Smart modular scaffolds (SMSs) for the realisation of 3D in-vitro organ models

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UNIVERSITY OF PISA

D

OCTORAL

S

CHOOL OF

E

NGINEERING

“L

EONARDO DA

V

INCI

”

P

H

.D.

PROGRAM IN

C

HEMICAL AND

M

ATERIALS

E

NGINEERING

XXVI

CYCLE

Smart modular scaffolds (SMSs) for the

realisation of 3D in-vitro organ models

(SSD:

ING-IND/34)

Author:

Giorgio M

ATTEI

Tutor:

Prof. Arti AHLUWALIA

Dr. Giuseppe GALLONE

Dr. Eng. Annalisa TIRELLA

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Abstract

Animal testing still continues to be the prevailing standard in the development of most of the drugs, treatments and cures for human diseases. In addition to elevated cost, laborious process and ethical issues associated with the use of animals, it is now well consolidated that animals are scarcely representative of humans, and thus poor models to predict the clinical efficacy and outcome of certain therapies in humans. These limitations have generated the demand for more affordable, efficient and standardised platforms for systematic, reliable and quantitative studies of tissue pathophysiology, disease mechanisms and development of new drugs and treatments, leading to the development of

in-vitro organ models. Although 2D in-in-vitro models have made significant

contribution to biological research, the current consensus is that 3D models provide a cellular micro-environment closer to the native one and are critical to replicate tissue functions in-vitro.

Owing to its central role in both endogenous and exogenous metabolism, the liver is one of the most studied organs in the human body. Several diseases as well as the physiological process of tissue ageing can cause liver fibrosis, which may lead to cirrhosis and subsequent liver failure: one of the major cause of illness and death across industrialised countries. No effective treatment for liver fibrosis is available yet, and extensive research is still on-going to investigate fibrotic processes and identify potential anti-fibrotic drugs, raising a continuing

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demand for new physiologically relevant in-vitro liver models. In this context, this thesis is focused on engineering a pathophysiological 3D in-vitro liver model able to mimic healthy hepatic tissue as well as ageing and fibrotic processes. The 3D liver model was developed following the classical Tissue Engineering approach, which is based on the successful interaction between i) cells of a given target tissue, ii) a scaffold serving as a supportive template for cells promoting their adhesion, proliferation and development, and iii) a bioreactor for the dynamic cultivation of the cell-scaffold construct providing cells with appropriate physicochemical cues to direct them towards the expression of the desired tissue phenotype. This thesis covers the entire design and engineering process to develop the pathophysiological 3D in-vitro liver model, according to the bottom-up strategy shown in Figure 0.1.

Figure 0.1: Bottom-up strategy followed to engineer the pathophysiological 3D

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Abstract

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In particular, since an ideal scaffold should mimic most of the properties of the native extracellular matrix (ECM), several decellularisation procedures were investigated to obtain liver matrices which were biochemically and mechano-structurally characterised to derive ideal design specifications for ECM-mimicking scaffolds. Given the critical role of matrix stiffness in regulating hepatic cell response and in directing the development of tissue fibrosis, attention was focused on the mechanical properties of decellularised hepatic tissue. Then, hydrogel-based smart modular scaffolds (SMSs) for hepatic cell encapsulation were designed to obtain supports that mimic the stiffness of healthy liver ECM and can subsequently be enzymatically stiffened to recreate fibrotic environments. The enzymatic stiffening was designed to recapitulate the enzyme-mediated hardening typical of ageing and fibrotic processes in hepatic tissue. Afterwards, a transparent bioreactor (TB) was designed and realised for dynamic cultivation and real-time monitoring of the hepatocyte-laden SMS constructs. Finally, experiments with HepG2 liver cells encapsulated within SMSs and cultured in the TB were conducted to assess the suitability of both SMSs and TB developed during this thesis as a platform for engineering pathophysiological 3D in-vitro liver models for a vast range of applications, such as drug development, prediction of the ADMET (i.e. adsorption, distribution, metabolism, elimination and toxicology) properties and the clinical efficacy of new potential drugs and treatments, mechanistic studies, chemical testing and disease modelling.

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Acknowledgements

This PhD was a very great experience for me, from both scientific and life standpoints. During the last three years I learned a lot of new things and I would like to thank very much all the people who helped, sustained and encouraged me and my research, allowing me to grow as a research scientist. I will do my best to remember everyone, but knowing my mind, let me thank here all the people who are not mentioned below.

Foremost, I would like to express my sincere gratitude to my tutors Prof. Arti Ahluwalia, Dr. Giuseppe Gallone and Dr. Eng. Annalisa Tirella for their continuous support during my PhD project, for their motivation, enthusiasm and great knowledge. Their guidance, advices and encouragement as well as their insightful observations and comments helped me in all the time of research and writing of this thesis. Thank you very much.

Besides my Italian tutors, I would like to thank a lot also Prof. Nicola Tirelli for having given me the opportunity to join his research group at the University of Manchester and guided me in the world of polymer chemistry, hydrogel design and characterisation.

I am grateful also to my internal examiner Prof. Luigi Lazzeri for having reviewed my work and for his precious comments and advices.

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vi How to forget all the labmates and friends???

Many thanks to all the people at the Research Centre “E. Piaggio” (Serena, Tommaso, Annalisa, Chiara, Danielino, Gianni, Daniele, Nicole, Abu, Giovanni, Carmelo, Francesca, and all the other guys as well) and at the IFC-CNR in Pisa (Federico, Valentina, Dottor Domenici). Thank you all for your friendship and support, and for the moments we spent together.

I would like to thank also all the people I met in Manchester. Thanks Rob, Arianna, Ghislaine, Chris, Enrique, Jenny, Ale, Erwin, Aziz and Francesco for your help and the good times we had together (although too much rainy…). Moreover, I really want to thank all the guys who play and who have played with me in several bands, since I actually can’t live without music. Thank you "Studiolive" (Piera, Andrea, Francesco), "Mr. Piera & The Forever Young" (Piera, Francio, Giova, Rocco, Maestro Caruso, Pablo) and "Emmeciesse" (Andrea, Marco, Gianluca, Maestro Caruso).

Many thanks also to my closest friends (Marco, Francesca, Nicola and Cristina) for all the happy moments and the great holidays we spent together.

A special thank goes to my family. Words cannot express how grateful I am to my father Marco, my mother Roberta and my brother Gianluca for having supported and encouraged me to strive towards my goals. To them I dedicate this thesis.

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Introduction

Most of the drugs, treatments and cures for human diseases available to date have been developed using animal models. In spite of many concerns about availability, feasibility and ethics related to the use of living subjects, it is now well consolidated that animals are scarcely representative of humans, and thus poor models to predict the clinical efficacy and outcome of certain therapies in humans. Although the use of animals gives rise to ethical concerns [1], the scientific community attempts to justify this to society focusing on its critical role in predicting human response to drugs and disease [2,3], in agreement with Giles’ comment in Nature [4]:

“In the contentious world of animal research, one question surfaces time and again: how useful are animal experiments as a way to prepare for trials of medical treatments in humans? The issue is crucial, as public opinion is behind animal research only if it helps develop better drugs. Consequently, scientists defending animal experiments insist they are essential for safe clinical trials, whereas animal-rights activists vehemently maintain that they are useless.”

How far can we go with animal models?

Animal experiments are often poorly designed, conducted and analysed. Methodologically adequate reviews and summary of evidence from animal research are generally lacking, contributing to the failure in translating animal

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results to humans [5]. Pound et al. pointed out some potential drawbacks in using animal models including: i) differences in drug administration between animals and humans, ii) variability in the selection and randomisation of animals used for the study as well as in the choice of comparison therapies and data reporting, and iii) selection of measured parameters and length of the follow up that may be not relevant for the investigation of the human clinical outcome [6].Just to give an example, a large number of vaccines were found to be effective against a HIV-like virus in animal models, but none of them have prevented HIV in humans [7]. The failure of animal models can also be noticed in the extremely low number of basic research papers leading to a new class of drugs [8] and in the poor number of animal researches translated at the level of human randomised trials [9].

Biological systems are very complex and exhibit emergence, i.e. new properties arising from the interactions of the parts that cannot be explained or predicted with a reductionist approach dissecting such complex systems into their constituent parts and studying them individually [10]. Moreover, complex systems are resistant to changes thanks to their redundancy, exhibit self-organisation, have hierarchal levels of organisation and feedback loops, interact dynamically with their environment and are very dependent upon initial condition (e.g. genetic make-up). Even among humans, very small differences in genetic make-up can result in dramatically different outcomes to perturbations such as drugs and disease, as observed for different sex [11,12], ethnic groups [13,14] and even between monozygotic twins [15]. As a consequence, predicting intra-complex system response is very challenging, while predicting inter-complex system response is likely impossible. In this regard, since animals and humans are complex systems evolved following

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Introduction

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different evolutionary trajectories, it is possible to conclude that animal models would fail to be predictive for human response to drugs and diseases. Cadaveric tissues can partially address these problems, but still presents issues related to availability and maintenance of excised tissues, as well as donor-specific factors (e.g. disease or genetic variations) that can affect the experimental outcome. These limitations have drawn researchers’ attention to look for more standardised and affordable platforms for drug testing and cell characterisation studies, leading to the development of in-vitro models.

In-vitro models: a meaningful solution

Compared to animal models and cadaveric tissues, in-vitro models permit systematic, reliable and quantitative investigations of cell and tissue/organ pathophysiology to study disease mechanism and drug efficacy. Furthermore,

in-vitro models can be more tightly controlled and are generally less expensive

and less time consuming than animal models. Therefore, they are very suited for high throughput screening (HTS, i.e. the investigation of a large number of different combinations of experimental parameters) which is often not feasible with animal models. In addition to these advantages, the establishment and validation of innovative in-vitro models can contribute significantly to refine, reduce and replace the use of animals in biomedical research, according to the 3Rs’ principle introduced by Russel and Burch [16] and in line with the recent policies of the European Union (EU) and of the United States (US), which solicit more innovative approaches to toxicity testing and the reduction of animal based studies [17–19].

In-vitro models: 2D vs 3D systems

The most widespread in-vitro model is represented by the cell monolayer, i.e. a layer of cells cultured onto bi-dimensional (2D) substrates, such as tissue

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culture plates or flasks. Although 2D models have made significant contributions to biological research, cells or tissues cultured on 2D substrates do not mimic cell growth in-vivo and express tissue specific genes and proteins at different levels. This is owed to the fact that 2D cultures are too simple to represent the complexity of tissues/organs in-vivo, with cells immersed in a three-dimensional (3D) environment composed of the extracellular matrix (ECM), soluble factors and products coming from homo- and hetero-typical cell–cell interactions. These limitations have prompted the development of 3D models, providing a cellular micro-environment closer to the native one, which is critical to replicate tissue functions in-vitro since cell behaviour is strongly dictated by environmental cues [20–22]. Compared to micro-patterned and microfluidic 2D models, 3D systems are better representative of the in-vivo environment, specifically in terms of i) control of concentration gradients of signalling molecules and/or therapeutic agents, ii) ECM composition and structure around cells and iii) cell adhesion, morphology and arrangement [23]. Moreover, 3D models can mimic physiological barriers that hinder species transport to target cells in-vivo, being critical in many research studies, such as drug discovery (2D models may lead to candidates that cannot reach target cells

in-vivo). In the last few years, 3D in-vitro models have gained increasing

interest from the research community thanks to the development of new enabling technologies (e.g. computer-aided tissue engineering, rapid prototyping, micro-fabrication techniques) [24–26], and successfully used to engineer 3D constructs of bone [27,28], liver [29,30], skin [31,32], cardiac [33,34] and other tissues. Offering valuable advantages for the investigation of micro-environmental effects and spatiotemporal constraints on different types of cells, 3D in-vitro systems have become complementary to cell monolayer

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Introduction

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and animal models in pathophysiological organ modelling and drug screening as well as in pharmacokinetic and pharmacodynamics studies.

Implementing 3D in-vitro models: the Tissue Engineering approach The field of Tissue Engineering (TE), which is aimed at recreating tissues and organs in-vitro for replacing diseased or missing parts, is based on the successful interaction between three components: i) cells of a given target tissue, ii) an appropriate scaffold that serves as a template to host these cells and promoting their adhesion, re-arrangement, proliferation and development, and iii) a bioreactor to cultivate the cell-scaffold construct providing cells with suited physicochemical cues to direct them towards the expression of the desired tissue phenotype [35,36]. This approach can be attractive also to realise 3D in-vitro organ models, which are thus represented by the entire cell-scaffold-bioreactor system.

It is known that an ideal scaffold should: i) support cell growth and proliferation, ii) provide cells with an appropriate environment mimicking most of the mechanical, chemical and biological features of the native ECM, and iii) allow for an effective transfer of nutrient, gas exchange (e.g. O2 and CO2),

metabolic waste removal and signal transduction. The ECM is a mixture of glycosaminoglycans and fibrous proteins (e.g. collagen, elastin, fibronectin and laminin) that fill the extracellular space between cells. It is a dynamic structure that provides structural and anchoring support to the cells, and it is constantly remodelled by the latter during tissue development, homeostasis and wound healing by balancing its synthesis and degradation through a variety of enzymes (e.g. matrix metallo-proteinases, MMPs) [37]. Moreover the ECM contributes to direct cell fate and function through cell–ECM interactions. Other signals from the microenvironment can come to a cell through i) soluble factors in the

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interstitial fluid, ii) cell-cell interactions, or iii) mechanical forces. These signals are transmitted within the cell, inducing a cascade of interactive events at several hierarchical levels, such as i) molecular (e.g. changes in gene expression), ii) subcellular (e.g. cytoskeleton remodelling) and iii) cellular (e.g. changes in cell shape, motility, proliferation, differentiation, apoptosis, and in the morphogenesis of cellular structures) [21,38]. The communication between parenchymal and stromal cells is also important in studying cellular micro-environment, and is mediated by i) adhesion molecules, ii) ECM connecting these cells, or iii) soluble factors secreted by them. Over the last few decades, several materials have been proposed to fabricate scaffolds for many tissue engineered constructs. There was a significant interest in hydrogels due to their advantages in replicating the features of the native ECM [39,40]. Hydrogels are 3D cross-linked insoluble, hydrophilic networks of polymers, particularly suited for both tissue engineering and in-vitro organ model purposes, since their physicochemical and mechano-structural properties can be easily modified to desirable values for several applications (e.g. cell encapsulation [41], immobilisation [42] and drug delivery [43]). To date, the biocompatibility of various natural (e.g. collagen, hyaluronic acid, fibrin, alginate, agarose, chitosan) and synthetic (e.g. poly(ethylene glycol), poly(lactic acid), poly(glycolic acid) and copolymers poly(lactic-glycolic) acid) hydrogels has been well characterised. Moreover, available nano- and micro-fabrication technologies (e.g. nano- and micro-fluidics, micromolding, lithography and biopatterning) allow the engineering of complex 3D shaped scaffolds [44,45]. Combinations of natural and synthetic hydrogels were also proposed to fabricate bio-artificial (or semi-synthetic) scaffolds with enhanced biological, biophysical and mechanical properties [46]. Several hydrogel-based cell-encapsulating constructs were investigated, proving their suitability as

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Introduction

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promising tools for the generation of 3D functional tissues for organ repair/regeneration or other applications, such as in-vitro models for investigating tissue pathophysiology or for drug testing and toxicological assays [47].

In general, three-dimensional cell constructs have significantly higher metabolic requirements than traditional cell monolayers (2D static cultures). In this regard, bioreactors are necessary to increase the mass transfer rate and facilitate the long-term supply of gases and nutrients as well as a proper removal of metabolic waste to and from the entire volume of the 3D engineered construct. A cell culture bioreactor can be generally defined as an engineered device for culturing cell constructs in a tightly controlled and closely monitored environment, providing cells with suitable spatiotemporal biochemical and physical cues to promote their reorganisation and differentiation into the target tissue/organ. Thanks to the great progress of new enabling technologies and the contribution of computational fluid dynamics, many bioreactors have been proposed to date, from the simplest stirred culture systems, to more complex versions providing cultures with a combination of different physicochemical cues and allowing for real-time monitoring of the growing tissue throughout the culture period [48]. However, despite all of these technological efforts, providing optimal conditions for the long-term maintenance of tissue-like 3D constructs still remains a challenge at the moment, given the complexity of many tissues and organs as well as the number of environmental cues affecting cell behaviour and function [20–22].

The liver: role and pathophysiology

In the context of 3D in-vitro organ models, this thesis is focused on the liver, which is one of the most well studied organs in the human body, by way of its

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central role in both endogenous and exogenous metabolism. It is our largest visceral organ and it is involved in more than 500 different functions, playing a key role in the metabolism of carbohydrates, lipids and proteins, regulating homeostatic functions (e.g. endocrine activity and haemostasis) and participating in the systemic reaction to injury by modulating the immune response and synthesizing acute phase proteins [49,50]. Moreover, the liver is the major site for inactivation of toxins and xenobiotics, firstly removing them from blood and consequently eliminating them from the organism through bile secretion.

Chronic hepatic injury can cause fibrosis, characterised by a progressive accumulation of connective scar tissue in the liver. Hepatitis B or C, autoimmune diseases, metabolic disorders as well as alcohol abuse and drug toxicity are among the most common causes of liver fibrosis [51]. Moreover, even the physiological process of tissue ageing is associated with a mild fibrotic and inflammatory state in the liver [52]. Liver fibrosis may lead to cirrhosis and liver failure, which is one of the major cause of illness and death across industrialised countries [53], due to the impossibility of substituting the liver’s multiple activities artificially. The development of liver fibrosis is the result of a multicellular process leading to an increased deposition of ECM, mainly caused by activated hepatic stellate cells (HSCs) [51]. This extra ECM can be cross-linked via non-enzymatic glycation as well as by enzymes such as transglutaminases, lysyl oxidases and lysyl hydroxylases, resulting in a stiffening of aged or fibrotic tissue with respect to a healthy young one [49]. The progressive changes in ECM composition as fibrosis proceeds, instigates several positive-feedback pathways, which further amplifies the fibrotic process unless the causal factor is removed. No effective treatment for liver

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Introduction

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fibrosis is available yet, and extensive research is still on-going to further investigate fibrotic processes and to identify potential anti-fibrotic drugs [54]. In-vitro liver models

Several in-vitro liver models have been reported in the literature [55–57]. The first models were based on the use of hepatic sub-cellular fractions (single enzymes, micro- and super-some, cytosolic and mixed fractions) for the study of single metabolic functions [58–62]. Despite their ease of use, these models were not suited for mimicking the complex metabolism of hepatocytes and were followed by liver-derived cell lines generated from malignant tumours or obtained from transformed cells. However, many of these cell lines were not fully reliable as in-vitro liver models, due to the loss of most of their original phenotypic features. Similarly, engineered cells or systems based on reporter gene transfer were not good enough in mimicking hepatocyte functions [63]. Owing to these limitations, human primary hepatocytes are often employed in the development of experimental in-vitro human liver models [64]. However, obtaining a suitable in-vitro liver model is challenging due to the high level of specialisation and complexity of its parenchyma, and the extreme sensibility of hepatic cells to even minimal environmental changes [49]. While research has demonstrated the failure of traditional 2D in-vitro models (i.e. static cultures in monolayer of one or more cell types) in reproducing the behaviour and the physiological response of liver, 3D in-vitro models seem to give promising results. Several reports demonstrate that 3D models allow long-term cell viability and the in-vitro expression of a liver specific phenotype (e.g. albumin and urea secretion as well as enzymatic activities) [50,56]. Some of the first 3D

in-vitro liver models were based on spheroidal aggregates proposed by Landry

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enable the spatial organisation of cells [66] and, more recently, tissue-like 3D structures for toxicity studies were proposed using a hydrogel inverted colloidal crystal (ICC) scaffold [67]. Although not properly 3D, sandwich cultures, obtained culturing a monolayer of hepatocytes between two layers of extracellular matrix (typically collagen and/or Matrigel), were commonly used as platforms for analysing hepatocyte function, since the ECM layers create a barrier to mass transfer, mimicking the in-vivo situation in the liver [68,69]. Given the advantages and encouraging results of sandwich cultures, in-vitro models were further improved combining modified culture media formulations with innovative culturing techniques in well-controlled 3D microenvironments. Miranda et al. showed that a 3D culture of hepatocytes in a bioreactor system with serum supplemented medium maintain liver specific functions (such as enzymatic activities, albumin and urea secretion) better and for a longer time with respect to standard monolayer cultures (21 days against 3-4 days for 3D and 2D cultures, respectively) [70]. Hepatocytes have also been co-cultured with hepatic stellate cells (HSCs), participating in cell-cell signalling critical for liver regeneration and homeostasis in-vivo, highlighting the importance of stromal cells and culture conditions in retaining metabolic functions and better mimicking liver tissues in-vivo [29,71,72]. Thomas et al. showed that rat HSCs promote the formation of hepatic ultrastructure (including bile canaliculi, desmosomes and tight junctions) in the 3D spheroidal aggregates through ECM synthesis and direct signalling to hepatocytes [72]. Moreover, the co-culture conditions promote a faster formation of spheroids (with diameters of 100-150 μm) compared to hepatocyte monocultures, also characterised by a higher enzymatic activity (7.2 fold increase in P450-catalysed metabolism of testosterone in co-cultures, compared to hepatocyte monocultures). Guzzardi et al. studied the cross-talk between hepatocytes and endothelial cells using a

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Introduction

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multi-compartmental bioreactor [73]. Their results showed an increase in albumin and urea production in co-cultures with respect to hepatocyte monocultures. Moreover, the dynamic connected culture implemented with the multi-compartmental bioreactor promoted a higher metabolite synthesis and secretion with respect to traditional co-cultures.

Liver disease models

At present, the research of new anti-fibrotic drugs is based on in-vivo animal experiments (typically restricted to rodents), designed to mimic different causes of liver fibrosis in human, e.g. by i) chronic administration of toxic compounds such as carbon tetrachloride [74], thioacetamide [75], dimethylnitrosamide [76] and galactosamide [77], or ii) ligation of the bile duct [78]. These in-vivo models can account for possible effects of the immune and central nervous system as well as other organs on the development of liver fibrosis. However, liver functional characteristics and susceptibility to injury are highly species-specific, making these animal models inadequate, poorly informative and predictive for humans [79,80].

Few disease liver models are available to date for simulating fibrotic processes

in-vitro [81]. Although they can never completely mimic the in-vivo situation,

continuous optimisation and investigation have increased their potential during the last years. Liver cell mono-cultures are the simplest models to study the effects of pro- and anti-fibrotic compounds in-vitro [82,83] and permit the investigation of responses of single cell types. Co-cultures of different liver cell-types enable the incorporation of cell-cell interactions to study indirect effects of potential pro- or anti-fibrotic compounds on HSC activation [84,85]. Furthermore, cell culture models can be used to study the effect of ECM proteins on HSC activation and fibrogenesis in-vitro [86,87]. Besides cell

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culture models, precision-cut liver slices can better mimic the in-vivo hepatic environment, providing a multicellular in-vitro model which maintains cell-cell and cell-ECM interactions [88–90]. The first studies with precision-cut liver slices highlighted their suitability as in-vitro models for liver fibrosis, filling the gap between in-vivo and cell culture models. Moreover, sinusoid-like structures were recently engineered using micro-scale patterning, representing a promising advanced tool for the study of liver fibrosis in-vitro [50]. Combining these different cell and tissue culture models may help in studying the mechanisms responsible for multicellular fibrosis development and in investigating potential pro- or anti-fibrotic properties of different compounds, by reducing, refining and possibly replacing the use of laboratory animals. In addition, one of the main advantages of these in-vitro models is the possibility of using human cells or (healthy or fibrotic) tissues to address inter-species differences, thus improving the relevance and predictiveness for humans. However, these are just singular examples and there are in fact few and very limited instances of fibrotic and aged liver models.

Aim of the thesis

In an effort to meet the challenges described, a pathophysiological 3D in-vitro liver model was engineered during this thesis, following the classical TE approach. In particular, scaffolds for hepatic cell encapsulation and a bioreactor for the dynamic cultivation of hepatocyte-laden constructs were developed, aiming at recreating both healthy and diseased hepatic systems able to mimic ageing and fibrotic processes. First of all, since an ideal scaffold should replicate most of the properties of the native ECM, several decellularisation procedures were investigated to obtain decellularised liver matrices (dECMs) from portions of untreated hepatic tissue. Such liver dECMs were then

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Introduction

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biochemically and mechano-structurally characterised to derive ideal scaffold design specifications. Then, focusing on mechanical properties, a library of photo-crosslinkable hydrogel-based smart modular scaffolds (SMSs) was developed to encapsulate hepatic cells for obtaining 3D liver-like constructs that mimic the stiffness of the healthy tissue. The SMSs were designed such that upon subsequent exposure to lysyl oxidase, an enzyme-mediated hardening typical of fibrotic and ageing processes could be activated. The proposed hydrogel design strategy along with the adopted two-step crosslinking method allowed for a fine tuning of the healthy SMSs mechano-structural properties and of the enzymatic stiffening towards different stages of aged/diseased models. Once SMSs were developed and characterised, a bioreactor was designed to allow i) in-situ photo-crosslinking of the cell-laden SMS directly within its culture chamber, facilitating the maintenance of sterility, and ii) real-time in-situ imaging of the 3D cell culture environment during the dynamic cultivation. Finally, preliminary experiments were performed with hepatocytes by forming the cell-laden SMS directly in the bioreactor culture chamber and culturing the cells over 7 days.

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Chapter 1

Liver decellularisation and

physicochemical characterisation

Abstract

Under physiological conditions, resident cells of each tissue or organ secreted a tissue-specific extracellular matrix (ECM) that provides optimal supportive framework and micro-environmental cues for appropriate cells in-growth and differentiation. Focusing on liver, aiming to preserve most of the architecture, structure and composition of the native hepatic ECM, in this thesis several decellularisation procedures were defined and optimised to obtain well-characterised and reproducible decellularised liver matrices (dECMs). For each procedure, the “decellularised state” was identified through histological analysis and DNA quantification. Since an ideal scaffold for tissue engineering applications should replicate most of the properties of the native ECM, the liver dECMs obtained were also biochemically and mechano-structurally characterised. The results obtained were used to identify the best decellularisation procedure, thus deriving the optimal scaffold design specifications for hepatic tissue replica.

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1.1 Introduction

The liver, by way of its central role in both endogenous and exogenous metabolism, is one of the most studied organs in the human body. Hepatic tissue and its derivatives are widely used in many applications ranging from in-vitro liver models for studying hepatic drug metabolism and diseases, to tissue regeneration and incorporation into bio-artificial liver devices [30,91,92]. One of the main challenge in engineering in-vitro liver models is to maintain the native hepatic functional characteristics ex-vivo. A number of scientists suggested that this has been elusive because hepatocytes in-vitro are deprived of the multi-parametric, multi-stimuli in-vivo milieu [93]. Although the two-dimensional (2D) in-vitro liver models (i.e. cell monolayers) have made significant contributions to the biological research, the main features of the

in-vivo environment considered absent are the three-dimensional (3D)

organisation of the liver and the continuous turnover and flow of nutrients brought about by the copious blood supply to this organ. A wide variety of methods have been developed to obtain an in-vitro environment for hepatocytes more similar to the in-vivo context. Many of them are based on devising complex cocktails of culture medium, which often include inducing agents to drive hepatocytes towards the expression of P450 cytochromes [68].

The classical Tissue Engineering approach, based on the use of cells, scaffolds and bioreactors to recreate tissues and organs in-vitro for replacing diseased or missing parts, can be attractive also to realise 3D in-vitro organ models [94]. However, reproducing the 3D features of the native liver is critical since the

in-vivo environment affects cell behaviour and function through a variety of

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Introduction

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mechanical cues (e.g. oxygen concentration, substrate stiffness) [20,95]. These environmental signals are transmitted within the cell, inducing a cascade of interactive events at various levels (i.e. molecular, subcellular and cellular levels) [96,97]. Beyond cell-cell communications, cell-ECM interactions are of primary importance since they play a fundamental role in hepatocyte growth [98], liver organ development [99], tissue regeneration [100], wound healing [100] and liver diseases [101]. Under physiological conditions, resident cells of each tissue and organ secreted an optimal tissue-specific ECM including collagen, fibronectin, laminin, glycosaminoglycans (GAGs), proteoglycans (PGs) and non-soluble growth factors. The ECM is in a state of dynamic reciprocity with these cells in response to changes in the microenvironment and has been shown to provide cues that affect cell migration, proliferation and differentiation [102,103]. The 3D ultrastructure, surface topology and composition of the native ECM all contribute to these effects, therefore the consensus is that an optimal scaffold for tissue engineering should mimic these physicochemical features to provide cells with an appropriate supportive framework and micro-environment. Moreover, to be widely employed for high or medium throughput testing, a scaffold should also be well characterised, reproducible, easy to fabricate, sterilise and store [104].

Biological scaffolds derived from decellularised tissues and organs have been successfully used for a myriad of applications in reconstructive surgery and regenerative medicine [105,106]. Since the presence of residual cellular material attenuates or even fully negates the advantages of using biological scaffolds [107], tissue decellularisation procedures to achieve complete cell removal are critical [106]. A variety of methods have been proposed to remove cells from whole organs or organ portions collected from small animals such as

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mice, to larger species such as pigs. Many different factors (e.g. tissue’s density, cellularity, lipid content and thickness) have to be taken into account while choosing the most effective decellularisation agents. These decellularisation agents can be generally classified in: i) chemical (e.g. ionic, non-ionic and zwitterionic detergents, acids and bases, hypotonic and hypertonic solutions), ii) biological (e.g. enzymes) and iii) physical (e.g. temperature, force and pressure). The optimal techniques to apply decellularisation agents are again dependent on tissue characteristics (e.g. thickness and density) as well as the agents being used and the desired application of the decellularised tissue. Most of the proposed methods for removing cells from whole liver or portions of hepatic tissue involve washing and saponification phases to respectively remove blood and debris, and hydrolyse cell membranes. In general, the outcome of a detergent-based decellularisation procedure for a given tissue is strongly dependent on the delivering method of decellularisation agents and the exposure time, which increases removal of cell residues, but also ECM disruption. Residual chemicals must be flushed out from ECM after decellularisation, in particular detergents such as sodium dodecyl sulphate (SDS) that penetrate into thick or dense tissues. In fact, such agents can be cytotoxic for subsequent cell cultures even in low concentrations, inhibiting or completely negating the beneficial properties of the decellularised scaffold [108]. Among the several decellularisation methods reported in the literature, whole organ perfusion, based on perfusing detergents through native organ vasculature, has been recently used by Uygun et al. [109] and Shupe et al. [110] to remove cellular material from whole livers. By perfusing SDS and Triton X-100 through the hepatic portal vein, both studies showed a complete cell removal (confirmed by histological results and DNA quantification), while microvasculature and ECM ultrastructure and constituents (e.g. collagen I and

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Introduction

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IV, fibronectin and laminin) were preserved in the decellularised matrices. Moreover, Uygun et al. re-seeded hepatocytes into decellularised livers by portal vein perfusion, assessing the metabolic activity of the recellularised liver graft through albumin production, urea secretion and total bile acid synthesis. The cumulative production of these metabolites was found to be similar to that of static sandwich culture controls, demonstrating the usability of the perfusion-culture system as an in-vitro model. Besides whole organ perfusion [109,111,112], common decellularisation techniques include pressure gradients [113–115] and immersion and agitation approaches [108,116,117]. Following an immersion and agitation approach, Lang et al. described a study to identify the most suitable combination of washing and saponification cocktail to decellularise thin slices of pig liver [116]. The authors optimised different decellularisation/oxidation procedures based on Triton X-100, ammonium hydroxide, phosphate buffered saline (PBS) and sodium chloride, eventually coupled with a peracetic acid (PAA) treatment. They report 93 % removal of cellular components from porcine liver tissue and preservation of the key molecular components in the ECM, including collagen I, III and IV, proteoglycans, glycosaminoglycans, fibronectin, elastin and laminin. Despite the recent explosion of interest, there is little consensus on the optimum method of decellularising hepatic tissue able to preserve the micro-architecture and protein content of the matrix as far as possible. Indeed, it has been recognised that aggressive physical and chemical treatments may alter the protein structure, content and porous architecture of the matrices.

In this context, the first part of this thesis aimed at developing a method for the production of well-characterised and reproducible liver matrices, which best preserve the structure and composition of the native ECM. Focusing on an

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immersion and agitation approach, several decellularisation procedures were investigated to obtain liver dECMs from discs of porcine hepatic tissue. Liver dECMs were then characterised using biochemical, histological, mechanical and structural analyses to assess the effect of different detergents and treatment duration on matrix physicochemical properties. The results were then used to identify the best procedure which ensures a complete cell removal while preserving most of the native ECM features. The “aggressiveness” of different decellularisation procedures was evaluated comparing the properties of resultant matrices with those of untreated liver. Particular attention was dedicated to the testing and analysis framework in order to enable meaningful comparisons between different decellularisation methods and controls, characterising both decellularised and untreated samples in the same reproducible initial state (i.e. the equilibrium swollen state). A number of different livers were sampled to determine whether individual variations amongst pigs could contribute to the scatter in the structural and biochemical parameters. Liver dECMs with highly conserved intra-lobular micro-structure and protein content (using the sample equilibrium swollen state as a reference) were obtained in a consistent and reproducible manner. Finally, several sterilisation procedures were investigated and cytotoxicity tests performed to obtain sterile liver dECMs as scaffolds for 3D cell cultures or for further applications such as to produce tissue derivatives (e.g. solubilised or powdered form of decellularised hepatic ECM).

Overall, the results obtained contribute to define the ideal design criteria for the fabrication of hepatic ECM-mimicking scaffolds for engineering 3D in-vitro liver models. Given the critical role of matrix stiffness in regulating cell response, the attention was focused on mechanical properties of both

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Experimental section

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decellularised and untreated hepatic tissue. However, due to the challenges in testing very soft and highly hydrated biological tissues, the adopted testing and analysis framework will be discussed in detail in the next chapter of this thesis (Chapter 2) along with the results obtained.

1.2 Experimental section

1.2.1 Hepatic tissue collection

Fresh livers were collected from one year old healthy pigs as a slaughter by-product. Pig liver is composed of five lobes (right lateral, right medial, left medial, left lateral and caudate lobe) wrapped in a tough fibrous capsule (i.e. the Glisson’s capsule) that ensures structural integrity [118]. Individual lobes were dissected from each liver, apart from the small caudate one. Some fresh samples were isolated and immediately processed for histology or tested for mechanical properties to assess eventual intra- and inter-pig variability (Chapter 2). The remnant portions of liver lobes were stored at -20 °C until use. Frozen livers were thawed at 4 °C overnight and then punched with a 14 mm diameter tool to obtain regular cylinders which were subsequently cut in 3 mm thick liver discs using a custom slicer frame and a microtome blade. The Glisson’s capsule was not present in these samples and particular attention was dedicated to avoid macroscopic vasculature. Liver discs were then stored at - 20 °C until use.

1.2.2 Liver decellularisation procedures

Focusing on an immersion and agitation approach, several decellularisation protocols were investigated varying chemical detergent and treatment duration (Table 1.1).

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Table 1.1: Decellularisation protocols investigated to obtain liver dECMs.

Percentage values are referred to the weight/volume ratios (w/v) of detergent solutions prepared in deionised water (dH2O). The final washing day for

detergent-free protocols involved only PBS 1x, since the contribution of 0.1 % w/v Triton X-100 to cell removal is very poor and not significant compared to that of 1 % w/v Triton X-100 or 0.1 % w/v SDS used in the other protocols.

Family Protocol ID Day 1 Day 2 Day 3 Day 4 Day 5

Ionic I3 PBS 1x SDS 0.1 % ½ d Triton X-100 0.1 % + ½ d PBS 1x I4 PBS 1x SDS 0.1 % SDS 0.1% ½ d Triton X-100 0.1 % + ½ d PBS 1x I5 PBS 1x SDS 0.1 % SDS 0.1 % SDS 0.1 % ½ d Triton X-100 0.1 % + ½ d PBS 1x Non-ionic NI3 PBS 1x Triton X-100 1 % ½ d Triton X-100 0.1 % + ½ d PBS 1x NI4 PBS 1x Triton X-100 1 % Triton X-100 1 % ½ d Triton X-100 0.1 % + ½ d PBS 1x NI5 PBS 1x Triton X-100 1 % Triton X-100 1 % Triton X-100 1 % ½ d Triton X-100 0.1 % + ½ d PBS 1x Detergent free DF3 PBS 1x PBS 1x PBS 1x DF4 PBS 1x PBS 1x PBS 1x PBS 1x DF5 PBS 1x PBS 1x PBS 1x PBS 1x PBS 1x Not treated (control)

FF Fresh liver frozen at -20 °C, then thawed at 4 °C overnight

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In particular 20 liver discs (about 10 g of samples) were placed into each bottle, then 200 mL of decellularisation solution were added. Filled bottles were placed on an orbital shaker (SO3, Stuart Scientific, Stone, UK) at 200 rpm in a cold room at 4 °C, changing the decellularisation solution twice a day for at least 3 days.

The investigated decellularisation procedures can be grouped into three major families depending on the nature of the chemical detergent used: detergent-free

(DF), ionic (I) and non-ionic (NI). The detergent-free family was provided as a

control to decouple the decellularising effect related to mechanical agitation from that due to chemical agents (i.e. detergents). Two different chemical detergents, i.e. Triton X-100 and sodium dodecyl sulphate (SDS), were chosen on the basis of their different action modes and reported effects on ECM [119]. Specifically, Triton X-100 is a non-ionic detergent that disrupts DNA-protein interactions, lipid-lipid, lipid-protein and protein-protein interactions. SDS is an ionic detergent (negatively charged, or anionic) that solubilises cytoplasmic and nuclear cellular membranes, better removing cell nuclei from dense tissues and organs than Triton X-100, but tends to denature proteins, disrupting the ultrastructure [120] and eliminating key growth factors [121]. The rinsing day at the beginning of each protocol served mainly to bleed liver discs, while the final washing day is critical to remove detergent residues from decellularised samples. The capability of non-ionic Triton X-100 to form micelles with anionic SDS [122] was used to enhance residual SDS removal from the ionic protocols during the last washing day.

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1.2.3 Decellularisation assessment

Cell removal from liver discs obtained with each of the nine protocols was assessed first histologically with haematoxylin and eosin (H&E) staining and then further investigated by DNA quantification (Section 1.2.5). Untreated (control) and decellularised liver samples were formalin fixed, paraffin embedded and cut into 5 µm sections. Histological sections were stained with H&E and examined using an Olympus IX81 microscope (Olympus Italia, Milan, IT).

1.2.4 Swelling experiments

As anticipated in the introduction, the sample’s equilibrium swollen state was taken as a reproducible testing condition and reference to enable meaningful comparisons between matrices obtained with different decellularisation procedures and untreated frozen and thawed liver samples, henceforth termed fresh-frozen (FF), used as controls. FF and decellularised liver discs were freeze-dried at - 50 °C, 0.45 mbar for 48 h to determine their dry weight, 𝑊𝑑.

Then they were swollen in PBS 1x at room temperature and weighed every 12 hours until obtaining a stable weight (i.e. the equilibrium swollen weight, 𝑊𝑒𝑞).

The equilibrium mass swelling ratio was calculated as 𝑄𝑒𝑞 = 𝑊𝑒𝑞/𝑊𝑑. All

measurements were performed in triplicate.

1.2.5 Total DNA quantification

The total DNA content for fresh-frozen and all decellularised liver samples was estimated using the NanoDrop system (NanoDrop Technologies, Wilmington, DE) and expressed as µg of DNA per mg of equilibrium swollen sample. Total DNA was extracted using the DNeasy Blood & Tissue kit (Qiagen, Valencia, CA) according to the manufacturer's instructions. In particular, 25 mg of

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equilibrium swollen tissue were used for the DNA quantification of FF and DF samples, while 25 mg of freeze-dried tissue were used in case of ionic and

non-ionic dECMs to concentrate any residual DNA. Liver samples were placed into

1.5 mL tubes, then the lysing solution was added and the DNA content estimated with the NanoDrop system. In case of ionic and non-ionic dECMs, the total DNA per mg of equilibrium swollen sample was obtained multiplying the measured DNA content by the inverse of the equilibrium mass swelling ratio (1/𝑄𝑒𝑞).

1.2.6 Biochemical characterisation

The “aggressiveness” of decellularisation procedures was biochemically investigated through total protein content (TPC) quantification and Western Blot analysis, comparing liver dECMs results to those of FF untreated samples. The TPC was determined in triplicate using the Bio-Rad Protein Assay (Bio-Rad Laboratories Inc, Hercules, CA). Liver samples were first equilibrium swollen in PBS 1x to determine the equilibrium wet weight (𝑊𝑒𝑞) and then

solubilised in 1 M NaOH solution at 60 °C for 2 h, using 10 mL of sodium hydroxide solution per gram of wet sample (i.e. alkaline solubilisation). Assuming a liver density equal to that of water (i.e. 1 g/cm3_{), the resultant}

NaOH concentration can be calculated as 1/11 = 0.91 M. To avoid any interference with the Bio-Rad assay, solubilised samples were diluted 1:10 v/v in dH20, obtaining solutions with a sodium hydroxide concentration compatible

with the maximum allowed (0.1 M), as reported in the manufacturer’s guidelines [123]. Therefore, the minimum wet sample dilution was 1/110, while further dilutions with 0.091 M NaOH were necessary in case of samples with very high protein concentrations (e.g. untreated cellularised controls). The TPC

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calibration curve was established using bovine serum albumin (BSA, Sigma-Aldrich, Milan, IT) standard solutions prepared as described for liver samples to account for any eventual effect related to the adopted alkaline solubilisation protocol. Briefly, BSA powder was solubilised in 1 M NaOH at 60 °C for 2 h and then diluted 1/11 in deionised water (dH2O) to obtain 0.091 M NaOH

solutions with known protein concentrations. The absorbance was read at 595 nm using a FLUOstar Omega spectrophotometer (BMG Labtech GmbH, Offenburg, Germany) and expressed as mg of protein per gram of wet sample, considering sample dilution and 𝑊𝑒𝑞, thus obtaining comparable data between

decellularised and/or fresh-frozen liver samples.

The presence of specific liver ECM proteins (i.e. laminin, fibronectin and collagen IV) in decellularised matrices was selectively analysed using Western Blot. On the basis of decellularisation outcomes and TPC results (Sections 1.3.1 and 1.3.4), Western blot analysis was performed in triplicate only for matrices obtained with protocol I3, NI3 and DF3, respectively representative of ionic,

non-ionic and detergent-free families, and compared to that of FF samples

(used as control). All samples were homogenised on ice using an Ultra-Turrax®

T25 (IKA GmbH, Germany) in a 10 mM Tris–HCl buffer (pH 7.8) containing protease inhibitors (4 μg/mL leupeptin, 1 μg/mL aprotinin and 1 mM phenylmethylsulfonyl fluoride) and then passed through a 22 gauge needle. Total protein content was determined in triplicate using the Bio-Rad Protein Assay (Bio-Rad Laboratories Inc, Hercules, CA). To enable meaningful comparisons between protocols, the same total protein quantity (10 μg) was loaded into each gel lane. In order to do that, 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

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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), subsequently stained with Ponceau S to verify loading and transfer efficiency. Non-specific 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 60 min at room temperature. Membranes were then incubated overnight at room temperature with i) 1:1000 dilution of goat polyclonal antibody raised against porcine fibronectin, ii) 1:1000 dilution of goat polyclonal antibody against porcine laminin or iii) 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. 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 enhanced chemiluminescence (ECL) detection system (Roche, Basel, Switzerland).

1.2.7 Architectural analysis

The architectural analysis of decellularised liver matrices is fundamental to characterise their pore size, porosity and permeability. Aiming to better investigate liver dECM features, further histological analysis was performed on 10 μm slices stained with silver, Mallory’s trichrome and Alcian blue/PAS (thinner sections, e.g. 5 μm, were lacking in contrast). Moreover, to control the preservation of 3D hepatic micro-structure, decellularised samples were also analysed with μ-CT scanning and confocal imaging. The 3D architecture of

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freeze-dried matrices was reconstructed using a SkyScan 1174 μ-CT scan (Skyscan, Aartselaar, Belgium) with a resolution of 6.5 μm/pixel, 180° rotation. Conversely, confocal imaging was performed on eosin-stained equilibrium swollen matrices using a Nikon A1 confocal microscope (Nikon, Tokyo, Japan): the scanning parameters were adjusted to maintain a pixel size less than 0.2 μm.

1.2.8 Liver dECMs sterilisation and cytotoxicity tests

Aiming at obtaining sterile liver dECMs usable as scaffolds for 3D cell cultures or further processable to produce tissue derivatives, protocol NI3 matrices were freeze-dried and then treated with four different sterilisation procedures: i) overnight exposure to chloroform (Carlo Erba, Milan, Italy) vapour, ii) overnight exposure to chloroform vapour followed by gas plasma sterilisation (Sterrad 100S, Advanced Sterilization Products, Irvine, CA), iii) gas plasma sterilisation only or iv) 2 h soaking in 0.1 % peracetic acid (Sigma-Aldrich, Milan, IT). Chloroform sterilisation was performed pouring 20 mL of solvent on the bottom of a 5 L desiccator and placing NI3 matrices on the desiccator ceramic support plate, about 3 cm above the liquid. Then the desiccator lid was closed and samples exposed to chloroform vapour overnight at room temperature.

Cytotoxicity experiments were then performed in triplicate placing so treated matrices in contact with HepG2 cell cultures. Briefly, HepG2 cells from American Type Culture Collection (ATCC), were cultured in Eagle’s minimal essential medium (EMEM, Sigma-Aldrich, Milan, IT) supplemented with 10 % fetal bovine serum (FBS, Sigma-Aldrich, Milan, IT), 1 % penicillin, 1 % streptomycin and 1 % L-glutamine (Invitrogen, Milan, IT). 50.000 cells per

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well were seeded in a 24 well plate with 2 mL of medium and cultured for 48 h allowing them to adhere and proliferate. Protocol NI3 liver dECMs treated with different sterilisation procedures were copiously rinsed in sterile PBS and placed in the cell-seeded wells. Cell viability was then assessed at 24 h and 48 h using CellTiter Blue (Promega, Madison, WI) and compared to that of HepG2 cells cultured on tissue culture plates (used as controls).

1.2.9 Statistical analysis

All experiments were carried out at least in triplicate using samples from different pig livers. Comparisons between n groups of data (e.g. total protein content, equilibrium mass swelling ratio) referring to one factor only (e.g. different decellularisation protocols) were performed using one-way ANOVA followed by Tukey’s post hoc multiple comparison test. Two-way ANOVA followed by Tukey’s post hoc multiple comparison test was used to analyse the viability of cells placed in contact with liver dECMs sterilised with different methods (1st_{factor of variability) at different culture times (2}nd_factor).

Differences were considered significant at p < 0.05. Statistical analysis was performed using OriginPro (OriginLab, Northampton, MA).

1.3 Results

1.3.1 Histological analysis

Haematoxylin and eosin (H&E) micrographs for each of the investigated protocols (Table 1.1) are shown in Figure 1.1. No cells were detected in either

ionic (I) or non-ionic (NI) dECMs, but several cells were observed in all the detergent-free (DF) protocols. Notably, there is a gradual collapse in the lobule

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independent of the nature of the detergent. The use of SDS clearly promotes a gradual removal of the intra-lobular matrix, extending from the centre of the lobule outwards.

Figure 1.1: H&E micrographs for liver dECMs obtained with all tested

decellularisation protocols as a function of treatment duration and nature of detergent used (protocol family). Scale bars 200 μm.

Focusing on 3 day long decellularisation protocols, additional micrographs at higher magnification are provided in Figure 1.2, better showing the inefficacy of DF3 protocol at cell removal, the aggressiveness of the I3 and the intra-lobular ultrastructure preservation of protocol NI3 dECMs.

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Figure 1.2: H&E micrographs of A) DF3 liver dECM (20x) with several cells

indicated by the arrows, B) I3 liver dECM (10x) with arrows showing largely vacant lobules and C) NI3 liver dECM (10x). D) Silver stained NI3 liver dECM (10x), better showing the preserved hepatic intra-lobular ultrastructure than H&E staining, with pore sizes of cellular dimensions (indicated by arrows). Scale bars 50 μm.

1.3.2 Swelling behaviour

Samples equilibrium mass swelling ratios (𝑄𝑒𝑞) were not found to be

significantly affected by the duration of the decellularisation procedures (3 to 5 days), but only dependent on the protocol family, as shown in Figure 1.3. The

ionic samples were characterised by an equilibrium swelling ratio of 10.3 ± 2.1,

which was significantly higher than that of non-ionic dECMs, equal to 6.7 ± 0.5 (p = 0.018). Moreover, the ionic samples took a shorter time to be

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equilibrium swollen (24 h) than the non-ionic ones (36 h), starting from the freeze-dried state.

Figure 1.3: Equilibrium swelling ratios (𝑄𝑒𝑞) for different sample families.

Significant differences are marked with an asterisk (* = p < 0.05).

These differences in swelling behaviour can be related to the ECM-rich intra-lobular network of non-ionic liver dECMs that hindered PBS absorption within the matrix, resulting in a lower equilibrium mass swelling ratio, reached after a longer time with respect to ionic samples. The latter were indeed characterised by a more open porous network composed principally of inter-lobular connective tissue (Section 1.3.1), which presumably allowed for a greater and more rapid penetration of water. The 𝑄𝑒𝑞 of FF samples was found to be equal

to 1.9 ± 0.4, similar to that of DF matrices, equal to 2.3 ± 0.4 (p = 0.96). Both FF and DF samples took 48 h to reach their equilibrium swollen state, characterised by a significantly lower 𝑄𝑒𝑞 than that of ionic (p = 7.19∙10-5 for

FF vs. I, and p = 1.03∙10-4_{for DF vs. I) and non-ionic samples (p = 3.35∙10}-3

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structural differences between samples as seen from histology, DNA and total protein content analyses. In particular, FF and DF samples were characterised by a high cell density and hence a tighter network with fewer pores for water to penetrate with respect to the cell-free I and NI liver matrices.

1.3.3 Total DNA content

The total DNA content for all samples is shown in Figure 1.4, confirming both histological and swelling results. The detergent-free (DF) protocols are clearly not sufficient to remove cells from hepatic tissue, demonstrating the need to use chemical detergents along with mechanical agitation. Since differences between ionic and non-ionic procedures were insignificant, either reagent can be used for complete cell removal, which was reached after 3 days.

Figure 1.4: Total DNA content of fresh-frozen and decellularised liver samples,

expressed as μg of DNA per mg of equilibrium swollen sample. Different letters indicate significant differences between samples (p < 0.05), whereas the same letter means non significant differences.

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1.3.4 Biochemical characterisation

Total protein contents (TPCs) for fresh-frozen and decellularised liver samples are summarised in Figure 1.5. All TPC data are expressed as mg of proteins per gram of equilibrium swollen sample, apart from FF-NS (i.e. fresh-frozen not-swollen sample) which was not equilibrium not-swollen, but processed as taken to account for eventual protein lost in the swelling medium during the swelling process. As expected, the TPC for FF liver sample (65 ± 7 mg protein/g wet sample) was found to be significantly lower than that of FF-NS (185 ± 7 mg protein/g wet sample), mainly due to the elimination of blood proteins during swelling. The TPC of FF-NS is in agreement with published data [124], however, to be consistent with results obtained for dECMs, the FF value was chosen as control, enabling meaningful comparisons between TPCs.

Figure 1.5: Total protein content (TPC) of fresh-frozen and decellularised liver

samples. Data are expressed as mg of protein per gram of equilibrium swollen sample, except for the fresh tissue (FF-NS) which was processed as it was. Different letters indicate significant differences between samples (p < 0.05).

Smart modular scaffolds (SMSs) for the realisation of 3D in-vitro organ models

UNIVERSITY OF PISA

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Smart modular scaffolds (SMSs) for the

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Author:

Giorgio M

Table of Contents

Abstract

Acknowledgements

Introduction

Chapter 1

Liver decellularisation and

physicochemical characterisation

Abstract

1.1 Introduction

1.2 Experimental section

1.2.1 Hepatic tissue collection

1.2.2 Liver decellularisation procedures

1.2.3 Decellularisation assessment

1.2.4 Swelling experiments

1.2.5 Total DNA quantification

1.2.6 Biochemical characterisation

1.2.7 Architectural analysis

1.2.8 Liver dECMs sterilisation and cytotoxicity tests

1.2.9 Statistical analysis

1.3 Results

1.3.1 Histological analysis

1.3.2 Swelling behaviour

1.3.3 Total DNA content

1.3.4 Biochemical characterisation