UNIVERSITÀ DI PISA
Research Doctorate School in Biological and Molecular
Sciences
Doctorate Program: Molecular Biotechnology
PhD thesis
Development of an in vitro model of healthy
and
diseased hepatic tissue using 3D matrices
Supervisors
Candidate
Prof. Arti Ahluwalia
Dr. Valentina Di Patria
Dr. Pascale Beffy
Advisor
Prof. Claudio Domenici
Contents
CHAPTER 1 ... 1
IN VITRO MODELS AS ALTERNATIVES TO ANIMAL TESTING ... 1
1.1. In vitro models ... 1
1.2. In vitro liver models ... 2
CHAPTER 2 ... 5
THE LIVER ... 5
2.1. Liver structure and metabolism ... 5
2.2. Liver extracellular matrix: characterization of the micro-architecture and protein content ... 7
2.3. Healthy and diseased liver: the role of matrix stiffness in regulating cells behavior ... 8 2.4. Liver fibrosis ... 9 2.5. Cytochrome P450 system ... 11 2.6. Multidrug resistance (MDR) ... 12 2.7. Epithelial-mesenchymal transition ... 15 CHAPTER 3 ... 19
DEVELOPMENT OF ENGINEERED CONSTRUCTS ... 19
3.1. Scaffolds for healthy and fibrotic hepatic in vitro models tissue ... 19
CHAPTER 4 ... 25
MATERIALS AND METHODS ... 25
4.1. Decellullarization of pig liver ... 25
4.1.1. Cell removal assessment: Histological analysis ... 26
4.1.2. Swelling tests ... 26
4.1.3. Cell removal assessment: DNA extraction and quantification ... 27
4.1.4. Protein content: Western Blot analysis ... 27
4.1.5. Micro-Tomografy (microCT) ... 28
4.1.6. Potential application as scaffolding material: dECM sterilization and cytotoxicity assay 30 4.2. Engineered scaffold for hepatic in vitro models ... 31
4.2.1. Collagen hydrogels and 3D Porous Protein Scaffolds (PPS) ... 31
4.2.2. Gelatin microspheres (μPPS) ... 33
4.2.3. Mechanical tests ... 34
4.3. Hepatocytes cell culture ... 34
4.3.1. HepG2 cell line and seeding on collagen hydrogels and 3D PPS ... 35
4.3.2. Primary Human Hepatocytes and seeding on collagen hydrogels ... 36
4.3.3. Human Upcyte® Hepatocytes and seeding on collagen 3D PPS ... 37
4.4.1. Viability, metabolic and morphology of hepatic in vitro models ... 38
4.4.2. Cell viability: CellTiter Blue® assay ... 39
4.4.3. Cell metabolism activity: urea synthesis, albumin secretion, and cytochrome P450 activity... 40
4.5.1. Cell morphology: immunofluorescence staining ... 41
4.6.1. Total cellular RNA extraction... 42
4.7.1. Statistical analisys ... 44
CHAPTER 5 ... 45
RESULTS ... 45
5.1. Pig liver decellularization ... 45
5.1.1. Histological analysis ... 45
5.1.2. Swelling behavior ... 46
5.1.3. DNA quantification ... 47
5.1.4. Protein content and Western Blot ... 47
5.1.5. 3D Microarchitecture ... 49
5.2. Role of substrate stiffness on hepatocytes ... 52
5.2.1. HepG2 model ... 52
5.2.2. Primary Human Hepatocytes model ... 61
5.2.3. Upcyte® Hepatocyte model ... 64
DISCUSSION AND CONCLUSION ... 73
AUTHOR’S PUBLICATIONS ... 77
Publications ... 77
Conferences ... 77
List of Abbreviations
ABC = ATP-binding cassette ATP = Adenosine triphosphate BSA = Bovine Serum Albumin cDNA = Complementary DNA DNA = Deoxyribonucleic acid CYP = Cytochromes P450 dECM = Decellularized ECM DPP IV = Dipeptidyl peptidase IV E = Young‟s elastic modulus E-cadherin = Epithelial cadherin ECM = Extra-Cellular matrix
EMEM = Eagle‟s Minimal Essential Medium EMT = Epithelial-Mesenchymal Transition EROD = Ethoxyresorufin-O-deethylase FBS = Fetal Bovine Serum
GTA = Glutaraldehyde
H&E = Hematoxylin and eosin HA = Hydroxyapatite HCC = Hepatocellular carcinomas HKGs = Housekeeping genes HRP = Horseradish peroxidase MDR = MultiDrug Resistance MDR1 = Multidrug resistance I
microCT = Micro-Computed Tomography μPPS = microspheres PPS
mRNA = messanger RNA
MRP = Multidrug resistance protein N-cadherin: Neural cadherin
PBS = Phosphate buffer saline PFA = Paraformaldehyde
PLGA = Poly lactic-co-glycolic acid PLLA = Poly-L-lactic acid
PPS = Porous protein scaffolds
qRT-PCR = Quantitative RealTime Polymerase Chain Reaction RGD = Arg-Gly-Asp
RNA = Ribonucleic acid RT = Room temperature
RT-PCR = Reverse transcription-PCR SDS = Sodium dodecyl sulfate
TCP = Tri-calcium phosphate TGF-b = Growth factor-b TMB = Tetramethylbenzidine TPC = Total protein content
1
Abstract
In vitro liver models are an important tool in hepatic tissue engineering as well as in
drug and chemical toxicity testing.
Recent studies have shown the importance of the several factors to preserve a differentiated hepatotypic phenotype, such as extracellular matrix (ECM) and its adhesive factors, the three dimensional architecture, the enrichment of the cell culture medium (e.g. growth factors, hormones, vitamins), the high cell density and the presence of other hepatic cell types.
The aim of this thesis is to develop new in vitro models of healthy and fibrotic human hepatic tissue for drug toxicity study. In particular the attention is focused on studying the influence of the hepatic fibrosis in the modulation of drug toxicity.
The research program is divided in two main sections: the development of protocols for the decellularization of pig liver in order to obtain matrices to be used as template for human liver scaffolds disegn (activity performed during the first year) and the development of hepatic in vitro models (second and third year), (Fig. 0.1). The first part of this project was also useful in determining the main properties of the hepatic extracellular matrix, which were then used for the selection of the scaffolding materials used in further in vitro models. In this latter section, the study was focused on the selection of „an optimal‟ hepatocyte cellular model and scaffolding material(s): different hepatocyte sources were grown on several engineered constructs evaluating differences in cell function and morphology.
The results obtained allowed to select a „healthy‟ and „diseased‟ hepatic in vitro models, able to preserve the hepatic phenotype of cultured cells in a 3D environment. These
2
In vitro models as alternatives to animal testing
1
Chapter 1
In vitro models as alternatives to animal testing
1.1. In vitro models
In vitro models are the obvious solution to the replacement of animal testing, and for
this reason are an important tool in tissue engineering as well as in drug and chemical toxicity testing, given that the complexity of the physiological environment is not replicated in petri dishes or microplates. “In vitro” refers to studies in experimental biology that are conducted using components of an organism that are isolated from their biological context in order to permit a more direct or more convenient analysis than that performed in-vivo using whole organisms.
Common examples of in vitro experiments include:
cells derived from multicellular organisms (cell culture or tissue culture) subcellular components (e.g. mitochondria or ribosomes)
cellular or subcellular extracts (e.g. wheat germ or reticulocyte extracts) [1].
In this way scientists can isolate specific cells, bacteria, and viruses and study them without the complexity of a whole organism. This permits an enormous level of simplification of the system under study [2].
It is common use to refer to in vitro models as cells cultured on a plastic 2D substrate (i.e. petri dish). Typically, these models use cell lines (derived from tumors or immortalized cells, and therefore different from the native ones) or primary cells, but in this case cells that can be kept in culture only for short time, thus posing a limit for some experiments.
In recent years many researchers highlighted the need of reproducing a more physiological in vitro cell microenvironment controlling the substrate (to be more similar to the extracellular matrix, ECM), the cell-cell interaction, and other important soluble factors (growth, adhesion, etc.).
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1.2. In vitro liver models
To date, no effective treatment for liver fibrosis has been reported, with the research mainly focused on the understanding of the mechanisms involved in the development of diseases or toxicity-induced liver fibrosis, and on the identification of potential pro- or anti-fibrotic properties of compounds. Studies on the development of anti-anti-fibrotic drugs are generally carried out on in vivo animal experiments, whereas in vitro models are mainly used in the understanding of the mechanisms involved in the development of liver fibrosis.
Several in vivo animal models (i.e. mouse) are used to mimic different causes of the development of liver fibrosis in humans as close as possible, e.g., by chronic administration of toxic compounds such as carbon tetrachloride [3], thioacetamide [4], dimethylnitrosamine [5], and galactosamine [6], or infection with Schistosoma [7].
An important disadvantage of experimental animal models, however, is that these models are largely restricted to rodents and may be of limited predictive value for human disease and drug toxicity leading to interspecies differences. In addition, in vivo fibrosis models give high discomfort to the animals used in experiments, thus they should be avoided when possible.
In fact, in 1959, Russel and Burch published “The Principles of Humane Experimental Technique” in which “the 3 R‟s principles” were proposed to encourage scientist in reducing the impact of research on animals.
“The 3 R‟s” stand for “Reduce”, “Refine” and “Replace”: Reduction:
Reducing the number of animals used in experiments by: - Improving experimental techniques;
- Improving techniques of data analysis; - Sharing information with other researchers; Refinement:
Refining the experiment or the way the animals are cared for so as to reduce their suffering by:
- Using less invasive techniques; - Better medical care;
- Better living conditions; Replacement:
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3
- Experimenting on cell cultures instead of whole animals; - Using computer models;
- Studying human volunteers; - Using epidemiological studies.
In November 2008, the European Union proposed to revise the directive of 1986 for the protection of animals used in scientific experiments in line with the three R‟s principles of replacing, reducing and refining the use of animals in experiments.
The proposals have two principal aims:
Improve the welfare of animals used in scientific procedures;
Ensure fair competition for industry to boost research activities in the European Union [8].
EU rules for animal experimentation are more restrictive, requiring that experiments using animals must be authorized and state that alternatives to testing on animals must be used when available and that the number of animals used in projects must be reduced to a minimum.
Moreover, in the legislative decree 26/2014 (art.13), procedures and methods for animal experiments in Italy have been reviewed [9].
The main advantage of in vitro models, apart from reduction of animal use, is the possibility to use human cells or human tissue to avoid species differences, the results will be of higher relevance to toxicity and disease in human.
In recent years, Van de Bovenkamp et al. [10], described some in vitro methods for this purpose, through the use of cells culture models (including primary cells and cells line), culture matrices, co-cultures and precision-cut liver slices. Several studies have shown the importance of the microenvironment with its extracellular matrix (ECM), other cells and other factors (i.e. growth factors, hormones, vitamins) to obtain functional and physiologically relevant in vitro models for studying human diseases, drugs, infections and molecular therapeutics.
The development of three-dimensional structures and dynamic systems using human liver cells may represent a valuable tool in the understanding of the molecular mechanism of some clinical aspects of inflammatory diseases [11].
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Chapter 2
The liver
2.1. Liver structure and metabolism
The liver is structurally and functionally heterogeneous and it is to be considered second only to brain in its complexity. It is one of the most important and complex organ and represents about 2% of the weight of an adult. It is involved both in endocrine and exocrine functions secreting factors such as albumin and urea into the blood, and secreting bile to the intestine respectively. It is involved in the storage function for glycogen, and in carbohydrate, lipid and amino acid regulation [12],[13]. In addition to its metabolic activity, the liver is one of body‟s fist lines of defense, inactivating toxins and xenobiotics absorbed by the intestine and clearing particles from blood [14].
The liver is an organ highly organized organ, in fact at least 15 different cell types can be found in normal liver (as summarized in Tab. 1)[15].
Table 1: List of cells found in the liver [5]
Hepatocytes (parenchymal cells) are the most numerous and comprise 60% of the total cells and 80% of the volume of liver. Other cell types include non-parenchymal cells such as
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the phagocytic Kupffer cells (macrophages), sinusoidal endothelial cells and stellate cells (fibroblasts).
Hepatocytes are large cuboidal epithelial cells with a diameter of 20-25 μm. Although they are epithelial cells, they do not form a surface, but rather form a tubular space between adjacent hepatocytes, which is delimited by tight junctions. These apical spaces fuse along the hepatic plate forming bile canaliculi, which drain secreted bile into the bile duct (Fig.1). The hepatocyte basal surface faces a 1.4 μm wide space (space of Disse), a thin reticular basement membrane composed of fibronectin, laminin, proteoglycan, collagen IV and I, this is the site of albumin secretion and lipoprotein uptake [13].
Figure 1: A basic diagram of interdependent liver cell types. Adapted from Mc Donnell et al. [16]
Hepatocytes are arranged in plates in repeating units named lobules, which appear in cross-section as hexagonal units spreading outward from a central vein (Fig.2).
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At the lobule vertices, a bile duct and a branch of the hepatic artery and portal vein are located close to one other in an arrangement often called the portal triad; blood flows from the two vessel branches toward the central vein through small vascular channels, termed sinusoids, lined with a fenestrated layer of endothelial cells. Plasma filters through the endothelium into the space that separates it from the hepatocytes (i.e. the space of Disse), and exchanges nutrients and metabolites with the hepatocytes through their basal surface.
Bile is secreted into the canaliculi formed between the apical surfaces of adjacent hepatocytes, and flows through the bile ducts into the common bile duct that dumps it into the duodenum [18].
Hepatic architecture and sinusoidal organization are essential for proper liver function. Loss of liver architecture due to trauma or disease, such as fibrosis, leads to loss of tissue function.
Kuppfer cells, ECM-producing stellate cells, biliary epithelial cells, hepatocyte precursor cells and fibroblasts are also present and perform important metabolic functions [19] . Liver cells are spatially organized to optimize communication and transport. Cells communicate directly through cellular and gap junctions, and via chemical signals dissolved and blood-borne or present in the macromolecules forming the ECM that surrounds them. The signals that cells exchange promote differentiation, proliferation and functions [20],[21]. Furthermore, metabolic (e.g., carbohydrate metabolism) and detoxification (e.g., CYP450 enzymes) activities of the hepatocytes change spatially along the length of the sinusoid, apparently regulated by gradients of oxygen, hormones and ECM composition, a phenomenon termed “liver zonation” [22].
Nevertheless, information on the structure-function relationship for normal and pathological liver tissue is still lacking.
2.2. Liver extracellular matrix: characterization of the
micro-architecture and protein content
The extracellular matrix (ECM) is the major component of the basement membrane, it is not only a physical scaffold, but also a crucial modulator of biologic process including cell attachment, migration, differentiation, repair, and development. It possesses several physiological and pathological functions such as modulate hepatic development, regeneration and even the maintenance of the normal architecture and differentiated states. Most of its components are large macromolecules with domains for various functions affecting
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differentiation, cell motility, adhesion, and other activities of cell. The extracellular matrix refers to the insoluble protein complex between cells that is composed of families of macromolecules consisting of collagens, elastin, glycoproteins, and proteoglycans (Fig.3). In addition, many other macromolecules are functionally related to the extracellular matrix, such as cell adhesion molecules together with their receptors, members of the transforming growth factor- (TGF- ) superfamily, and enzymes such as metalloproteinases [23].
The cell-ECM interaction plays a fundamental role in cell growth, organ development, tissue regeneration and wound healing, as well as in malignant growth processes: as in vivo cells attach to protein and carbohydrate domains present in the ECM [24].
Figure 3: Extracellular matrix. Adapted from Shimada et al. [25]
Many studies have shown that these structures provide support and anchorage for cells, but once cells are isolated from tissue and removed from the native matrix, differentiated cells rapidly lose their phenotype. In cell culture one of the main issues is to maintain functional hepatocytes, however when they are cultured with hormones, growth factors, serum free medium and ECM-derivatives, the hepato-cellular physiology (such as albumin synthesis and urea metabolism) can be maintained [13].
For this reason, in recent years, many researchers try to mimic in vitro 3D microenvironment using polymers with similar properties to extracellular matrix.
2.3. Healthy and diseased liver: the role of matrix stiffness in
regulating cells behavior
Recent studies have shown that the ECM biomechanical properties play an important role in regulating cell behavior [16].
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Tissue biomechanical properties are typically characterized by the Young‟s elastic modulus (E), expressed in Pascal (1 Pa = 1 N/m2). The E can be determined by sample deformation either in tension or compression, considering a sample with an area A and initial length L0, and measuring the resultant force (F). The force-displacement data are then
normalized to sample‟s area and initial length, respectively, to obtain stress and strain. The slope of the stress versus strain plot determines E. Although biological soft tissues (including liver [28]) typically exhibit an increase in stiffness with increasing strain (known as strain-stiffening effect), this slope is usually constant in the physiologically relevant small strain region.
To complicate the study, it has to be highlighted that the liver is one of the softest organs of our body. Several research groups have studied the elastic properties of liver, reporting elastic modulus values ranging from 0.6 to 1.0 kPa for a healthy liver [29], [30], [31], [32]. Several liver diseases, such as hepatitis B, hepatitis C, or hepatocarcinomas, cause a significant tissue hardening [30], with elastic modulus values increasing up to 1.6 kPa for fibrotic livers, depending on the fibrosis degree [32].
A high number of studies have showed that stiffness is one of the main factors in regulating cell growth and viability [33], and some of these showed that tumor markers expression is increased by matrix stiffness. To date only few studies were reported on the effects of matrix stiffness on hepatic metabolism and drug toxicity.
2.4. Liver fibrosis
Liver fibrosis is the common outcome of numerous chronic liver diseases with distinct etiologies, such as exposure to chemicals (e.g., alcohol liver disease, chronic drug toxicity), metabolic derangements (e.g., non-alcoholic steatohepatitis, Wilson disease, hemochromatosis), infectious diseases (e.g., viral hepatitis, schistosomiasis), and autoimmunity (e.g., primary biliary cirrhosis, autoimmune hepatitis) [34].
Liver fibrosis is defined by changes in both biochemical and physical properties of the cellular microenvironment and myofibroblasts are the major effector cells during the development of liver fibrosis. These multifunctional cells exhibit a lot of features: high proliferation rate, fibrogenicity through production and release of connective tissue proteins in the extracellular environment and secretion of proteases and their natural inhibitors involved in matrix re-modeling, contractility and immune-modulatory properties through the secretion and signal transduction of potent inflammatory cytokines, chemoattractants, growth factors
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and other bioactive mediators [35], [36]. This pathological process results from a chronic damage to the organ with a consequent excessive accumulation of ECM proteins, which is a characteristic of most types of chronic liver diseases. The accumulation of these proteins cause a deformation of the hepatic architecture by forming a fibrous scar, and hepatic fibrosis is the results of the wound-healing response of the liver to repeated injury [37]. After an acute liver injury (e.g. viral hepatitis) parenchymal cells regenerate and replace the necrotic or apoptotic cells, but if the damage persists the liver regeneration fails and hepatocytes are substituted with abundant ECM, and this accumulation causes an increasing of liver stiffness [31], [34], (Fig.4).
Figure 4: Hepatic architecture in normal and fibrotic liver, adapted from [34].
Recent studies demonstrated that modifications of mechanical properties of the liver could be related to the development of the hepatocellular carcinomas (HCC), in fact in many cases advanced levels of fibrosis preceded the generation of this type of cancer. Schrader et al. showed that the high stiffness environment encountered in chronic fibrotic liver disease fosters HCC progression by promoting cellular proliferation, a mesenchymal phenotype and resistance to chemotherapy. They demonstrated that human HCC cell lines Huh7 and HepG2 lose mesenchymal features, such as stress fibers, N-cadherin, and vimentin expression, and the cells up-regulate markers of hepatocyte differentiation when maintained in a soft environment [38].
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2.5. Cytochrome P450 system
As it is considered the major site of drug metabolism, liver diseases are know to be a possible cause of modifications of the pharmacodynamics and pharmacokinetics of drugs. The family of isozymes responsible for the biotransformation of several drugs is the cytochromes P450 (CYP). The name “cytochrome P450” (CYP) is derived from the spectral characteristics of this superfamily of enzymes, which present a maximum of absorbance at 450 nm – rather than at 420 nm as other hemoproteins – when in the reduced forms bind carbon monoxide. This spectral property of P450 is given by the presence of a thiolate cysteine linked to the heme group. Traditionally, biotransformation pathways are classified into two groups: phase I and phase II reactions, and CYPs constitute a superfamily of hemoproteins that catalyse the phase I biotransformation of many substrates and xenobiotics.
Animals, plants and microorganisms contain this multienzymatic complex P450. In mammals it has been found in all tissues examined, but P450 is found predominantly in the endoplasmic reticulum and mitochondria, and in greatest abundance in the liver, catalyzing reactions such as monooxygenation, hydroxylation, epoxidation, deamination. Therefore it can exert a detoxifying action, transforming a toxic or pharmacologically active substance in a harmless product. On the other hand, the P450 system can also catalyze oxidative reactions in which a substrate, originally inactive, can be activated and be mutagenic or carcinogenic [39]. For each isoform of P450 it is used a nomenclature based on the homology of the aminoacid sequence. The P450 superfamily is subdivided into families, subfamilies and individual isoforms. Six different P450 isozymes (i.e. CY1A2, CYP2C19, CYP2C9, CYP2D6, CYP2E1, and CYP3A4) play important roles in drug metabolism [40], [41]. Of these 6 isozymes, in this research project we focus the attention of the analysis of CYP-1A1, -1A2 and -3A4. In fact, it is well known that liver diseases are associated with decreased elimination of many drugs, particularly for those metabolized by enzymes of the cytochrome P450 family [42]. The capacity of the liver to metabolize drugs depends on hepatic blood flow and liver enzyme activity, moreover liver failure can influence the binding of a drug to plasma proteins [43]. Human CYP1A1, CYP1A2, CYP2B6, CYP2C8, CYP2C9, CYP2C19 and CYP3A4 are known to be inducible, whereas CYP2D6 is not.
CYP1A1 is expressed only at very low levels in mouse, rat and human liver, and it is essentially an extrahepatic enzyme that is present predominantly in the small intestine [44], [45], [46], lung [47], placenta [36] and kidney [37]. According to McDonnell et al. [38] the CYP1A1 catalytic activity varied considerably in human small intestine and microsomal
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preparations, and its expression may not be constitutively expressed, but only expressed after induction [39]. In fact, higher levels of CYP1A1 are often associated with increased smoking, physical exercise and ingestion of chargrilled meats.
CYP1A2 is expressed mainly in the liver and is not, or weakly, expressed in extrahepatic tissues in human [40], rat and mouse [37]. In human liver, CYP1A2 accounts for 13% of the total CYP content [41], [42] , and is involved in the metabolism of ∼ 4% of drugs on the market [43]. In human, CYP1A2 metabolizes several drugs, including phenacetin, tacrine, ropinirole, acetaminophen, riluzole, theophylline and caffeine [44].
The CYP3A subfamily is the most important of all human drug-metabolizing enzymes because this subfamily is involved in the biotransformation of ∼ 50% of therapeutic drugs on the market at present [52], although its content in the liver is only 30% of total P450. Some examples of drugs metabolized by CYP3A are terfenadine, the benzodiazepines midazolam and triazolam, quinidine, lidocaine, carbamazepine, nifedipine, tacrolimus, dapsone, erythromycin and dextromethorphan [52], [54]. In addition to drugs, CYP3A is involved in the oxidation of a variety of endogenous substrates, such as steroids, bile acids and retinoic acid [55]. Humans express four CYP3A enzymes, CYP3A4, -3A5, -3A7 and -3A43. CYP3A4 and its related -3A5 are the most abundant CYP isoforms in human liver, and are involved in the biotransformation of the majority of drugs [56].
Nakai et al., analyzed liver tissue of chronic hepatitis C patients and they have showed that mRNA levels of CYP1A2, CYP2E1, CYP3A4, NTCP, OATP-C, and OCT1 decreased as the fibrosis stage progressed and were measured after 24 h of cytokine exposure [57].
On the other hand Congiu et al. studied the mRNA levels in the liver of patients with various forms of liver disease. Results from patients with low fibrosis or inflammatory scores were compared with those with high scores, according to the Metavir scoring system [58]. They observed that levels of mRNA for CYP1A2, tended to be elevated in fibrosis groups with a score of 1 and 2 (mild and medium fibrosis respectively), and these were significantly higher than levels in the fibrosis groups with a high score (3 to 4), [59].
2.6. Multidrug resistance (MDR)
Many endogenous and exogenous compounds, including drugs, are eliminated from the body by the liver via metabolism and/or excretion. For this purpose, numerous transport proteins are available on the basolateral membrane hepatocytes to mediate uptake of amphipathic and polar organic compounds, as well as some lipophilic molecules, from
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sinusoidal plasma to hepatic cytosol. Moreover, these transport proteins play an important role in the excretion of drugs and metabolites from the hepatocyte. Unidirectional or bidirectional basolateral transport systems translocate polar molecules from hepatic cytosol into blood, whereas active canalicular transport systems are responsible for the biliary excretion of drugs and metabolites. Active pumps in the canalicular domain of hepatocytes help to remove drugs and their metabolites from the cell interior, and these pumps are called multidrug resistance protein (MRP). They are members of the ATP-binding cassette (ABC) superfamily of transmembrane transporters [60], [61], the most broadly expressed protein superfamilies known. ATP binding cassette (ABC) transporters bind ATP and use the energy of ATP hydrolysis to transport various compounds across cell membranes [62], including phospholipids, ions, peptides, steroids, polysaccharides, aminoacids, organic anions, bile acids, drugs, and other xenobiotics. These pumps are in the intestine, liver, kidney, and in other organs (Tab 2), and they have a large impact on the pharmacokinetics of numerous drugs.
Table 2: Distribution of human P-glycoprotein, MRP1 and MRP2 in selected tissue, adapted from Gilbert et al. [63].
The ABC transporter family has been divided into seven subclasses (A-G) [64].Members of the ABCB superfamily that are expressed in the liver include MDR1 or P-glycoprotein (encoded by ABCB1), an efflux pump for hydrophobic compounds; MDR3 (ABCB4), a translocator of phosphatidylcholine; and BSEP (ABCB11), the export pump for bile salts. These transporters are all located in the apical membrane of hepatocytes.
The ABCC subfamily consists of 12 members, five of which have been identified in liver tissue. The best characterized are the apical multi-drug resistance-related protein 2 (MRP2, ABCC2) and its basolateral homologues MRP1 (ABCC1) and MRP3 (ABCC3) [65], [66], (Fig.5).
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1) Importers, which mediate the uptake of nutrients into the cell (amino acids, sugars, ions and other hydrophilic molecules);
2) Exporters/Effluxers, which pump toxins and drugs out of the cell;
3) ABC proteins, which are involved in translation and DNA repair processes [67].
Figure 5: Schematic representation of the localization of MDR1, MRP1, MRP2, MRP3, MRP5. The physical barier between
the apical and baslateral membrane is formed by thigh junction, adapted from Ott et al. [68].
It is increasingly recognized that these and other transporters are involved in MultiDrug Resistance (MDR), which is one of the major causes of failure chemotherapy of human malignancies. The precise nature of chemotherapy resistance, and the potential role of drug resistance genes involved in the transport of anticancer agents, is still unclear. To date is known that drug resistance can be mediated by different mechanisms:
1) Increasing in the activity of ATP-dependent efflux pumps resulting in reduced intracellular drug concentrations. Agents commonly associated with this type of resistance include doxorubicin, daunorubicin, vinblastine, vincristine and paclitaxel [69];
2) Reduction of cellular drug uptake. Water-soluble drugs may attach to transporters carrying nutrients and therefore fail to accumulate within the cell. Resistance to drugs like cisplatin, 8-azaguanine and 5-fluorouracil is mediated by this mechanism [70];
3) The activation of regulated detoxifying systems such as the cytochrome P450 mixed function oxidases, and also of increased DNA repair.
4) In addition, resistance can result from defective apoptotic pathways due to malignant transformation [71], a change in the apoptotic pathway during exposure to chemotherapy [72], or changes in the cell cycle mechanisms that activate checkpoints and prevent initiation of apoptosis.
Other mechanisms involved in drug resistance include lack of drug penetration, modification of the ability to activate prodrugs and alterations in drug targets.
Human P-glycoprotein, encoded by the multidrug resistance I (MDR1) gene has a wide tissue distribution [73] and was the first ABC transporter identified to be overexpressed in breast cancer cell lines displaying MDR [74], and in other kind of tumors.
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But, in addition to their role in drug resistance, MRP1, MRP2 and MDR1 are expressed in non-malignant tissue, and are belied to be involved in protecting tissues from xenobiotic accumulation and resulting toxicity [60].
MRP1 is expressed at very low levels in healthy human liver, but its basolateral expression is induced during liver regeneration, and during endotoxin and bile duct ligation-induced cholestasis [75], [76], [77]. Ros et al. have showed that during severe human liver disease, MRP1, MDR1 and MRP3 expression results to be increased [75], so in this project we focused our attention on the expression of three pumps: MDR1 (apical) and MRP1 and MRP2 (basolateral).
2.7. Epithelial-mesenchymal transition
Epithelial cells are adherent cells that closely attach to each other, forming coherent layers in which cells exhibit apico-basal polarity. As all epithelial cells, hepatocytes must be polarized to be functional. Hepatocyte polarity is very peculiar and complex [13] compared to the “simple polarity” of most epithelial cells such as intestinal cells (Fig. 6). Usually, “simple polarized” cells have one apical pole located at the cell apex and one basolateral pole corresponding to the remaining domain of the plasma membrane. On the contrary, hepatocytes are organized in plates in the liver, so have several apical and basolateral poles [78].
Numerous studies have been carried out to find a way to keep the hepatocytes isolated from normal liver in a “well differentiated and polarized” status. For example, hepatocytes cultured on different substrates or in different matrices may preserve or change their polarized functional features [79].
Figure 6: Schematic representation of a simple epithelial cell and an hepatocte. In red are represented tight junction, adapted
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Mesenchymal cells, in contrast, are non-polarized cells, capable of moving as individual cells because they lack intercellular connections.
During the Epithelial-Mesenchymal Transition (EMT), the epithelium loses its polarity (e.g. adherent junctions, tight junctions, desmosomes, and cytokeratin intermediate filaments) in order to rearrange the F-actin stress fibers and expressing filopodia and lamellopodia. This phenotypic conversion requires the molecular reprogramming of epithelium with new biochemical instructions, during which cells gradually lose typical epithelial characteristics and acquire mesenchymal traits [80], [81], [82], in this transition it is observed an enhanced migratory capacity, invasiveness, elevated resistance to apoptosis, and greatly increased production of ECM components [83].
EMT can be classified into three different subtypes that cause different consequences, as described below.
Type I EMT is associated with implantation, embryo formation, and organ development in order to generate diverse cell types that share common mesenchymal phenotypes. This class of EMTs, neither causes fibrosis nor induces an invasive phenotype resulting in systemic spread via the circulation [84].
The EMTs associated with wound healing, tissue regeneration, and organ fibrosis are classified as the second type. In these type II EMTs, the program begins as part of a repair-associated event that normally generates fibroblasts and other related cells in order to reconstruct tissues following trauma and inflammatory injury. In contrast to type I EMTs, these type II EMTs are associated with inflammation and during organ fibrosis, such as during liver fibrosis, type II EMTs can continue to respond to ongoing inflammation, leading eventually to organ destruction.
Type III EMTs occur in neoplastic cells that have previously undergone genetic and epigenetic changes, specifically in genes that favor clonal outgrowth and the development of localized tumors [81].
Liver fibrosis can be classified as type II EMT, which is associated with tissue repair, but if the damage continues, it lead to the progressive loss of epithelial markers (E- cadherin, ZO-1) and gain mesenchymal markers (vimentin, alpha smooth muscle actin, FSP1 and β-catenin), (Fig.7) [81].
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Figure 7: EMT involves a functional transition of polarized epithelial cells into mesenchymal cells, adapted from Kalluri et
al. [81].
The most recognized inducers of EMT are growth factors acting through receptor tyrosine kinases, secreted signaling molecules in the Wnt and Notch families, and cytokines, such as transforming growth factor- (TGF- ) [85]. These transcription factors drive EMT by repressing expression of epithelial genes and activating expression of mesenchymal genes. Down-regulated genes include those encoding proteins maintaining epithelial cell-cell adhesions, such as the adherens junction protein E-cadherin, and the tight junctions proteins claudins and occludin. Up-regulated genes include those encoding proteins promoting cell migration and invasion, such as the mesenchymal cell-cell adhesion protein N-cadherin, the intermediate filament protein vimentin, and the extracellular matrix proteins fibronectin and collagen. On the other hand, remodeling of the actin cytoskeleton and how this is regulated are less well understood. Actin filaments in epithelial cells are organized in cortical thin bundles. In contrast, actin filaments in transdifferentiated mesenchymal cells are bundled into thick contractile stress fibers at the ventral cell surface [86].
Development of engineered constructs
19
Chapter 3
Development of engineered constructs
3.1. Scaffolds for healthy and fibrotic hepatic in vitro models
tissue
Primary human hepatocytes are considered to be the “gold standard” for in vitro toxicity tests [87], [88]. However, many studies have demonstrated that hepatocytes rapidly lose their phenotype and their specific liver functions when they are removed from their microenvironment. It is well-known that human hepatocytes are influenced by ECM interactions, soluble growth factors and cytokines, physical factors (e.g. stress and strain) [89], [90], and cell-cell communications [91]. Mainly, cell-ECM interactions play a fundamental role in hepatocyte growth [92], [93], liver organ development [94],[95], tissue regeneration [92], [96], [97], wound healing [96], [98], [99], and malignancy [94], [100]. Moreover, the liver ECM contains proteins and carbohydrates that provide support and anchorage for cells and regulate intercellular communication, for this reason, specific components from the ECM are commonly used as culture substrates and are commercially available. Some matrix components (e.g. collagen, fibronectin, laminin) have been used in cell culture for many years and have been showed to have profound effects on cells, both with respect to attachment and survival as well as for the maintenance of various functions [101], [102]. Some years ago Zhang. et al. [24] , reported that small differences in ECM composition between different tissue types can affect cellular interactions, hence it is highly recommended to use tissue-derived ECM substrates only for culturing cells of the specific tissue from which the ECM was obtained..
For this reason, many researchers focused their attention on the development of protocols for the decellularization of whole organs or tissues, maintaining the ECM architecture and content [103], [104], [105]. In fact, decellularized extracellular matrix represents an ideal material for tissue engineering applications as it retains relevant aspects of complex structure and chemical composition of the ECM. Moreover, what really makes decellularization an attractive technique for scaffold preparation in tissue engineering is the
Development of engineered constructs
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possibility to retain functional aspects of the native microvasculature and microarchitecture [106].
The potential application of decellularized matrix in tissue engineering has been demonstrated for a number of tissues including bladder [107], artery [108], esophagus [109], myocardium [110], skin [111], trachea [112], and whole heart [113]. Many decellularization methods have been described, such as the whole organ perfusion [114], [115], [116], using ionic or anionic detergents, and common decellularization techniques including pressure gradients [117], [118], [119] and immersion and agitation approaches [120], [121], [122].
In this project, several decellularization methods were tested, and finally a decellularization protocol was established and standardized. This protocol was then used to obtain 3-D substrates for in vitro applications, resembling the architecture and the protein content of a healthy liver. Decellularized ECM (dECM) samples were studied; retrieving important parameters (e.g. protein content, stiffness) which were then used to form 3D matrices able to mimic healthy and fibrotic hepatic tissue.
3.2. Three-dimensional structures: state of art
A number of methods able to reproduce in vitro organ architectures are widely described in literature. A class of these is represented by the porous 3D scaffolds, which are extensively used as a biomaterial in the field of tissue engineering for both in vitro study of cell-scaffold interactions and tissue synthesis, or for in vivo study of induced tissue and organ regeneration.
Instead of describing all the factors considered by bioengineers to produce a scaffold, below are listed only the main ones, while a deep description is given for the variables that are mainly involved in the preservation of the hepatic phenotype – as main topic of this research project.
Biocompatibility is the most important property for a scaffold since cells have to
adhere, migrate and proliferate the material, maintaining their viability and functions. Another important characteristic (mainly for tissue engineering applications) is the biodegradability, in fact once implanted, the scaffold must degrade and allowing cells to produce their own extracellular matrix.
As a matter of fact, the mechanical properties and the architecture of a scaffold are of critical importance for many in vitro studies.
Development of engineered constructs
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From an architectural requirement point of view, a scaffold should have an interconnected porous structure, as well as a high porosity, to ensure cellular penetration and adequate diffusion of nutrients to cells within the construct. In this 3D space there is also room for extracellular matrix deposition (that generally occur after few days/weeks of cell culture): in this way cells tend to re-model the microenvironment, in fact cells primarily interact with scaffolds via chemical groups (ligands) on the material surface. Therefore scaffolds produces using natural extracellular materials (e.g. collagen, gelatin, fibrin) naturally possess these ligands in the form of Arg-Gly-Asp (RGD) binding sequences, whereas scaffolds made from synthetic materials may require to include such ligands through, for example, protein adsorption. Additionally, cell adhesion and activity have been observed to vary considerably depending on the cell type, as well as the scaffolds composition and pore size [123], [124], [125]. Pore sizes of the scaffolds have to be large enough to allow cells to migrate into the structure, establishing a sufficiently high specific surface [126], [125].
Typically, three individual groups of biomaterials, ceramics, synthetic polymers and natural polymers, are used in the fabrication of scaffolds for tissue engineering with their advantages and disadvantages.
Ceramic scaffolds, such as hydroxyapatite (HA) and tri-calcium phosphate (TCP), are characterized by high mechanical stiffness (Young‟s modulus), very low elasticity, and a hard brittle surface, are typically used for bone regeneration.
Synthetic polymers, including the biodegradable polyesters PLLA (poly-L-Lactides) and PLGA (poly(lactic-co-glycolic acid)), and PLLA-PLGA copolymers, are used for the fabrication of 3D scaffolds for many organ in vitro models.
Biological materials and natural polymers are successfully used as scaffold biomaterials, such as collagen and derivatives, proteoglycans, alginates and chitosan.
Derivatives of the ECM have also been used as biomaterials, with few reports in liver studies. In particular ECM-derivatives are recognized as one of the optimal biomaterials since
in vivo cues - such as ECM moieties, cell signaling and soluble factors - are critical to
reproduce in a synthetic and functional hepatic in vitro construct. Collagen and hyaluronic acid, both ECM components, have been shown to play a key role in cell adhesion, proliferation, and signaling in vivo and they have been largely used to make hydrogel-based scaffolds. Other natural hydrogels such as Matrigel and alginate have been extensively used for 3D culture of hepatocytes due to their biocompatibility, mild gelling conditions and improved cell entrapment proprieties.
Development of engineered constructs
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Matrigel is composed of solubilized basement membrane proteins extracted from mouse chondrosarcomas. It consists of laminin, collagen IV, and heparin sulphate proteoglycan and is characterized by unknown ingredient concentration and quality that varies from batch to batch. Hepatocytes cultured on Matrigel have been shown to cluster into multicellular spheroids, but appear less polygonal than hepatocytes in vivo.
Alginate is generally extracted from seaweed, and it is considered an attractive biomaterial for its gelation proprieties (which can occur through the addition of ionic cross linkers or divalent cations such as Ca2+, Ba2+ and Sr2+). It is widely used as a drug delivery system as well as a scaffold for in vitro and tissue engineering studies [127].
Collagen is the most abundant structural protein of the extracellular matrix (ECM), and it has a high density of RGD sequences for cell adhesion and differentiation [128], [129]. Due to its biocompatibility and structure, collagen is widely used in biological and tissue engineering applications as a scaffold for cell adhesion and in vitro regeneration of tissues such as skin, cornea and vascular tissue [130], [131], [132], [125], [133]. For its biocompatibility and adhesion properties, type I collagen (derived from rat tail) was chosen as the candidate biomaterial for the production of 3D substrates – named porous protein scaffolds (PPS). A detailed description of PPS‟s properties is given below.
It has to be highlighted that in recent years, a number of studies report on the relationship between substrate stiffness and cell function and fate [134], [27], [135]. It is nowadays accepted that to reproduce a target tissue, it is necessary to match its mechanical properties. In this, dealing with collagen, it is critical to first understand and the control all the physical and chemical factors involved in collagen fibers alignment and interaction. Several studies have investigated methods for enhancing the mechanical strength of collagen-based hydrogels (e.g. controlling cross-link density) using different gelation and cross-linking methods [136], [133]. Bigi et al., described a cross-linking method for proteins involving the use of glutaraldehyde (GTA): aldehydes react with the amines of the collagen chains generating covalent inter-chain (i.e. cross-links) giving rise to a 3D network. Cross-linked collagen scaffolds are then freeze-dried, and protein porous structures are thus obtained (Fig. 8) [137]. Using this method, we developed 3D porous protein scaffolds (PPS) reproducing different grades of stiffness miming a healthy and fibrotic liver tissue (widely described in next chapters).
Development of engineered constructs
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Figure 8: SEM images of 3D collagen porous scaffolds: a) physically cross-linked collagen, b) collagen cross-linked with
Materials and methods
25
Chapter 4
Materials and methods
4.1. Decellullarization of pig liver
Livers were harvested from 1 year old healthy pig (kindly provided by slaughtrhouse Desideri, Pontedera, Italy). 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 [139]. Individual lobes, except for the small caudate one were sectioned, and some fresh samples had been taken for mechanical tests, liver lobes were frozen at -20°C until use. Frozen livers were thawed at 4°C overnight, then punched with a tool to obtain 14-mm-diameter cylinders, which were subsequently cut in 3-mm-thick liver discs, avoiding macroscopic vasculature and Glisson‟s capsule. Liver discs were frozen at -20°C until use. Several decellularization methods (Tab 3), based on immersion and agitation were investigated, varying chemical detergent and treatment duration.
Liver disc samples were thawed at room temperature and placed in 500 mL plastic bottle. The decellularization solution was added in a 20:1 v/w ratio with respect to the weight of liver disc samples,(i.e. 200 mL solution per 10 g of liver samples, about 20 discs). Then bottles were placed on an orbital shaker (SO3, Stuart Scientific, Stone, UK) at 200 rpm in a cold room, changing the decellularization solution twice a day for at least 3 days. Detergent solutions were prepared in deionised water.
The decellularization methods tested can be grouped into three major families: detergent-free (DF), ionic (I) and non-ionic (NI). First, a DF control family was provided to decouple the decellularizing effect due to mechanical agitation from that due to chemical agents (i.e., detergents). Triton X-100 (Sigma-Aldrich, Milan, IT) and sodium dodecyl sulfate (SDS) (Sigma-Aldrich, Milan, IT) were chosen as chemical decellularization detergents on the basis of their different modes of action and the reported effects on ECM [63]. Triton X-100 is a NI detergent that disrupts DNA–protein interactions as well as lipid–lipid, lipid– protein and, to a lesser degree, protein–protein interactions. SDS is an I detergent (i.e., anionic) that solubilizes cytoplasmic and nuclear cellular membranes, better at removing cell
Materials and methods
26
nuclei from dense tissues and organs than Triton X-100 is, but tends to denature proteins, disrupting the ultrastructure [68] and eliminating key growth factors [140]. Each protocol began with a rinsing day, to remove blood residues, and ended with a final washing day, to remove detergent residues. Furthermore, the capability of NI Triton X-100 to form micelles with anionic SDS [141] was used in this work to enhance residual SDS removal from the I protocols.
Table 3: Decellularization protocols used to obtain liver dECM; percentages refer to the weight/volume ratio (w/v) of
detergent solution deionized H2O. Only PBS 1X was used in the final washing day for DF protocols, since the contribution of 0.1% w/v Triton X-100 to cell removal is very poor and not significant compared with that of 1% w/v Triton X-100 or 0.1% w/v SDS used in the other.
4.1.1. Cell removal assessment: Histological analysis
To assess cell removal after the decellularization of liver discs, hematoxylin and eosin (H&E) staining and DNA quantification were performed for each of the nine protocols. Untreated frozen and thawed liver samples, henceforth termed fresh-frozen (FF), were used as controls. Decellularized liver ECM (dECM) discs were fixed and paraffin (Sigma, Milan, IT) embedded. The dECM samples were then cut into 5 µm sections, stained with H&E (Sigma, Milan, IT) and examined using an Olympus IX 81 micro- scope (Olympus Italia, Milan, IT).
4.1.2. Swelling tests
Liver dECM samples obtained with I and NI protocols were tested in triplicate (6 protocols × 3 samples). Decellularized liver discs were freeze-dried at -50°C, 0.45 mBar for 48 hours to determine their dry weight Wd. Then they were swollen in PBS 1X (Sigma,
Materials and methods
27
obtained, i.e., the equilibrium swollen weight Weq. The equilibrium mass swelling ratio was
calculated as Qeq ¼Weq=Wd. All measurements refer to the samples‟ equilibrium swollen
state, unless stated otherwise.
4.1.3. Cell removal assessment: DNA extraction and quantification
Total DNA content (micrograms per milligram equilibrium swollen sample) was extracted from samples obtained with all protocols and compared to that of fresh-frozen (FF) liver. Liver dECM was lyophilized after incubation at -80°C for 24 h, weighed and then placed into sterile 1.5 mL microcentrifuge tubes. Total DNA was isolated from 25 mg of equilibrium swollen tissue of FF liver and DF matrices, while 25 mg of freeze-dried tissue were used in case of ionic and non-ionic dECMs, hence concentrating any residual DNA, using the commercially available Kit Dneasy Blood & Tissue Qiagen (Valencia, CA), following the supplied protocol. The DNA concentration was estimated at 260 nm using a Nanodrop spectrophotometer (NanoDrop Technologies, Wilmington, DE), its purity was evaluated checking the absorbance ratio at 260/280 nm.
Total DNA content was expressed as μg of DNA per mg of fully swollen sample, multiplying the measured DNA content by the inverse of the equilibrium mass swelling ratio (1/Qeq) in case of ionic and non-ionic dECMs.
4.1.4. Protein content: Western Blot analysis
For biochemical characterization, total protein content (TPC) was determined using the Bio-Rad Protein Assay (Bio-Rad Laboratories Inc, Hercules, CA). FF and all decellularized liver samples were first equilibrium swollen in PBS 1X to determine their equilibrium wet weight (Weq), then solubilized in 1M NaOH solution (Sigma, Milan IT) at 60°C for 2 hours,
using 10 mL of sodium hydroxide solution per gram of wet sample (alkaline solubilisation). To obtain an absolute and repeatable measurement of protein concentration, care was taken to obtain a reproducible initial matrix and a meaningful standard curve subject to the same solubilization process as the samples. Solubilised samples were diluted 1:10 v/v in dH2O,
obtaining solutions with a sodium hydroxide concentration less than 0.1 M (i.e. 0.091M), so as not to interfere with the assay (as indicated by the manufacturer in http://www.bio-rad.com). The minimum wet sample dilution was then 1/110, assuming a liver density equal to that of water. In case of high protein concentrations, samples were further diluted with
Materials and methods
28
0.091 M NaOH. A protein calibration curve was established using bovine serum albumin (BSA, Sigma-Aldrich, Milan, IT) standard solutions solubilised in 1M NaOH at 60 °C for 2 hours and then diluted 1/11 in dH2O, as described for liver samples. The absorbance was read
at 595 nm (FLUOstar Omega, BMG Labtech GmbH, Offenburg, Germany) and expressed as mg of protein per gram of wet sample, considering sample dilution and
W
eq, thus obtainingcomparable data between decellularised and/or FF liver samples.
Moreover, key liver ECM proteins (i.e., laminin, fibronectin and collagen IV) were selectively analysed using Western blot.
Samples were homogenised on ice by Ultra-Turrax (Ultra-Turrax® T25, IKA, Germany) in a 10 mM Tris–HCl buffer (pH 7.8) containing protease inhibitors (leupeptin 4 μg/ml, aprotinin 1 μg/ml and phenylmethylsulfonyl fluoride 1 mM), then passed through a 22 gauge needle. In order to load the same total protein quantity (10 μg) for each of the analysed samples, all homogenates were diluted to the same protein concentration in a 0.1 M Tris-HCl buffer (pH 6.8) containing 143 mM β-mercaptoethanol, 0.4% w/v SDS, 7% v/v glycerol and 0.01% w/v bromophenol blue (final concentrations) and heated at 95°C for 5 min. All samples were separated by a 10% SDS-PAGE and gels were blotted onto nitrocellulose membranes (Whatman, GE Healthcare, UK). Membranes were stained with Ponceau S to verify loading and transfer efficiency. Nonspecific binding on the membranes was blocked with 5% w/v non-fat milk/Tris-buffered saline (pH 7.5) containing 0.1% Tween-20 (TBS-T) for 1 hour at room temperature. Membranes were then incubated with 1:1000 dilution of goat polyclonal antibody raised against porcine fibronectin, 1:1000 dilution of goat polyclonal antibody against porcine laminin or 1:500 dilution of goat polyclonal antibody against porcine collagen IV (Santa Cruz Biotechnology, Santa Cruz, CA) in TBS-T with 1% w/v (fibronectin) or 0.5% w/v (laminin and collagen IV) non-fat milk, overnight, at room temperature. The blot was washed three times in TBS-T and then incubated for 1 hour at room temperature with donkey anti-goat IgG secondary antibody conjugated to horseradish peroxidase (Santa Cruz Biotechnology, Santa Cruz, CA) diluted 1:5000 in TBS-T with 0.5% w/v non-fat milk. Bound proteins were visualised using an ECL detection system (Roche, Basel, Switzerland).
4.1.5. Micro-Tomografy (microCT)
Firstly we investigated the role of sample harvesting site and differences between animals on the bulk compressive modulus. Threecylindrical fresh porcine liver samples were collected from each of the 4 major liver lobes of four different animals, obtaining a total of 48
Materials and methods
29
fresh liver samples (4 animals × 4 lobes × 3 samples). A 14 mm diameter punch was used to collect regular liver cylinders that were subsequently cut in 3 mm thick samples using a custom slicer frame and a microtome blade. The Glisson‟s capsule was not present in tested samples and particular attention was dedicated to avoid macroscopic vasculature. To ensure a repeatable sample testing state, all liver samples were equilibrium swollen in PBS 1× at 4 °C, then brought to room temperature and carefully measured using a calliper prior to testing [32]. Hepatic tissue was considered as mechanically isotropic [142], [143] . To investigate the effect of a freeze-thawing cycle on bulk liver compressive modulus, frozen lobes were thawed at 4 °C overnight and then samples were collected and tested as previously outlined. Finally, liver dECM samples obtained decellularizing each of the four swine livers with protocols NI3 and I3 were tested in triplicate (4 animals × 2 decellularization protocols × 3 samples). The ECM discs maintained their gross cylindrical shape throughout all immersion and agitation decellularization procedures, in agreement with results from Lang et al. [120]. Then, they were equilibrium swollen in PBS 1× at 4 °C and brought to room temperature, as for non-treated samples, to ensure a repeatable testing state. The actual sample dimensions were measured with a calliper just prior to testing, hence accounting for any variations in diameter and thickness due to the decellularisation procedures. All samples (equilibrium swollen fresh, fresh-frozen and decellularised porcine liver) were placed in a glass Petri dish and partially immersed in PBS 1× to preserve their hydration during the unconfined compression test. As with most soft biological tissues, mechanical testing of liver samples is a challenge. We adopted the M testing configuration which we have recently proposed to overcome some of the issues in testing extremely soft and highly hydrated materials [144]. Briefly, the Petri dish was positioned above the bottom plate of the testing machine (Zwick/Roell ProLine Z005), while the upper plate (connected to the load cell) was manually approached to the top of the sample, avoiding any contact with it, thus preventing sample pre-stress. Force (F) and compressive displacement ( l ) data were recorded over time while compressing samples at a fixed quasi-static strain rate ( ) of 0.01 s-1 at room temperature. Measuring the mechanical properties of these soft samples is very challenging and the adopted testing configuration allowed precise determination of the initial point of the compressive phase. As the upper plate of the testing machine approaches the sample the measured force is equal to zero. Then it becomes negative at the instant of contact, due to water surface tensions. The instant in which the plate starts to actually compress the sample is taken as the point at which the force-displacement curve crosses the abscissa and then the force increases monotonically. In this point F is zero by definition, while l can be easily re-zeroed subtracting an offset (
l
offset),Materials and methods
30
giving the actual sample compression (
l
reall
l
offset). Considering this as the initial point ofthe compressive phase, measured force and actual compressive displacement data were respectively normalised to the cross-sectional area (A) and the initial length (
l
0) of the sample, obtaining engineering stress ( F /A) and strain (l
real/ l
0) data. Compressive moduli were then derived as the slope of the first linear tract ( 0.03) of the stress-strain curve.4.1.6. Potential application as scaffolding material: dECM
sterilization and cytotoxicity assay
Sterilization procedure
Freeze dried NI3 dECM samples were sterilized using four different methods: 1) overnight exposure to chloroform (Carlo Erba, Milan, Italy) vapour,
2) overnight exposure to chloroform vapour followed by gas plasma sterilisation (Sterrad 100S, Advanced Sterilization Products, Irvine, CA),
3) gas plasma sterilisation only,
4) for 2 hours soaking in 0.1 % peracetic acid (Sigma-Aldrich, Milan, IT). In particular chloroform sterilization was performed by pouring 20 mL of solvent on the bottom of a 5 L dessicator. The liver matrices were placed on the dessicator ceramic support plate about 3 cm above the liquid and the dessicator lid closed in order to expose the samples to chloroform vapour overnight at room temperature.
Cytotoxicity assay:
Cytotoxicity experiments were performed using HepG2 cells from ATCC (American Type Culture Collection, passage 83) in Eagle's minimal essential medium (EMEM, Sigma-Aldrich, Milan, IT) supplemented with 10% fetal bovine serum (Sigma-Sigma-Aldrich, Milan, IT) and 1% penicillin, 1% streptomycin and 1% L-glutamine (Invitrogen, Milan, IT). 50 000 cells per well were seeded in a 24 well plate with 1 mL of medium and left to adhere and proliferate for 48 h. Liver dECMs, from protocol NI3, were well washed in sterile PBS 1× before being placed in the cell-seeded wells. Cell viability was assessed at 24 and 48 hours using CellTiter Blue (Promega, Madison, WI). HepG2 seeded on tissue culture plates were used as controls.
Materials and methods
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4.2. Engineered scaffold for hepatic in vitro models
4.2.1. Collagen hydrogels and 3D Porous Protein Scaffolds (PPS)
Collagen-based scaffolds, as hydrogels or as PPS, were fabricated modulating the final stiffness by controlling the cross-links (herein expressed as glutaraldeyde/collagen ratio [mol/g]). With this approach collagen substrates were conceived in order to mimic both healthy and fibrotic hepatic tissues, and specifically the latter with different grades of fibrosis. As already mentioned, collagen is the most abundant structural protein of the ECM, and it is commonly used in biological and tissue engineering applications as a scaffold for cell adhesion and in vitro regeneration of tissue [130], [133]. In this study, type I collagen was previously extracted from rat tails using the method described by Elsdale and Bard [145]. An acidic solution of collagen is cross-linked first using a physical gelation mechanism in order to shape the construct (physical gelation occur changing the pH in a temperature controlled system); and later a second crosslinking step (i.e. chemical gelation) is used to modulate the number of crosslinks, thus the stiffness (i.e. using glutaraldehyde to link amines in collagen chains). Crosslinks number was expressed as molGTA/gcoll.
Briefly, type I collagen was diluted to a final concentration of 5.56 mg/mL with a 0.02 N solution of acetic acid. Then, 10X concentrated M199 medium (previously adjusted to pH 9.3) was added keeping the collagen solution in contact with ice (this is critical to slow down the gelation). Both hydrogels and 3D porous collagen scaffolds were prepared in a multiwell plate (24- or in 48-multiwell plate, accordingly to the further cell experiment) obtaining samples of about 1 mm in thickness. Samples were kept at 37°C for 1 hour to form a stable physical hydrogel. After this first step (physical gelation), samples were chemically crosslinked by adding a solution of GTA. Different GTA concentrations (ranging from 2 mM to 200 mM, as reported in tab. 4) were used to obtain various sample crosslinking grades (expressed as molGTA/gcoll) recapitulating the stiffness of both healthy and fibrotic livers.
Non-crosslinked scaffolds (0 mM GTA) were used as control (Fig. 9). Please note that the maximum concentration limit of 200 mM GTA was chosen to avoid any toxic effect [146].
