Ex vivo fibrosis research: 5 mm closer to human studies Bigaeva, Emiliia

Ex vivo fibrosis research: 5 mm closer to human studies Bigaeva, Emiliia

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5 mm closer to human studies

Emilia Bigaeva

2019

The research was carried out at the Groningen Research Institute of Pharmacy, department of Pharmaceutical Technology and Biopharmacy, in collaboration with University Medical Center Groningen (The Netherlands), the Institute of Translational Immunology, Johannes Gutenberg University Mainz (Germany) and Boehringer Ingelheim Pharma GmbH & Co. KG (Germany).

Printing of this thesis was financially supported by the University Library and the Graduate School of Science and Engineering of the University of Groningen.

PARANYMPHS

Emilia Gore

Susana Figueroa-Lozano

Ex vivo fibrosis research 5 mm closer to human studies

Layout & cover design: Emilia Bigaeva

Printing: Ipskamp Printing, Enschede

ISBN (printed): 978-94-034-1832-2 ISBN (digital): 978-94-034-1831-5

© Copyright 2019, Emilia Bigaeva

All rights are reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic or mechanical, without permission of the author.

Ex vivo fibrosis research:

5 mm closer to human studies

PhD Thesis

to obtain the degree of PhD at the University of Groningen

on the authority of the Rector Magnificus prof. E. Sterken

and in accordance with the decision by the College of Deans.

This thesis will be defended in public on Friday 12 July 2019 at 16.15 hours

by

Emilia Bigaeva

born on 23 May 1990

in Saint-Petersburg, Russia

Co-supervisors

Dr. M. Boersema Dr. H.A.M. Mutsaers

Assessment Committee

Prof. dr. J.L. Hillebrands Prof. dr. K.N. Faber Prof. dr. W. Jiménez

Dedicated to the ones I love the most: my family.

With all of you by my side, I am complete.

In memory of my father

Валерий Рафаэльевич Бигаев

23.04.1965 – 04.08.2018

PART I CHARACTERIZATION OF THE PCTS MODEL

CHAPTER 2 33 Exploring organ-specific features of fibrogenesis using murine precision-cut tissue slices

CHAPTER 3 65 Next generation sequencing reveals in-depth features of murine and human precision-cut tissue slices

CHAPTER 4 105 Ex vivo human fibrosis model: characterization of healthy and diseased precision-cut tissue slices by next generation sequencing

PART II PCTS AS AN EX VIVO SCREENING PLATFORM FOR ANTIFIBROTIC THERAPIES

CHAPTER 5 145 Inhibition of tyrosine kinase receptor signalling attenuates fibrogenesis in an ex vivo model of human renal fibrosis

CHAPTER 6 175 Predictive value of precision-cut kidney slices as an ex vivo screening platform of antifibrotic therapies for human renal fibrosis

PART III OPTIMIZATION OF THE PCTS CULTURE CONDITIONS CHAPTER 7 223 Growth factors of stem cell niche extend the life-

span of precision-cut intestinal slices in culture:

a proof-of-concept study

CHAPTER 8 247 General discussion and perspectives CHAPTER 9 269 Summary (English, Dutch and Russian)

CHAPTER 10 284 Abbreviations 288 Author affiliations

291 Curriculum vitae 293 List of publications 296 Acknowledgements

CONTENT S

OF THIS THESIS

1

1

“Science, my lad, is made up of mistakes, but they are mistakes which it is useful to make,

because they lead little by little to the truth”.

Jules Verne, Journey to the Center of the Earth, 1864

ORGAN FIBROSIS AT A GLANCE

The evolutionary conserved ability to repair tissues is essential for the survival of an organism [1].

As a fundamental step of the wound healing process, scar formation represents a reaction that aims to limit organ damage and maintain architectural integrity of the remnant tissue [2]. However, when normal wound healing fails, usually under sustained or repetitive injury, scarring becomes a pathological condition, known as fibrosis, that leads to the excessive accumulation of extracellular matrix (ECM) components at the site of injury (Figure 1). Fibrosis disrupts the normal tissue architecture and ultimately results in a loss of organ function [3,4]. Fibrotic diseases account for 45%

of deaths worldwide [5]. Fibrosis is a dynamic, highly integrated and immensely complex process that is triggered and orchestrated by a cornucopia of different cells and signalling molecules [6,7]. It can affect virtually any human organ, including the lung, skin, heart, kidney, liver and intestine. Considering that different types of tissues and organs share common mechanisms of fibrosis and a large variety of pathophysiologically distinct diseases converge into this single process, fibrosis represents an attractive therapeutic target [8,9]. Therefore, understanding the mechanism of fibrogenesis is crucial for the development of therapeutic options that allow us to prevent, stop or revert the progression of fibrosis and improve organ function in numerous diseases.

Injury and inflammation

Mechanical trauma, ischemic injury, infections, toxins, or autoimmune reactions, can all cause tissue damage. Irrespective of the initiating insult and affected organ, tissue injury induces the progressive loss of parenchyma (i.e., hepatocytes in the liver, nephrons in kidney) and activation of local inflammatory processes [10]. Upon injury, damaged epithelial and endothelial cells release various danger signals – damage-associated molecular patterns (DAMPs) – and chemokines that recruit inflammatory cells to the site of injury [11,12]. As a result, lymphocytes, monocytes, macrophages and dendritic cells migrate to the area to facilitate tissue repair by removing pathogens and/or apoptotic cells. Therefore, acute inflammation, as a response to a disrupted epithelial surface, is a physiological part of normal wound healing [13]. However, persistent inflammation is a powerful stimulus that triggers fibrosis [14].

Once inflammatory cells have infiltrated the wound, they become activated and start to produce a large variety of cytokines and chemokines, including interleukins IL-1β, IL-6 and tumor necrosis factor TNFα, as well as other profibrotic molecules, such as reactive oxygen species (ROS) [15,16]. These factors, in turn, promote fibrogenesis by triggering the proliferation and activation of fibroblasts that continuously sustain ECM deposition.

Figure 1. Schematic overview of wound healing and fibrosis. Epithelial and/or endothelial damage caused by various insults triggers complex interconnected tissue repair programs in an attempt to restore homeostasis. Early responses comprise acute inflammation and activation of innate immune cells, such as resident macrophages, monocytes and neutrophils. Released inflammatory mediators (cytokines, chemokines) and growth factors promote fibrogenesis by activating cellular progenitors into myofibroblasts that orchestrate the production of ECM components. Failure to eliminate factors causing the injury perpetuates wound healing and inflammation, ultimately resulting in fibrosis.

TNFα, tumor necrosis factor alpha; IL-1β, interleukin-1 beta; TGFβ, transforming growth factor beta; PDGF, platelet- derived growth factor; ECM, extracellular matrix.

Myofibroblasts and tissue remodelling

Activation of fibroblasts is a central driving force of fibrogenesis and tissue remodelling [17,18]. In healthy tissues, fibroblasts are primarily involved in homeostatic ECM turnover [19]. During both physiological and pathological wound healing, fibroblasts differentiate into myofibroblasts that produce large amounts of ECM components, including fibrillar collagens (mainly collagen type I and III), cellular fibronectin, laminin, elastin, glycoproteins and proteoglycans [7,20]. A key feature of myofibroblasts is their contractile function, which is aided by the expression of a-smooth muscle actin (a-SMA) [21,22]. Myofibroblasts not only deposit matrix, but also directly contribute to tissue damage by producing an array of inflammatory cytokines and growth factors, and generating ROS [23,24]. The latter can further amplify inflammation and fibrosis [25,26].

Interestingly, the myofibroblast is not a specific cell type, but rather it represents a functional cell

fate that develops in response to injury [4,27]. The cellular origin of myofibroblasts has been a major

focus of fibrosis research over the past 20 years, yet, it remains a matter of an ongoing debate. The

main hurdle for elucidating the origin of myofibroblasts is the lack of specific cell markers, and inter-

organ and inter-species differences in existing markers [24]. Figure 2 illustrates possible cellular

progenitors of myofibroblasts. Until very recently, it was widely accepted that injured epithelial cells 1

served as the primary source of myofibroblasts [28,29]. In experimental models of organ fibrosis, fully mature epithelial and endothelial cells acquire a mesenchymal-like phenotype and function [30,31].

This phenomenon of sequential loss of epithelial/endothelial markers and de novo acquisition of mesenchymal markers is termed epithelial-to-mesenchymal (EMT) or endothelial-to-mesenchymal transition (EndoMT). However, state-of-the-art lineage tracing studies have provided strong evidence that mesenchymal cells – resident interstitial fibroblasts and pericytes – are the major, if not the only, source of ECM-producing myofibroblasts in multiple organs [32–40]. The exact definition of fibroblasts is somewhat elusive: while it is well recognized that fibroblasts are a heterogeneous cell population that can differ even within an organ, some argue that historically defined fibroblast is actually not a cell type, but a general term for heterogeneous subsets of mesenchymal cells [8,40,41].

Other potential myofibroblast progenitors include circulating bone marrow-derived fibrocytes that migrate to the area of tissue injury, and resident mesenchymal stem cells (MSCs) that can be found in perivascular niches in human tissues [42–44].

Figure 2. Proposed origin of myofibroblasts in fibrosis. Activated myofibroblasts can originate from mesenchymal progenitors, such as tissue-resident fibroblasts and pericytes, that undergo differentiation directly under the influence of the profibrotic environment. Mesenchymal cells derived from circulating fibrocytes or bone marrow-derived stem cells contribute to the pool of activated myofibroblasts. Epithelial and endothelial cells also can become myofibroblasts once they acquire a mesenchymal-like phenotype through epithelial-to-mesenchymal (EMT) or endothelial-to- mesenchymal transition (EndoMT), respectively. Damaged epithelial and endothelial cells release various danger signals and proinflammatory mediators that attract immune cells to the site of injury. Sustained inflammatory environment promotes differentiation of progenitor cells into myofibroblasts that produce excessive amounts of ECM.

Not only myofibroblasts, but also deposited ECM actively contributes to fibrogenesis. As fibrosis progresses, ECM proteins, particularly fibrillar collagens, undergo extensive enzymatic cross-linking by collagen oxidases, including lysyl oxidase-like (LOXL) family members and transglutaminases [20]. This biochemical modification increases rigidity of the fibrotic matrix, rendering it resistant to proteolysis and degradation [45]. Successful matrix degradation is a critical component of the resolution phase of normal wound healing, whereas organ fibrosis is characterized by an aberrant ECM turnover [46,47]. In the latter case, an imbalance in the expression and activity of ECM remodelling enzymes, such as matrix metalloproteinases (MMPs) and tissue inhibitors of metalloproteinases (TIMPs), favours pathological deposition of ECM. MMPs degrade various matrix components, cell surface receptors, cytokines and growth factors [48]. For instance, collagenases MMP-1, MMP13 and MMP-14 as well as gelatinases MMP-2 and MMP-9 are able to process intact or denatured collagen fibrils, respectively [49]. Generally, increased levels of MMPs are associated with fibrosis progression, and have been implicated in other diseases, such as cancer, atherosclerosis and osteoarthritis; however, their multifunctional nature makes it challenging to precisely define the role of MMPs in these disease states [49–51].

Failure to degrade accumulated ECM leads to increased tissue stiffness, which in turn stimulates fibrogenesis by activating mesenchymal cells and enhancing a variety of fibrosis-associated signalling pathways via a positive feedback loop [29,47,52]. In other words, excessive ECM deposition and increased tissue stiffness, even in the absence of ongoing injury or inflammation, are capable of sustaining progressive fibrosis via a vicious circle [53]. Similarly, Herrera et al. [54] suggest that at some point during the course of fibrosis progression, mesenchymal cells acquire “autonomy”, so that they no longer respond to ECM cues that would normally terminate the fibrotic response. Therefore, at a certain, as of yet undefined, stage in the fibrotic process, disease progression becomes irreversible and self-sustainable.

Molecular pathways in fibrogenesis

Fibrosis is not just defined by the matrix and cells that produce matrix, but also by the core molecular

pathways that drive fibrogenesis. These pathways involve a large variety of growth factors – proteins

that stimulate cell growth and proliferation – that are secreted by epithelial/endothelial cells, immune

cells and myofibroblast progenitors, and are able to orchestrate cellular responses [55]. Among these

growth factors, transforming growth factor beta (TGFβ) dominates the pathobiology of organ fibrosis

and is therefore considered as the central profibrotic mediator [56,57]. TGFβ strongly promotes the

production of ECM components, including collagen I and III, and fibronectin, and drives differentiation

of myofibroblast progenitors [58]. Sustained TGFβ activation contributes to deregulated matrix

turnover by inhibiting the synthesis of MMPs and increasing the synthesis of TIMPs and plasminogen

activator inhibitor 1 (PAI-1). On the other hand, TGFβ exerts immunoregulatory properties and acts as

a negative regulator of pro-inflammatory responses in various organs, reflecting its pleiotropic nature

[59]. Multifunctionality of TGFβ is further demonstrated by its critical involvement in the regulation of

embryogenesis, carcinogenesis, cell proliferation and migration, among others [10].

TGFβ exists as an inert complex covalently bound to the ECM, and is released/activated upon injury 1

via interaction with integrins – mechanosensing cell-surface receptors that mediate cell-adhesion [60]. This mechanism is directly correlated to the contraction of myofibroblasts and stiffness of the ECM, which represents a reservoir of latent TGFβ in the tissue [27,61]. While controlled amounts of TGFβ play an essential role in the physiological wound healing process, overexpression of TGFβ in fibrotic tissues is detrimental, with matrix stiffness acting as an amplifier of TGFβ activity [18,62,63].

TGFβ mediates signalling through two types of transmembrane serine/threonine kinases, TGFβ type I (TGFβRI, ALK5) and type II (TGFβRII) receptors [64,65]. Interaction of TGFβ with its receptors elicits a multitude of cellular responses via both SMAD and non-SMAD-dependent signalling pathways [56,66].

In the canonical SMAD-dependent pathway, binding of TGFβ to TGFβRII induces the recruitment and phosphorylation of TGFβRI, activating its kinase activity [67]. Signal transduction is propagated through phosphorylation of receptor-regulated SMAD proteins, namely SMAD2 and SMAD3, which then translocate as a complex with SMAD4 into the nucleus and regulate the transcription of the target genes [66,68]. Alternatively, TGFβ modulates downstream cellular responses via non-SMAD pathways [69,70], such as mitogen-activated protein kinases (MAPK) signalling cascades involving p38, extracellular signal-regulated kinase (ERK) and c-Jun N-terminal kinase (JNK). TGFβ also activates Rho-like GTPases, phosphatidylinositol-3-kinase PI3K/Akt and nuclear factor-κB (NF-κB) pathways.

Although non-SMAD pathways have been reported to be involved in TGFβ-induced fibrosis, TGFβ mainly drives fibrosis via SMAD signalling [57].

Other important soluble profibrogenic mediators include platelet-derived growth factor (PDGF), connective tissue growth factor (CTGF), vascular endothelial growth factor (VEGF), fibroblast growth factor (FGF) and epithelial growth factor (EGF). These mediators function as ligands for the corresponding receptor tyrosine kinases (RTKs). Activation of RTKs after injury plays an important role in organ fibrosis [52,71–74]. Binding of the ligand results in RTK dimerization and autophosphorylation, with subsequent activation of the downstream signalling pathways [75]. Among the mentioned growth factors, members of the PDGF family are potent mitogens and chemo-attractants for mesenchymal cells in most organs, driving their recruitment and proliferation during fibrosis [76,77]. Similar to TGFβ, PDGF stimulates synthesis of major ECM components.

Altogether, the diversity in molecular pathways driving fibrogenesis highlights the complexity of the regulation of organ fibrosis.

ANTIFIBROTIC STRATEGIES UNDER CLINICAL DEVELOPMENT

Decades of research significantly advanced our understanding of the central cellular and molecular mechanisms underlying the progression of organ fibrosis. This has laid the foundation for the development and clinical validation of rational mechanism-based antifibrotic strategies. Here, we outline current therapies that are under clinical development for fibrotic diseases in liver, kidney and gut, as these organs comprise the main focus of this thesis (Figure 3).

Figure 3. Pipeline for therapeutic strategies to combat organ fibrosis. Agents are classified based on their mode of action, and colours code for a particular organ – liver (yellow), kidney (red) and intestine (green). Therapeutics denoted by circles are experimental treatments, while stars mark those that are commercially available for indications other than fibrotic diseases of liver, kidney or gut. Data are derived from clinicaltrials.gov and literature; for more details see [14,16,77–86].

mAb, monoclonal antibody; CTGF, connective tissue growth factor; PDGF, platelet-derived growth factor; TGFβ, transforming growth factor beta; TKI, tyrosine kinase inhibitor; IL-13, interleukin-13; PDE, phosphodiesterase; NRF2, nuclear factor erythroid 2–related factor 2; CNI, calcineurin inhibitor; TNF, tumor necrosis factor; CCR2/CCR5, C-C chemokine receptor type 2/ type 5; MCP, monocyte chemoattractant protein; ASK1, apoptosis signal-regulating kinase 1;

NOX, nicotinamide adenine dinucleotide phosphate (NADPH) oxidase; PPAR, peroxisome proliferator-activated receptor;

FXR, farnesoid X receptor; HIF-PHI, hypoxia-inducible factor prolyl hydroxylase inhibitor; mTOR, mammalian target of

rapamycin; JAK, Janus kinase; ACC, acetyl-CoA carboxylase; AT1 inhibitor, angiotensin II type 1 receptor inhibitor; LOXL2,

lysyl oxidase-like 2; ERAs, endothelin-1 receptor antagonists; MMP9, matrix metalloproteinase 9.

Main antifibrotic strategies include: 1) eliminating the pathological trigger; 2) suppressing 1

inflammation; 3) preventing myofibroblast activation; and 4) promoting ECM degradation and remodelling [78]. The most straightforward way to halt the progression of fibrosis is by removing the cause of tissue damage or inflammatory stimulus, as has been observed in individuals with chronic hepatitis B infection treated with entecavir, a highly effective oral antiviral drug [10]. However, this is not always feasible, since in many fibrotic disorders the tissue-damaging stimulus is either unknown or cannot be easily eliminated. Therefore, as illustrated in Figure 3, current clinical efforts are invested in antifibrotic strategies and agents that target downstream profibrotic pathways (e.g. inhibitors of TGFβ, PDGF, CTGF), inflammation (e.g. antibodies against IL-13; monocyte chemoattractant protein, MCP; thymic humoral factor, THF), oxidative stress (e.g. inhibitors of NADPH oxidase, NOX; activator of nuclear factor erythroid 2–related factor 2, NRF2) or pathologic ECM (e.g. targeting ECM molecules and their receptors, crosslinking enzymes). Notably, while multiple pharmacological interventions are under evaluation for the treatment of liver and kidney fibrosis, there are hardly any trials investigating antifibrotic therapies for intestinal fibrosis.

Given the prominent role of TGFβ and PDGF signalling in organ fibrosis, strategies to inhibit these pathways will be addressed in this thesis. Essentially, there are three main approaches for pathway inhibition: 1) sequestering the ligand (e.g. with antisense oligonucleotides); 2) disrupting ligand- receptor interaction (e.g. with neutralizing monoclonal antibodies against the ligand – TGFβ or PDGF – or their receptors); and 3) blocking receptor kinase activity to prevent signal transduction (e.g.

with tyrosine kinase inhibitors, TKI) [87,88]. Two experimental monoclonal antibodies against TGFβ, fresolimumab and LY2382770, have reached clinical studies in patients with fibrosis-associated kidney diseases, but failed to show clear efficacy [18,89]. Evaluation of pirfenidone for liver and renal fibrosis falls under the category of “drug repurposing”, which is a pharmaceutical development strategy that oversees the reuse of existing licensed drugs for new medical indications. The indisputable benefits of drug repurposing include reduced development time and costs for the drugs, since they already have been proven safe and effective for other diseases. Pirfenidone is the first ever antifibrotic drug approved for the treatment of idiopathic pulmonary fibrosis [90]. The proposed mechanism of action of pirfenidone is inhibiting the nuclear accumulation of SMAD2/3, thus disrupting the TGFβ signalling, although the exact molecular targets remain unknown [91]. As illustrated in Figure 3, pirfenidone is not the only repurposed therapeutic agent. For instance, TKI that block the enzymatic activity of the PDGF receptor (PDGFR) represent a majority of current cancer therapies that inhibit PDGF signalling.

These small molecules bind to the ATP-binding pocket of kinases, which is a conserved structure in this group of enzymes; therefore, TKI are not entirely specific and show multi-targeted effects [77].

Among the TKI, imatinib was the first to be approved for clinical use in 2001, and it is now the frontline treatment of chronic myeloid leukaemia. Imatinib primarily targets PDGFR, BCR-Abl kinase and c-kit.

Second generation TKI have an even broader spectrum of action: for example, sorafenib, clinically approved for the treatment of renal and liver cancer, inhibits PDGFRs, FLT3, CSF1R and VEGFR [88].

TKI have been evaluated in various fibrotic diseases (e.g. sorafenib in liver fibrosis [92–94], imatinib in liver and lung fibrosis [95–98]), but only nintedanib, which targets PDGFR, VEGFR and FGFR, has been approved for the treatment of pulmonary fibrosis.

Despite overwhelming scientific efforts, none of the numerous drugs currently in clinical development have been approved for the treatment of hepatic, renal or intestinal fibrosis. Moreover, the sole drugs that are clinically available – pirfenidone and nintedanib – only slow down the progression of lung fibrosis, while the disease continues to progress and is ultimately fatal [1,99]. Hence, the resolution of organ fibrosis remains an urgent and unmet clinical need. Targeting both the inflammatory component and basic elements of the fibrotic program (i.e., growth factors, ECM, etc.) proved to be challenging.

Indeed, a lot of attention has been given to therapeutics with an anti-inflammatory/antioxidant mode

of action, highlighting the prominent role of inflammation, proinflammatory mediators and ROS in the

progression of fibrosis. However, there is growing evidence that once fibrosis becomes self-sustainable,

the mere control of inflammation is not enough to revert this process [14]. In addition, the clinical

development of drugs that directly interfere with key players in the fibrotic process is hindered by the

multifaceted nature of these factors and their role in maintaining total body homeostasis. A classic

example of this is pharmacological modulation of TGFβ function. A plethora of therapies targeting

TGFβ have been explored (reviewed in [18,58,100]), yet, only a few reached clinical trials and resulted

in positive patient outcomes. Although systemic inhibition of the TGFβ pathway may ameliorate

fibrosis, such a therapy might also block its anti-inflammatory function and lead to deleterious side

effects in patients [100]. Instead of direct inhibition of TGFβ or its receptors, selective targeting of

downstream TGFβ signalling mediators could abolish some safety concerns [18]. Furthermore, due

to the lack of clinical success with compounds that directly targeted known drivers of fibrosis, drug

discovery and development has shifted towards indirect antifibrotic agents, some of which are now

entering phase I and phase II clinical trials, as reviewed elsewhere [89,101–105]. Moreover, emerging

strategies of targeted drug delivery (by using nanoparticles, liposomes, cell-penetrating peptides,

etc.), cell therapy (e.g. by using mesenchymal stem cells) and combination therapy comprise an

exciting avenue of antifibrotic drug development [16].

1

RECIPE FOR SUCCESSFUL CLINICAL DRUG DEVELOPMENT

Before we dive into brewing a successful clinical trial, let’s take a look at our cooking pot. A simple PubMed database search tells us that for the past 40 years, more than 130.000 articles addressing fibrosis have been published, not counting reviews and articles dedicated to cystic fibrosis. Since 2016, each year is marked by more than 8000 of publications dealing with fibrosis. At the same time, more than 2000 clinical studies have been listed on clinicaltrials.gov with the keyword “fibrosis”

(excluding those mentioning “cystic fibrosis”), of which over 660 are active/ongoing. A multitude of antifibrotic strategies that had promising effects in preclinical experimental models, proved inefficient and/or unsafe in human clinical trials. To date, clinical development yielded only two approved drugs that slow down the progression of fibrosis in a single organ, the lung, increasing the life expectancy for patients with lung fibrosis with a few months. Still, no therapy exists for any other type of organ fibrosis. Oh, tearful onions.

Much like in any cooking recipe, there are several key ingredients that are required to achieve the desired outcome, in our case, clinical trials that provide strong evidence for effective antifibrotic therapies. Critical factors must be considered in the design of clinical trials, including: 1) patient recruitment and selection; 2) identification of appropriate end-points, and 3) availability of biomarkers.

Ideally, study subjects should be matched according to aetiology, age, gender, medication use and risk factors, and diagnosed with an intermediate stage of fibrosis, where dynamic changes are best detectable [101]. At present, most clinical trials continue to enrol a non-homogeneous population of patients with advance disease stages, multiple comorbidities and different pathogenic processes [102]. Diagnostically conclusive primary end-points are a major determinant of the outcome of clinical trials, but studies often rely on clinical parameters that do not fully reflect disease progression [106].

For example, renal fibrosis is poorly captured by glomerular filtration rate (GFR) or proteinuria, which are generally considered as primary end-points in almost all antifibrotic trials in kidney patients [85].

Similarly, liver function parameters, such as albumin or prothrombin time, are usually altered in the late stages of liver disease, and serum transaminases are not predictive [101]. The situation is even more complicated regarding intestinal fibrosis, since no clinically useful serum markers, nor patient- reported endpoints for assessment of therapeutic response are available [79,86]. The best read-out for a trial investigating a potential antifibrotic therapy would be fibrosis itself. Currently, tissue biopsy is considered the gold standard for assessing hepatic, renal and intestinal fibrosis. Unfortunately, such a procedure is invasive and prone to sampling error, limiting the usefulness of this approach [14,106].

Therefore, therapeutics that specifically target fibrosis in these organs, without affecting conventional clinical parameters, are especially challenging to investigate in clinical trials. This is in sharp contrast with trials in idiopathic pulmonary fibrosis, as there is a very reliable end-point, termed “forced vital capacity”. The vital capacity, measured by the amount of expelled air from the lungs after maximum inhalation, is a simple, non-invasive test and can be easily repeated with no risk for the patient [107].

Luckily, the field of liver fibrosis is catching up: the recently developed FibroScan ^® (vibration-controlled transient elastography) is a novel, non-invasive technique to assess hepatic fibrosis and steatosis that has been validated in chronic hepatitis B and C, and non-alcoholic fatty liver disease (NAFLD) [108].

Nonetheless, specific, sensitive and non-invasive biomarkers of fibrosis are essential for successful antifibrotic drug development, as they could replace the current gold standard - invasive biopsy - for assessing and staging fibrotic changes, predicting disease progression and revealing drug efficacy [14,89].

Even before we can mix these key ingredients, we have to address the core element of the recipe:

adequate preclinical evaluation is undoubtedly the foundation of successful clinical dug development.

The conventional steps in the identification process of new therapeutic targets and drug testing significantly rely on the use of in vitro systems, such as cell mono- and co-cultures, and in vivo animal models. However, the low clinical success of putative drugs argues for the poor translational efficiency of basic biomedical research using conventional preclinical tools.

While two-dimensional cell cultures comprise a simple, cost-efficient and potentially high-throughput method to study fibrosis, they lack cellular heterogeneity and relevant cell-cell and cell-ECM interactions, which are a prerequisite to mimic the multicellular character of fibrosis. To minimize the gap between cell culture and the in vivo situation, and to tackle the limited physiological relevance of most experimental models, new in vitro models are being developed, such as co-cultures, sandwich cultures and three-dimensional (3D) tissue engineered models (e.g. spheroids, decellularized organs, 3D bioprinting), reviewed in [109–111]. The latter are gaining more and more attention; however, these models do not fully recapitulate the structural organisation of human organs and still face many technical hurdles, limiting their use in fibrosis research [112].

Animal models, particularly rodents, are the gold standard for basic and preclinical research in medical

and pharmacological science. From a regulatory perspective, their use is mandatory in the transition

from preclinical to clinical studies. In the last decade, a wide variety of animal models, including surgical,

genetic, toxic and nutritional models, have been established to study renal, hepatic and intestinal

fibrosis [113–116]. However, animal testing raises many ethical and scientific concerns. A major

criticism is that animal models do not fully replicate human pathology due to the distinct differences

in anatomy, (patho)physiology, metabolic capacity and immunology [45]. In many widely used animal

models of fibrosis, the disease is provoked by triggers that are rarely, if ever, seen in human disease

(e.g. toxic injury by CCl ₄ in liver or bleomycin in lung) [29]. The use of knockout and transgenic mouse

models, in which genes/proteins that regulate fibrosis have been knocked out or overexpressed,

also poses a challenge regarding clinical translation [5]. Furthermore, most in vivo studies initiate

treatment before or at the time of disease induction, hindering extrapolation to the clinical situation

[22]. In clinical practice, therapeutic interventions are frequently initiated during late stages of the

disease, when fibrosis already exists. In addition, animal studies are more often than not performed

in single-strain young male mice, in contrast to the heterogeneous population of patients enrolled

in most clinical trials [106]. Collectively, current in vivo models have a limited predictive capacity for

drug safety and efficacy in humans, as a staggering 9 out of 10 drugs that passed animal experiments

ended up failing in human clinical trials [117]. Nevertheless, even though the translational value of

animal models is continuously questioned, most preclinical research is executed in rodents because 1

human-based models are lacking.

In conclusion, the secret to brewing a deliciously successful antifibrotic therapy for humans entails that clinical drug development refines clinical trial design, establishes fibrosis-specific biomarkers and, most importantly, improves the translational aspect of preclinical tools, including the search for alternative, human-based models.

PRECISION-CUT TISSUE SLICES – 5 MILLIMETER ORGAN MINIATURES

In light of the great need for versatile, predictive and translational preclinical models, ex vivo precision- cut tissue slices (PCTS), a functional 3D organ model, has recently caught the attention of biomedical research. PCTS represent a more physiologically relevant model compared to most of the currently used in vitro systems. Each slice, only 5 mm in diameter, contains all cell types and acellular components of the whole organ in their original configuration, while preserving the intercellular and cell-matrix interactions. In other words, PCTS are miniaturized organs that retain native tissue architecture and intact cellular environment. Therefore, PCTS generated from rodent organs provide a link between currently used in vitro and in vivo models (Figure 4).

Such organotypic cultures have been established for various organs, including lung [118], liver [119], kidney [120,121], intestine [122], heart [123] and pancreas [124]. PCTS have emerged as a tool for pharmacological and toxicological research and have been proven useful in elucidating drug metabolism, drug transport, toxicity and drug-drug interactions [125–131]. As a more recent application, the PCTS model has found its use in elucidating the mechanisms of organ fibrosis and testing the efficacy of putative antifibrotic compounds [132–137]. Additionally, the use of PCTS has expanded into the fields of nanotechnology [138], virology [139] and gene silencing [140].

The most invaluable advantage of PCTS is the ability to use human tissue, which is considered to be more predictive for human responses than animal experiments. This not only increases the predictive capacity of the model, but also eliminates the need for animal-to-human extrapolation. Thus, PCTS provide a robust screening platform that reinforces preclinical evidence of potential therapeutic targets and compounds for future clinical trials. Therefore, human PCTS help to bridge the gap between animal models and the patient setting (Figure 4). Human material for the preparation of PCTS is usually obtained from surgical waste remaining from resection surgery or non-transplantable organs. Highlighting the versatility, PCTS can also be prepared from fibrotic tissue obtained from experimental animals or patients with severe fibrosis, allowing the study of advance disease stages and the effects of compounds in a pathological environment.

Other beneficial features of the PCTS model include reproducibility, simplicity in PCTS preparation and handling and the capacity to test several experimental conditions at once. Multiple slices can be obtained from a single organ, with an option to use various organs from the same animal.

Therefore, murine and human PCTS greatly reduce animal use in accordance with the 3Rs (reduction, replacement and refinement). Still, as with any research model, PCTS culture is subjected to several limitations, which are addressed in detail in the general discussion of this thesis (Chapter 8).

Figure 4. Precision-cut tissue slices (PCTS) in the drug development process. The PCTS model reflects the complex

three-dimensional architecture of the whole organ and maintains cellular diversity – two unique features that can make

in vitro drug testing more physiologically relevant. Hence, murine PCTS might be a translational link between currently

used in vitro and in vivo models. In turn, the ability to use human tissues makes PCTS an indispensable tool in preclinical

drug development that can bridge the translational gap between animal studies and human clinical trials.

1

AIM OF THE THESIS

Prompted by scarce therapeutic options for the treatment of organ fibrosis, the field of fibrosis research is in dire need of good translational human-based preclinical models. The ex vivo PCTS model promises to meet this urgent need. The aim of this thesis was threefold (Figure 5): to deepen our knowledge of the molecular processes that take place in PCTS cultures (Part I); to demonstrate the predictive value of PCTS as a screening platform for antifibrotic drugs (Part II); and to optimize current PCTS culture conditions (Part III).

Figure 5. The concept and aim of this thesis. Precision-cut tissue slices (PCTS) were prepared from tissues of various species (mouse, rat, human), organs (liver, kidney, gut) and states (healthy, diseased). The basis of the PCTS method is obtaining thin slices of defined thickness and diameter with the use of a tissue slicer, followed by incubation of the slices in culture plates. This thesis is divided into three parts: part I is dedicated to the characterization of murine and human PCTS from healthy and diseased organs in culture; part II discusses the use of PCTS as preclinical screening platform for antifibrotic drugs; and part III delineates the optimization of PCTS culture conditions.

Although numerous molecular pathways are common in the development of organ fibrosis, justifying the term “core pathways of fibrosis”, each organ responds differently to injury and displays unique organ-specific features of fibrosis. Chapter 2 delineates the diversity in culture-induced fibrogenic responses in murine PCTS, as well as organ-specific responses to antifibrotic treatment. An incomplete understanding of culture-driven processes in PCTS halts the use of this model in preclinical research to its full potential. Therefore, chapter 3 provides a comprehensive characterization of the dynamic transcriptional changes in murine and human PCTS, originating from liver, kidney and gut, during culture. In chapter 4, the molecular processes that unfold in human PCTS from diseased organs are described and compared to human PCTS from healthy tissues, defining the role of culture and pre- existing pathology.

The use of PCTS as preclinical screening platform for antifibrotic compounds is demonstrated in

chapter 5, which details the antifibrotic efficacy of nintedanib in murine and human kidney PCTS,

emphasizing the added value of using human tissues in the drug development process. Chapter 6

describes the use of an even more extensive panel of antifibrotics to showcase the predictive and

translational power of PCTS. To this end, the ex vivo effects of pirfenidone, galunisertib (LY2157299)

and imatinib were compared to the published in vivo effects and performance in clinical trials. One

of the major limitations of the PCTS model is the relatively short viability of the slices during culture,

which precludes the investigation of long-term drug toxicity and efficacy. The research presented

in chapter 7 aimed to optimize the current culture conditions of intestinal PCTS by modifying the

composition of the culture media leading to prolonged tissue viability. Lastly, chapter 8 summarizes

and discusses the findings described in this thesis, and suggests future directions for ex vivo fibrosis

research.

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29. Friedman, S. L., Sheppard, D., Duffield, J. S. & Violette, S. Therapy for Fibrotic Diseases: Nearing the Starting Line.

Sci. Transl. Med. 5, 167sr1-167sr1 (2013).

30. Kalluri, R. EMT: When epithelial cells decide to become mesenchymal-like cells. J. Clin. Invest. 119, 1417–9 (2009).

31. Stone, R. C. et al. Epithelial-mesenchymal transition in tissue repair and fibrosis. Cell Tissue Res. 365, 495–506 (2016).

32. Lin, S.-L., Kisseleva, T., Brenner, D. A. & Duffield, J. S. Pericytes and Perivascular Fibroblasts Are the Primary Source of Collagen-Producing Cells in Obstructive Fibrosis of the Kidney. Am. J. Pathol. 173, 1617–27 (2008).

33. Humphreys, B. D. et al. Fate Tracing Reveals the Pericyte and Not Epithelial Origin of Myofibroblasts in Kidney Fibrosis. Am. J. Pathol. 176, 85–97 (2010).

34. Duffield, J. S. & Humphreys, B. D. Origin of new cells in the adult kidney: results from genetic labeling techniques.

Kidney Int. 79, 494–501 (2011).

35. Mederacke, I. et al. Fate tracing reveals hepatic stellate cells as dominant contributors to liver fibrosis independent of its aetiology. Nat. Commun. 4, 2823 (2013).

36. Dulauroy, S., Di Carlo, S. E., Langa, F., Eberl, G. & Peduto, L. Lineage tracing and genetic ablation of ADAM12+

perivascular cells identify a major source of profibrotic cells during acute tissue injury. Nat. Med. 18, 1262–70 (2012).

37. Goritz, C. et al. A Pericyte Origin of Spinal Cord Scar Tissue. Science. 333, 238–242 (2011).

38. Thomas, H., Cowin, A. & Mills, S. The Importance of Pericytes in Healing: Wounds and other Pathologies. Int. J. Mol.

Sci. 18, 1129 (2017).

39. Greenhalgh, S. N., Iredale, J. P. & Henderson, N. C. Origins of fibrosis: pericytes take centre stage. F1000Prime Rep.

5, 37 (2013).

40. Di Carlo, S. E. & Peduto, L. The perivascular origin of pathological fibroblasts. J. Clin. Invest. 128, 54–63 (2018).

41. Sorrell, J. M. & Caplan, A. I. Chapter 4 Fibroblasts—A Diverse Population at the Center of It All. in International Review of Cell and Molecular Biology 276, 161–214 (Academic Press, 2009).

42. Bucala, R., Spiegel, L. A., Chesney, J., Hogan, M. & Cerami, A. Circulating fibrocytes define a new leukocyte subpopulation that mediates tissue repair. Mol. Med. 1, 71–81 (1994).

43. Galligan, C. L. & Fish, E. N. The role of circulating fibrocytes in inflammation and autoimmunity. J. Leukoc. Biol. 93, 45–50 (2013).

44. Crisan, M. et al. A Perivascular Origin for Mesenchymal Stem Cells in Multiple Human Organs. Cell Stem Cell 3, 301–13 (2008).

45. Jun, J.-I. & Lau, L. F. Resolution of organ fibrosis. J. Clin. Invest. 128, 97–107 (2018).

46. Afratis, N. A., Selman, M., Pardo, A. & Sagi, I. Emerging insights into the role of matrix metalloproteases as therapeutic targets in fibrosis. Matrix Biol. 68–69, 167–79 (2018).

47. Duffield, J. S. Fibrosis. in Kidney Development, Disease, Repair and Regeneration 293–314 (Elsevier, 2016).

doi:10.1016/B978-0-12-800102-8.00023-0

48. Apte, S. S. & Parks, W. C. Metalloproteinases: A parade of functions in matrix biology and an outlook for the future.

Matrix Biol. 44–46, 1–6 (2015).

49. McKleroy, W., Lee, T.-H. & Atabai, K. Always cleave up your mess: targeting collagen degradation to treat tissue fibrosis. Am. J. Physiol. Cell. Mol. Physiol. 304, L709–21 (2013).

50. Giannandrea, M. & Parks, W. C. Diverse functions of matrix metalloproteinases during fibrosis. Dis. Model. Mech. 7, 193–203 (2014).

51. Karamanos, N. K., Theocharis, A. D., Neill, T. & Iozzo, R. V. Matrix modeling and remodeling: A biological interplay regulating tissue homeostasis and diseases. Matrix Biol. 75–76, 1–11 (2019).

52. Bettenworth, D. & Rieder, F. Pathogenesis of Intestinal Fibrosis in Inflammatory Bowel Disease and Perspectives for Therapeutic Implication. Dig. Dis. 35, 25–31 (2017).

53. Wells, R. G. The role of matrix stiffness in regulating cell behavior. Hepatology 47, 1394–400 (2008).

54. Herrera, J., Henke, C. A. & Bitterman, P. B. Extracellular matrix as a driver of progressive fibrosis. J. Clin. Invest. 128, 45–53 (2018).

55. Li, X., Zhu, L., Wang, B., Yuan, M. & Zhu, R. Drugs and Targets in Fibrosis. Front. Pharmacol. 8, 855 (2017).

56. Meng, X., Nikolic-Paterson, D. J. & Lan, H. Y. TGF-β: the master regulator of fibrosis. Nat. Rev. Nephrol. 12, 325–38 (2016).

REFERENCES

1. Horowitz, J. C. & Thannickal, V. J. Mechanisms for the Resolution of Organ Fibrosis. Physiology 34, 43–55 (2019).

2. Wynn, T. A. Common and unique mechanisms regulate fibrosis in various fibroproliferative diseases. J. Clin. Invest.

117, 524–9 (2007).

3. Ghosh, A. K., Quaggin, S. E. & Vaughan, D. E. Molecular basis of organ fibrosis: Potential therapeutic approaches.

Exp. Biol. Med. 238, 461–81 (2013).

4. Pakshir, P. & Hinz, B. The big five in fibrosis: Macrophages, myofibroblasts, matrix, mechanics, and miscommunication. Matrix Biol. 68–69, 81–93 (2018).

5. Mehal, W. Z., Iredale, J. & Friedman, S. L. Scraping fibrosis: Expressway to the core of fibrosis. Nat. Med. 17, 552–3 (2011).

6. Duffield, J. S. Cellular and molecular mechanisms in kidney fibrosis. J. Clin. Invest. 124, 2299–306 (2014).

7. Rockey, D. C., Bell, P. D. & Hill, J. A. Fibrosis — A Common Pathway to Organ Injury and Failure. N. Engl. J. Med. 372, 1138–49 (2015).

8. Zeisberg, M. & Kalluri, R. Cellular Mechanisms of Tissue Fibrosis. 1. Common and organ-specific mechanisms associated with tissue fibrosis. Am. J. Physiol. Physiol. 304, C216–25 (2013).

9. Schaefer, L. Decoding fibrosis: Mechanisms and translational aspects. Matrix Biol. 68–69, 1–7 (2018).

10. Wynn, T. A. & Ramalingam, T. R. Mechanisms of fibrosis: therapeutic translation for fibrotic disease. Nat. Med. 18, 1028–40 (2012).

11. Zhang, Q. et al. Circulating mitochondrial DAMPs cause inflammatory responses to injury. Nature 464, 104–7 (2010).

12. Heymann, F. & Tacke, F. Immunology in the liver — from homeostasis to disease. Nat. Rev. Gastroenterol. Hepatol.

13, 88–110 (2016).

13. Borthwick, L. A., Wynn, T. A. & Fisher, A. J. Cytokine mediated tissue fibrosis. Biochim. Biophys. Acta - Mol. Basis Dis.

1832, 1049–60 (2013).

14. Allinovi, M., De Chiara, L., Angelotti, M. L., Becherucci, F. & Romagnani, P. Anti-fibrotic treatments: A review of clinical evidence. Matrix Biol. 68–69, 333–54 (2018).

15. Mack, M. Inflammation and fibrosis. Matrix Biol. 68–69, 106–21 (2018).

16. Weiskirchen, R., Weiskirchen, S. & Tacke, F. Organ and tissue fibrosis: Molecular signals, cellular mechanisms and translational implications. Mol. Aspects Med. 65, 2–15 (2019).

17. Hinz, B. et al. The myofibroblast: One function, multiple origins. Am. J. Pathol. 170, 1807–16 (2007).

18. Györfi, A. H., Matei, A.-E. & Distler, J. H. W. Targeting TGF-β signaling for the treatment of fibrosis. Matrix Biol.

68–69, 8–27 (2018).

19. Desmoulière, A., Darby, I. A. & Gabbiani, G. Normal and Pathologic Soft Tissue Remodeling: Role of the Myofibroblast, with Special Emphasis on Liver and Kidney Fibrosis. Lab. Investig. 83, 1689–707 (2003).

20. Ricard-Blum, S., Baffet, G. & Théret, N. Molecular and tissue alterations of collagens in fibrosis. Matrix Biol. 68–69, 122–49 (2018).

21. Darby, I., Skalli, O. & Gabbiani, G. Alpha-smooth muscle actin is transiently expressed by myofibroblasts during experimental wound healing. Lab. Invest. 63, 21–9 (1990).

22. Boor, P., Ostendorf, T. & Floege, J. Renal fibrosis: novel insights into mechanisms and therapeutic targets. Nat. Rev.

Nephrol. 6, 643–56 (2010).

23. Hinz, B. The role of myofibroblasts in wound healing. Curr. Res. Transl. Med. 64, 171–7 (2016).

24. Campanholle, G., Ligresti, G., Gharib, S. a & Duffield, J. S. Cellular Mechanisms of Tissue Fibrosis. 3. Novel mechanisms of kidney fibrosis. Am. J. Physiol. Physiol. 304, C591–603 (2013).

25. Parola, M. & Robino, G. Oxidative stress-related molecules and liver fibrosis. J. Hepatol. 35, 297–306 (2001).

26. Luangmonkong, T. et al. Targeting Oxidative Stress for the Treatment of Liver Fibrosis. in Reviews of physiology, biochemistry and pharmacology 175, 71–102 (2018).

27. Carthy, J. M. TGFβ signaling and the control of myofibroblast differentiation: Implications for chronic inflammatory disorders. J. Cell. Physiol. 233, 98–106 (2018).

28. Zeisberg, M. & Kalluri, R. Fibroblasts emerge via epithelial-mesenchymal transition in chronic kidney fibrosis.

Front. Biosci. 13, 6991–8 (2008).

1

86. Bettenworth, D. & Rieder, F. Medical therapy of stricturing Crohn’s disease: what the gut can learn from other organs - a systematic review. Fibrogenesis Tissue Repair 7, 5 (2014).

87. Neuzillet, C. et al. Targeting the TGFβ pathway for cancer therapy. Pharmacol. Ther. 147, 22–31 (2015).

88. Papadopoulos, N. & Lennartsson, J. The PDGF/PDGFR pathway as a drug target. Mol. Aspects Med. 62, 75–88 (2018).

89. François, H. & Chatziantoniou, C. Renal fibrosis: Recent translational aspects. Matrix Biol. 68–69, 318–32 (2018).

90. Azuma, A. et al. Exploratory analysis of a phase III trial of pirfenidone identifies a subpopulation of patients with idiopathic pulmonary fibrosis as benefiting from treatment. Respiratory Research. 12, 143 (2011).

91. Choi, K. et al. Pirfenidone inhibits transforming growth factor-β1-induced fibrogenesis by blocking nuclear translocation of Smads in human retinal pigment epithelial cell line ARPE-19. Mol. Vis. 18, 1010–20 (2012).

92. Hong, F., Chou, H., Fiel, M. I. & Friedman, S. L. Antifibrotic Activity of Sorafenib in Experimental Hepatic Fibrosis:

Refinement of Inhibitory Targets, Dosing, and Window of Efficacy In Vivo. Dig. Dis. Sci. 58, 257–64 (2012).

93. Deng, Y.-R. et al. STAT3-mediated attenuation of CCl4-induced mouse liver fibrosis by the protein kinase inhibitor sorafenib. J. Autoimmun. 46, 25–34 (2013).

94. Pinter, M. et al. The effects of sorafenib on the portal hypertensive syndrome in patients with liver cirrhosis and hepatocellular carcinoma - a pilot study. Aliment. Pharmacol. Ther. 35, 83–91 (2012).

95. Yoshiji, H. et al. Imatinib mesylate (STI-571) attenuates liver fibrosis development in rats. Am J Physiol Gastrointest Liver Physiol 288, 907–13 (2005).

96. Neef, M. et al. Oral imatinib treatment reduces early fibrogenesis but does not prevent progression in the long term. J. Hepatol. 44, 167–75 (2006).

97. Daniels, C. E. et al. Imatinib mesylate inhibits the profibrogenic activity of TGF-β and prevents bleomycin- mediated lung fibrosis. J. Clin. Invest. 114, 1308–16 (2004).

98. Daniels, C. E. et al. Imatinib Treatment for Idiopathic Pulmonary Fibrosis. Am. J. Respir. Crit. Care Med. 181, 604–10 (2010).

99. Martinez, F. J. et al. Idiopathic pulmonary fibrosis. Nat. Rev. Dis. Prim. 3, 17074 (2017).

100. Isaka, Y. Targeting TGF-β Signaling in Kidney Fibrosis. Int. J. Mol. Sci. 19, 2532 (2018).

101. Schuppan, D. & Kim, Y. O. Evolving therapies for liver fibrosis. J. Clin. Invest. 123, 1887–901 (2013).

102. Breyer, M. D. & Susztak, K. The next generation of therapeutics for chronic kidney disease. Nat. Rev. Drug Discov.

15, 568–88 (2016).

103. Yoon, Y., Friedman, S. & Lee, Y. Antifibrotic Therapies: Where Are We Now? Semin. Liver Dis. 36, 087–98 (2016).

104. Nanthakumar, C. B. et al. Dissecting fibrosis: therapeutic insights from the small-molecule toolbox. Nat. Rev. Drug Discov. 14, 693–720 (2015).

105. Declèves, A.-E. & Sharma, K. Novel targets of antifibrotic and anti-inflammatory treatment in CKD. Nat. Rev.

Nephrol. 10, 257–67 (2014).

106. Djudjaj, S. & Boor, P. Cellular and molecular mechanisms of kidney fibrosis. Mol. Aspects Med. 65, 16–36 (2019).

107. Wells, A. U. Forced vital capacity as a primary end point in idiopathic pulmonary fibrosis treatment trials: making a silk purse from a sow’s ear. Thorax 68, 309–10 (2013).

108. Singh, S., Muir, A. J., Dieterich, D. T. & Falck-Ytter, Y. T. American Gastroenterological Association Institute Technical Review on the Role of Elastography in Chronic Liver Diseases. Gastroenterology 152, 1544–77 (2017).

109. Mazza, G., Al-Akkad, W. & Rombouts, K. Engineering in vitro models of hepatofibrogenesis. Adv. Drug Deliv. Rev.

121, 147–57 (2017).

110. Breslin, S. & O’Driscoll, L. Three-dimensional cell culture: the missing link in drug discovery. Drug Discov. Today 18, 240–9 (2013).

111. Weinhart, M., Hocke, A., Hippenstiel, S., Kurreck, J. & Hedtrich, S. 3D organ models—Revolution in pharmacological research? Pharmacol. Res. 139, 446–51 (2019).

112. Lee, Y., Ahn, S. I. & Kim, Y. Organs-on-Chips. in Encyclopedia of Biomedical Engineering 384–93 (Elsevier, 2019).

113. Buhl, E. M. et al. The role of PDGF-D in healthy and fibrotic kidneys. Kidney Int. 89, 848–61 (2016).

114. Yang, H.-C., Zuo, Y. & Fogo, A. B. Models of chronic kidney disease. Drug Discov. Today Dis. Model. 7, 13–9 (2010).

115. Liedtke, C. et al. Experimental liver fibrosis research: update on animal models, legal issues and translational aspects. Fibrogenesis Tissue Repair 6, 19 (2013).

57. Kim, K. K., Sheppard, D. & Chapman, H. A. TGF-beta 1 Signaling and Tissue Fibrosis. Cold Spring Harb. Perspect. Biol.

10, a022293 (2018).

58. Walton, K. L., Johnson, K. E. & Harrison, C. A. Targeting TGF-β Mediated SMAD Signaling for the Prevention of Fibrosis. Front. Pharmacol. 8, 461 (2017).

59. Biancheri, P. et al. The role of transforming growth factor (TGF)-β in modulating the immune response and fibrogenesis in the gut. Cytokine Growth Factor Rev. 25, 45–55 (2014).

60. Danen, E. H. & Sonnenberg, A. Integrins in regulation of tissue development and function. J. Pathol. 200, 471–80 (2003).

61. Henderson, N. C. et al. Targeting of αv integrin identifies a core molecular pathway that regulates fibrosis in several organs. Nat. Med. 19, 1617–24 (2013).

62. Hinz, B. Tissue stiffness, latent TGF-β1 activation, and mechanical signal transduction: Implications for the pathogenesis and treatment of fibrosis. Curr. Rheumatol. Rep. 11, 120–6 (2009).

63. Zhang, Y. E. Mechanistic insight into contextual TGF-β signaling. Curr. Opin. Cell Biol. 51, 1–7 (2018).

64. Heldin, C.-H. & Moustakas, A. Signaling Receptors for TGF-β Family Members. Cold Spring Harb. Perspect. Biol. 8, a022053 (2016).

65. Katz, L. H. et al. TGF-β signaling in liver and gastrointestinal cancers. Cancer Lett. 379, 166–72 (2016).

66. Derynck, R. & Zhang, Y. E. Smad-dependent and Smad-independent pathways in TGF-β family signalling. Nature 425, 577–84 (2003).

67. Shi, Y. & Massagué, J. Mechanisms of TGF-β Signaling from Cell Membrane to the Nucleus. Cell 113, 685–700 (2003).

68. Massagué, J. TGFβ signalling in context. Nat. Rev. Mol. Cell Biol. 13, 616–30 (2012).

69. Mu, Y., Gudey, S. K. & Landström, M. Non-Smad signaling pathways. Cell Tissue Res. 347, 11–20 (2012).

70. Zhang, Y. E. Non-Smad pathways in TGF-β signaling. Cell Res. 19, 128–39 (2009).

71. Floege, J., Eitner, F. & Alpers, C. E. A New Look at Platelet-Derived Growth Factor in Renal Disease. J. Am. Soc.

Nephrol. 19, 12–23 (2008).

72. Strutz, F. et al. Basic fibroblast growth factor expression is increased in human renal fibrogenesis and may mediate autocrine fibroblast proliferation. Kidney Int. 57, 1521–38 (2000).

73. Zhuang, S. & Liu, N. EGFR signaling in renal fibrosis. Kidney Int. Suppl. 4, 70–4 (2014).

74. Beyer, C. & Distler, J. H. W. Tyrosine kinase signaling in fibrotic disorders. Biochim. Biophys. Acta - Mol. Basis Dis.

1832, 897–904 (2013).

75. Lemmon, M. A. & Schlessinger, J. Cell Signaling by Receptor Tyrosine Kinases. Cell 141, 1117–34 (2010).

76. Heldin, C.-H. & Westermark, B. Mechanism of Action and In Vivo Role of Platelet-Derived Growth Factor. Physiol.

Rev. 79, 1283–316 (1999).

77. Borkham-Kamphorst, E. & Weiskirchen, R. The PDGF system and its antagonists in liver fibrosis. Cytokine Growth Factor Rev. 28, 53–61 (2016).

78. Altamirano-Barrera, A., Barranco-Fragoso, B. & Méndez-Sánchez, N. Management Strategies for Liver Fibrosis.

Ann. Hepatol. 16, 48–56 (2017).

79. Rogler, G. Challenges of Translation of Anti-Fibrotic Therapies into Clinical Practice in IBD. in Fibrostenotic Inflammatory Bowel Disease 295–305 (Springer International Publishing, 2018).

80. Schwartz, D. M. et al. Erratum: JAK inhibition as a therapeutic strategy for immune and inflammatory diseases.

Nat. Rev. Drug Discov. 17, 78–78 (2018).

81. Rotman, Y. & Sanyal, A. J. Current and upcoming pharmacotherapy for non-alcoholic fatty liver disease. Gut 66, 180–90 (2017).

82. Schuppan, D., Ashfaq-Khan, M., Yang, A. T. & Kim, Y. O. Liver fibrosis: Direct antifibrotic agents and targeted therapies. Matrix Biol. 68–69, 435–51 (2018).

83. Sumida, Y. & Yoneda, M. Current and future pharmacological therapies for NAFLD/NASH. J. Gastroenterol. 53, 362–76 (2018).

84. Pinto, S. et al. Drug Discovery in Tissue Fibrosis. in Comprehensive Medicinal Chemistry III 7, 694–713 (Elsevier, 2017).

85. Klinkhammer, B. M., Goldschmeding, R., Floege, J. & Boor, P. Treatment of Renal Fibrosis—Turning Challenges into

Opportunities. Adv. Chronic Kidney Dis. 24, 117–29 (2017).

1

116. Wirtz, S. & Neurath, M. Mouse models of inflammatory bowel disease ☆. Adv. Drug Deliv. Rev. 59, 1073–83 (2007).

117. Hay, M., Thomas, D. W., Craighead, J. L., Economides, C. & Rosenthal, J. Clinical development success rates for investigational drugs. Nat. Biotechnol. 32, 40–51 (2014).

118. Henjakovic, M. et al. Ex vivo testing of immune responses in precision-cut lung slices. Toxicol. Appl. Pharmacol.

231, 68–76 (2008).

119. Graaf, I. A. de, Groothuis, G. M. & Olinga, P. Precision-cut tissue slices as a tool to predict metabolism of novel drugs. Expert Opin. Drug Metab. Toxicol. 3, 879–98 (2007).

120. Stribos, E. G. D., Seelen, M. A., van Goor, H., Olinga, P. & Mutsaers, H. A. M. Murine Precision-Cut Kidney Slices as an ex vivo Model to Evaluate the Role of Transforming Growth Factor-β1 Signaling in the Onset of Renal Fibrosis.

Front. Physiol. 8, 1026 (2017).

121. Stribos, E. G. D. et al. Precision-cut human kidney slices as a model to elucidate the process of renal fibrosis. Transl.

Res. 170, 8-16.e1 (2016).

122. Pham, B. T. et al. Precision-cut rat, mouse, and human intestinal slices as novel models for the early-onset of intestinal fibrosis. Physiol. Rep. 3, e12323 (2015).

123. Wang, K. et al. Cardiac tissue slices: preparation, handling, and successful optical mapping. Am. J. Physiol. Circ.

Physiol. 308, H1112–25 (2015).

124. Marciniak, A. et al. Using pancreas tissue slices for in situ studies of islet of Langerhans and acinar cell biology.

Nat. Protoc. 9, 2809–22 (2014).

125. Kanter, R., Monshouwer, M., Meijer, D. & Groothuis, G. Precision-Cut Organ Slices as a Tool to Study Toxicity and Metabolism of Xenobiotics with Special Reference to Non-Hepatic Tissues. Curr. Drug Metab. 3, 39–59 (2002).

126. Niu, X., de Graaf, I. a M., van der Bij, H. a & Groothuis, G. M. M. Precision cut intestinal slices are an appropriate ex vivo model to study NSAID-induced intestinal toxicity in rats. Toxicol. In Vitro 28, 1296–305 (2014).

127. Li, M., de Graaf, I. A. M. & Groothuis, G. M. M. Precision-cut intestinal slices: alternative model for drug transport, metabolism, and toxicology research. Expert Opin. Drug Metab. Toxicol. 12, 175–90 (2016).

128. Hadi, M. et al. Human Precision-Cut Liver Slices as an ex Vivo Model to Study Idiosyncratic Drug-Induced Liver Injury. Chem. Res. Toxicol. 26, 710–20 (2013).

129. Vatakuti, S., Olinga, P., Pennings, J. L. A. & Groothuis, G. M. M. Validation of precision-cut liver slices to study drug- induced cholestasis: a transcriptomics approach. Arch. Toxicol. 91, 1401–12 (2017).

130. Olinga, P. & Schuppan, D. Precision-cut liver slices: A tool to model the liver ex vivo. J. Hepatol. 58, 1252–3 (2013).

131. Lauenstein, L. et al. Assessment of immunotoxicity induced by chemicals in human precision-cut lung slices (PCLS). Toxicol. Vitr. 28, 588–99 (2014).

132. Westra, I. M., Pham, B. T., Groothuis, G. M. M. & Olinga, P. Evaluation of fibrosis in precision-cut tissue slices.

Xenobiotica 43, 98–112 (2013).

133. van de Bovenkamp, M., Groothuis, G. M. M., Meijer, D. K. F. & Olinga, P. Liver slices as a model to study fibrogenesis and test the effects of anti-fibrotic drugs on fibrogenic cells in human liver. Toxicol. In Vitro 22, 771–8 (2008).

134. Stribos, E. G. D., Hillebrands, J.-L., Olinga, P. & Mutsaers, H. A. M. Renal fibrosis in precision-cut kidney slices. Eur. J.

Pharmacol. 790, 57–61 (2016).

135. Luangmonkong, T. et al. Evaluating the antifibrotic potency of galunisertib in a human ex vivo model of liver fibrosis. Br. J. Pharmacol. 174, 3107–17 (2017).

136. Huang, X. et al. Molecular characterization of a precision-cut rat liver slice model for the evaluation of antifibrotic compounds. J Physiol Gastrointest Liver Physiol 316, 15–24 (2019).

137. Huang, X. et al. Molecular characterization of a precision-cut rat lung slice model for the evaluation of antifibrotic drugs. Am. J. Physiol. Cell. Mol. Physiol. 316, L348–57 (2019).

138. Sauer, U. G. et al. Applicability of rat precision-cut lung slices in evaluating nanomaterial cytotoxicity, apoptosis, oxidative stress, and inflammation. Toxicol. Appl. Pharmacol. 276, 1–20 (2014).

139. Dobrescu, I. et al. In vitro and ex vivo analyses of co-infections with swine influenza and porcine reproductive and respiratory syndrome viruses. Vet. Microbiol. 169, 18–32 (2014).

140. Ruigrok, M. J. R. et al. siRNA-Mediated RNA Interference in Precision-Cut Tissue Slices Prepared from Mouse Lung

and Kidney. AAPS J. 19, 1855–63 (2017).

PA RT I CH AR AC TE RI ZA TI ON OF TH E P CT S M OD EL

2 FEATURES OF FIBROGENESIS USING MURINE PRECISION-CUT TISSUE SLICES

Emilia Bigaeva Emilia Gore

Henricus A. M. Mutsaers Dorenda Oosterhuis Ruud A. Bank Miriam Boersema Peter Olinga

(Submitted)

2

ABSTRACT

Fibrosis is the hallmark of pathologic tissue remodelling in most chronic diseases. Despite advances in our understanding of the mechanisms of fibrosis, it remains uncured. Fibrogenic processes share conserved core cellular and molecular pathways across organs. In this study, we aimed to elucidate shared and organ-specific features of fibrosis using murine precision-cut tissue slices (PCTS) prepared from small intestine, liver and kidneys. PCTS displayed substantial differences in their baseline gene expression profiles: 70% of the ECM-related genes were differentially expressed across the organs.

Culture for 48h induced significant changes in ECM regulation and triggered the onset of fibrogenesis in all PCTS in organ-specific manner. TGFβ signalling was activated during 48h culture in all PCTS.

However, the degree of its involvement varied: both canonical and non-canonical TGFβ pathways

were activated in liver and kidney slices, while only canonical, Smad-dependent, cascade was

involved in intestinal slices. The treatment with galunisertib blocked the TGFβRI/SMAD2 signalling

in all PCTS, but attenuated culture-induced dysregulation of ECM homeostasis and mitigated the

onset of fibrogenesis with organ-specificity. In conclusion, regardless the many common features

in pathophysiology of organ fibrosis, PCTS displayed diversity in culture-induced responses and in

response to the treatment with TGFβRI kinase inhibitor galunisertib, even though it targets a core

fibrosis pathway. A clear understanding of the common and organ-specific features of fibrosis is the

basis for developing novel antifibrotic therapies.