic cells activity was measured using Alamar blue after 24 and 48 hours to assess the cell viability and proliferation. Alamar blue working solution 1 mg/ml (10μl) was added to each well and incubated at 37°C for 120 minutes. Thereafter, aliquots of 100μl were transferred into a Costar-96-well plate (flat bottom, black), and the fluorescence was measured at excitation 485 nm and emission 615 nm. Although Alamar Blue is not a terminal assay, separate samples were tested at each time point and the experiment was
T
RYPANB
LUE EXCLUSION ASSAYCell viability and proliferation were studied with the trypan blue exclusion test. As described above, a number of 30,000 cells were encapsulated in ECM gels, seeded in triplicate in a 96-wells plate and treated with mTG at the indicated concentration.
After 48, 96, 168 and 240 hours from the encapsulation, all the gels were solubilized with the addition of a solution containing the enzyme collagenase H (COLLH-RO-Sigma-Aldrich®, St. Louis, MO, USA) prepared in serum free MEM. After an incubation of 10 minutes at 37°C and 5% CO2, collagenase activity was inhibited by the addiction of a 20mM sodium-EDTA prepared in Phosphate Buffered Saline (PBS). As control for cell proliferation, monolayers of HepG2 (30,000 cells) were seeded in 6-wells plates and harvested by trypsinization after 48, 96, 168 and 240 hours in culture. Finally, cell obtained from both gels and monolayers were stained with trypan blue and then counted using a hemocytometer under a light microscopy (Leica). Trypan bleu positive cells were considered dead, whereas viable cells with intact cell membranes were not stained o colored. Each assay was performed in triplicate. The number of viable cells obtained was compared with the values obtained at time zero, in order to estimate the rate of growth.
LACTATE
D
EHYDROGENASE(LDH)
C
YTOTOXICITY ASSAYLactate Dehydrogenase (LDH) leakage test, is one of the most commonly method used as a marker of cell damage. Upon cell death, LDH, is released in the culture medium where it can be detected. Samples of culture medium were thus collected at each time points, centrifuged at 300 xg for 5 minutes to remove large debris and stored at -20°C until analysis.
The LDH assay was performed using the Lactate Dehydrogenase Activity Assay Kit (MAK066-Sigma-Aldrich®, St.Louis, MO, USA) according to the manufacturer’s instructions with minor modifications. Briefly, 0,2,4,6,8,10 µl of a NADH standard solution corresponding to 0; 2.5; 5; 7.5; 10; 12.5 nmolee of NADH were added to the wells of a 96-well plate. A suitable volume of each samples (40 µl) was also added to the plate. Both standards and samples were brought to a final volume of 50 µl with the assay buffer supplied by the manufacturer. Subsequently, 50 µl of Master Reaction mix (48 µl of LDH Assay Buffer and 2 µl LDH Substrate Mix) was insert to each well and the plate was mixed with horizontal shaker, protected from light, for 2-3 minutes. After that, the initial absorbance was read at 450 nm (A(450) initial). The plate was then incubated at 37°C and other measure were performed after 5, 20, 40, 60, 75 minutes, until the value of the most active sample was greater than the value of the highest standard. The background was corrected by subtracting the final measurement obtained by the blank from the final measurement (A(450) final)of the standards and the standard curve was plotted. The change in measurement from Tinitial to Tfinal
and compared to the standard curve to determine NADH amount generate by the kinase assay.
LDH activity was then calculated according to the following system:
LDH Activity= B x Sample Dilution Factor Reaction Time x V
B = Amount (nmole) of NADH generated between Tinitial and Tfinal.
Reaction Time = Tfinal – Tinitial (minutes)
V = sample volume (mL) added to well
One unit of LDH activity is defined as the amount of enzyme that catalyzes the conversion of lactate into pyruvate to generate 1 µmole of NADH per minute at 37°C.
3D
ATP
ASSAY.
Standard colorimetric methods based on resazurin reduction (Alamar blue assay) or tetrazolium reduction (MTT assay), frequently used to assess number of viable cells in 2D cell culture, have been found not applicable to 3D spheroids/microtissues and collagen or other type of matrices. The CellTiter-Glo® 3D Cell Viability Assay, (Promega) is a bioluminescent ATP detection assay. It is robust, sensitive, and scalable to high-throughput screens, and offer relatively simple work-flow and data analysis. This type of assay quantifies a luminescent signal generated by the conversion of luciferin by luciferase as a function of cytoplasmic ATP concentration. Determinations were performed according to manufacturer’s instructions, with minor modification.
Briefly, 100 µl (out of 150 µl) of culture medium was removed from each well of the 96-well plate were hydrogels were cultured, leaving the microtissues in the remnant volume of the medium. Then 50 µl of reagent was added into each well and mixed for five minutes with a horizontal shaker protecting from light. For an effective lysis, plate was then kept for further for 20 minutes at room temperature. Finally, luminescence was read with a luminometer (PerkinElmer)according to manufacturer’s instructions.
ECM
-
BASED MICROENCAPSULATION USING AS
PHERICALH
YDROGELG
ENERATORThe Spherical Hydrogel Generator (Sphyga) was developed in Research Centre “E.Piaggio” (Tirella A. et al., 2014). It is a device composed of a syringe-based extrusion module combined with an external air system. Sphyga working parameters were chosen to generate consistently spherical hydrogel constructs with controlled dimensions. Microbeads were fabricated extruding the inks through a needle with an internal diameter of 210 µm using a flow rate of 10 µL/s. To produce cell-ECM-based microcapsules, 333 µl of ECM (10 mg/ml) and cells were mixed with 667 µl alginate (1% (w/v),
NovaMatrix, Norrway) then cells were resuspended to a final concentration of 2x 106 cells per ml of mixture. The droplets generated by Sphyga were made to land in a sterile 0,1M calcium chloride solution (Sigma Aldrich, Italy) and left for 10 minutes. After this the alginate-ECM microspheres were maintained in a humidified environment at 37°C with 5% CO2 for 30 minutes to enable ECM solidification and formation of 3D network fibers inside
by incubating the alginate-ECM capsules in a HEPES sodium citrate solution (2%) for 10 minutes. Pure alginate microcapsules were used as a positive control.
Figure 4.2 The Spherical Hydrogel Generator (Sphyga) for microbead fabrication: The Sphyga layout and the working principle, whereby multiple parameters are controlled to obtain microspheres with well-defined and controlled shape and size (Mattei G. et al 2013)
IMAGING
4.10.1 DOUBLE STAIN APOPTOSIS DETECTION KIT
Qualitatively, the cell viability was determined with a vital double staining that provides a rapid and convenient detection for the compacted state of the chromatin in apoptotic cells. Hoechst 33342, a kind of blue-fluorescence dye (excitation/emission maxima ~350/461 nm ， when bound to DNA), stains the condensed chromatin in apoptotic cells more brightly than the chromatin in normal cells. Propidium iodide (PI), a red-fluorescence dye (excitation/emission maxima ~535/617 nm，when bound to DNA), is only permeant to dead cells. The staining pattern resulted from
the simultaneous use of these dyes makes it possible to distinguish normal, apoptotic, and dead cell populations by fluorescence microscopy.
Briefly, after 48, 96, 168 and 240hours from the encapsulation, all the gels were incubated with the two dyes for 10 minutes at room temperature. Subsequently, gels were transferred on a glass microscope slide and fluorescence was evaluated with a fluorescence microscope (Leica) at 200× magnification. Images were captured by an on-line Leica DFC320 camera.
S
TATISTICAL ANALYSISAll calculations were performed using GraphPad software (GraphPad software, Inc., San Diego). Student't t-test , One-Way or Two -Way analysis of variance (ANOVA) followed by Bonferroni post hoc testwere used to evaluate statistical significance. p-Values below 0.05 were considered statistically significant.
RESULTS AND DISCUSSION The aim of the present Thesis was to set up a suitable model for in vitro 3D cultures by using a natural hydrogel obtained from a decellularized porcine liver extracellular matrix and to test its applicability for mimicking the matrix stiffening associated to liver fibrosis. There is an increasing interest in exploring the use of smart materials, both of natural and synthetic origin, to develop 3D models of cell cultures. It is well known that tissues stiffness may change in response to different biochemical and physical stimuli associated to pathological conditions such as chronic fibrotic disease and cancer progression (Hinz P. et al. 2013), so the analysis of matrix/tissue stiffness might produce new insights in understanding the pathological mechanisms of tumor and fibrotic diseases. The development of suitable 3D in vitro models could thus help to better understand the interactions between cells and ECM and how cells respond to mechanical changes in their microenvironment. In this perspective, the microbial enzyme transglutaminase (mTG) was used to mimic the matrix stiffening associated to a healthy-to fibrotic transition. Another field of active research is focused on the development of small-sized in vitro models allowing the most adequate oxygen and nutrients supplies to encapsulated cells (Haghgooie R. et al., 2010).
The potential use of a new microencapsulation device (Spherical Hydrogel Generator) recently developed in E. Piaggio center (Tirella A. et al., 2014) was thus tested.
