Lysophosphatidic acid-mediated GPR35 signaling in CX3CR1+ macrophages regulates the intestinal cytokine milieu
Inauguraldissertation zur Erlangung der Würde eines Doktors der Philosophie vorgelegt der Philosophisch-Naturwissenschaftlichen Fakultät der Universität Basel
von
Berna Kaya
aus der Türkei
2020
Originaldokument gespeichert auf dem Dokumentenserver der Universität Basel edoc.unibas.ch
Genehmigt von der Philosophisch-Naturwissenschaftlichen Fakultät auf Antrag von
Prof. Dr. Christoph Hess Prof. Dr. Jan Hendrik Niess PD Dr. Cristian Riedel
Basel, 23.06.2020
Prof. Dr. Martin Spiess The Dean of Faculty of Science
Table of Contents
TABLE OF CONTENTS
ABBREVIATIONS ...... IV ABSTRACT ...... VII 1 INTRODUCTION ...... 1 1.1 The intestinal immune system ...... 1 1.1.1 Innate immunity in the intestine ...... 1 1.1.1.1 Intestinal epithelium...... 2 1.1.1.2 Mononuclear phagocytes ...... 5 1.1.2 Host- and microbiota-derived metabolites ...... 10 1.2 Inflammatory bowel diseases ...... 14 1.2.1 Host immune system and genetics ...... 15 1.2.2 Microbiome ...... 15 1.2.3 Diet and other environmental factors ...... 16 1.2.4 Current therapy options ...... 17 1.3 G protein-coupled receptor (GPR) 35 ...... 19 1.4 Aims of the study ...... 22 2 METHODS ...... 23 2.1 Human samples ...... 23 2.2 Animals ...... 25 2.2.1 Mouse lines ...... 25 2.2.2 Generation of Gpr35-IRES-tdTomato knock-in mice ...... 25 2.2.3 Construction of Gpr35-flox and -knockout (KO) mice ...... 27 2.2.4 Zebrafish lines ...... 29 2.3 In vivo experiments ...... 31 2.3.1 Dextran sodium sulfate-induced colitis mouse model ...... 31 2.3.2 Mouse endoscopy ...... 31 2.3.3 Treatment of zebrafish with 2,4,6-Trinitrobenzenesulfonic acid ...... 31 2.3.4 Treatment of zebrafish with antibiotics ...... 32 2.3.5 Treatment of mice with antibiotics ...... 32 2.3.6 Challenging of mice with E. coli-CFP ...... 32 2.3.7 Exposure of zebrafish with Vibrio anguillarum ...... 32 2.3.9 LPA treatment of zebrafish larvae for cytokine expression analysis ...... 33 2.3.8 In vivo macrophage migration assay in zebrafish ...... 33 2.4 Histology and imaging ...... 33 2.4.1 Preparation of RNA probes for Gpr35b in zebrafish ...... 33 2.4.2 In situ hybridization for gpr35b in zebrafish ...... 34 2.4.3 GPR35 staining on human biopsies and mouse tissues ...... 35 2.4.4 Hematoxylin-eosin (H&E) staining and histological scoring ...... 36 2.4.5 Autotaxin staining of mouse colon ...... 36 2.4.6 Ex vivo imaging of mouse tissues ...... 36 2.5 Cell isolation ...... 37
III Table of Contents
2.6 Antibodies, cell staining, and flow cytometry ...... 37 2.7 RNA extraction and quantitative PCR ...... 40 2.8 RNA sequencing ...... 41 2.9 In vitro assays ...... 42 2.9.1 3’-5’-Cyclic adenosine monophosphate (cAMP) assay ...... 42 2.9.2 Mouse bone marrow-derived macrophages (BMDMs) ...... 43 2.9.3 Trans-well migration assay for mouse BMDMs ...... 43 2.9.4 Enzyme-linked immunosorbent assay (ELISA) for corticosterone ...... 44 2.9.5 LPA ELISA using ex vivo colonic explants of DSS-treated mice ...... 44 2.10 Western blot LPA- or LPS- treated BMDMs ...... 44 2.11 Identification of GPR35T108M and TNF blocker response in IBD patients ...... 45 2.12 Statistical analysis...... 45 3 RESULTS ...... 46 3.1 Colonic macrophages express GPR35 ...... 46 3.1.1 Gpr35b is predominant in the intestine of zebrafish ...... 46 3.1.2 Gpr35 is expressed in the human and mouse gastrointestinal tract ...... 48 3.1.3 Macrophages in the intestinal lamina propria are GPR35+ ...... 48 3.1.4 GPR35+ and GPR35- colonic macrophages are transcriptionally distinct ...... 51 3.2 Regulation of GPR35 by the microbiota and intestinal inflammation ...... 52 3.2.1 Gpr35 expression is microbiota-dependent in zebrafish and mice ...... 52 3.2.2 Intestinal inflammation elevates GPR35 expression in zebrafish and mice ...... 54 3.2.3 Bacterial infection or colonization induces GPR35 in the intestine ...... 55 3.2.4 Increased number of GPR35+ cells in patients with ulcerative colitis ...... 56 3.3 Lysophosphatidic acid signaling depends on GPR35 in macrophages ...... 57 3.3.1 LPA inhibits the cAMP release in humanGPR35-transfected cells ...... 57 3.3.2 LPA leads to GPR35-dependent Tnf induction in zebrafish ...... 58 3.3.3 GPR35-deficiency leads to altered LPA signaling in BMDMs ...... 59 3.3.4 LPA facilitates GPR35-dependent chemotaxis of macrophages...... 63 3.4 Intestinal inflammation enhances Autotaxin and LPA production ...... 64 3.5 Macrophage-expressed GPR35 is protective in DSS colitis mouse model ...... 67 3.5.1 GPR35-deficient mice have augmented DSS-induced colitis ...... 67 3.5.2 Macrophage-specific GPR35 deletion leads to aggravated colitis ...... 68 3.5.3 LPA attenuates DSS colitis in a macrophage-expressed GPR35 manner ...... 70 3.5.4 Gpr35ΔCx3cr1 mice have enhanced neutrophil infiltration during DSS colitis ...... 71 3.5.5 GPR35 deletion in macrophages causes reduced TNF production in colitis ...... 73 3.6 TNF attenuates colitis and induces corticosterone synthesis in Gpr35ΔCx3cr1 mice 74 3.6.1 TNF is protective in Gpr35ΔCx3cr1 mice during DSS colitis ...... 74 3.6.2 TNF induces Cyp11b1 and corticosterone synthesis in Gpr35ΔCx3cr1 mice ...... 76 3.6.3 TNF does not influence DSS colitis severity in WT mice ...... 77 3.7 GPR35T108M variant might correlate with TNF blocker responses in IBD ...... 78 4 DISCUSSION ...... 79 4.1 Heterogeneous expression of GPR35 in intestinal lamina propria monocytes and macrophages ...... 79
IV Table of Contents
4.2 GPR35-mediated LPA signaling in macrophage cytokine and migration responses ...... 79 4.3 Regulation of ATX/LPA/GPR35 axis in the intestine ...... 81 4.4 Reduced TNF and corticosterone levels during DSS colitis upon macrophage- specific GPR35 deletion ...... 83 4.5 Conclusion and outlook ...... 84 6 APPENDIX ...... 86 6.1 Supplementary Information ...... 86 6.2 List of Tables and Figures ...... 90 6.2.1 List of Tables ...... 90 6.2.2 List of Figures ...... 90 6.3 Acknowledgements ...... 93 7 REFERENCES...... 94
V Abbreviations
ABBREVIATIONS
AMP Anti-microbial peptide
ATX Autotaxin
BMDM Bone marrow-derived macrophages
cAMP 3’-5’-Cyclic adenosine monophosphate
CCL C-C motif chemokine ligand
CD Crohn’s disease
cDC Conventional dendritic cell
CFP Cyan fluorescent protein co-LP Colonic lamina propria
CS Corticosterone
CXCL C-X-C motif chemokine ligand
Cyp11a1 Cytochrome P450 family 11 subfamily A member 1
Cyp11b1 Cytochrome P450 family 11 subfamily B member 1
DSS Dextran sodium sulfate
EC50 Effective concentration
EGF Epidermal growth factor
ER Estrogen receptor
GALT Gut-associated lymphoid tissue
GC Glucocorticoid
GF Germ-free
GFP Green fluorescent protein
GPCR G protein-coupled receptor
GPR35 G protein-coupled receptor 35
GWAS Genome-wide association study
IV Abbreviations
IBD Inflammatory bowel disease
IEC Intestinal epithelial cell
IFN Interferon
ILF Isolated lymphoid follicle
IgA Immunoglobulin A
IL Interleukin
ILC Innate lymphoid cell iNOS Pathogen-inducible nitric oxide synthase
ISC Intestinal stem cell
KYNA Kynurenic acid
LP Lamina propria
LPA Lysophosphatidic acid
Ly6C Lymphocyte antigen 6 complex locus C
M cell Microfold cell
MLN Mesenteric lymph node mRNA Messenger ribonucleic acid
NF-κB Nuclear factor kappa-light-chain-enhancer of activated B cells
NLRP NOD-like receptor, pyrin domain-containing
NO Nitric oxide
NOD Nucleotide-binding oligomerization
PBS Phosphate-buffered saline pDC Plasmacytoid dendritic cell
RA Retinoic acid
RNA Ribonucleic acid si-LP Small intestinal lamina propria
V Abbreviations
SNP Single nucleotide polymorphism
tdT tdTomato
TGF Transforming growth factor
Th T helper
TLR Toll-like receptor
TNBS 2,4,6-Trinitrobenzenesulfonic acid
TNF Tumor necrosis factor
Treg Regulatory T cell
UC Ulcerative colitis
WT Wild type
VI Abstract
ABSTRACT
Metabolites derived from the host or the microbiota regulate intestinal immunity via
their G protein-coupled receptors (GPCRs). An aberrant relationship with the gut
microbiota leads to gastrointestinal disorders, including inflammatory bowel diseases
(IBD) in genetically predisposed individuals. G protein-coupled receptor (GPR)35 has
been identified as one of the risk genes in IBD. Nonetheless, mechanisms by which
GPR35 contributes to intestinal immunity are understudied. In this Ph.D. thesis, we aim to describe a role for macrophage-expressed GPR35 in regulating the inflammatory cues during intestinal inflammation. Using RNA sequencing, we found that GPR35 in colonic macrophages correlated with the expression of pro- inflammatory cytokines. Aiming to deorphanize the GPR35, we have used a GPCR ligand identification assay where LPA led to the activation of inhibitory G protein- mediated signaling in GPR35-transfected cells. In zebrafish and mice, we observed that LPA causes upregulation of pro-inflammatory cytokine genes in a GPR35- dependent manner. Specific deletion of GPR35 in CX3CR1+ macrophages resulted in
exacerbated dextran sodium sulfate (DSS)-induced colitis, decreased TNF in colonic
macrophages, and lower corticosterone production in mice. The administration of TNF
alleviated the severity of the colitis and was associated with restored corticosterone
levels. In conclusion, the data presented in this thesis provide evidence that LPA
signaling mediates the intestinal cytokine milieu via GPR35 in CX3CR1+
macrophages.
VII Introduction
1 INTRODUCTION
1.1 The intestinal immune system
The intestine is considered as the largest immune organ in the body and is the primary
site where the immune system interacts with the luminal contents, including food
antigens and the microbiota. In the human body, the intestinal tract holds the largest reservoir of the microbiota containing bacteria, fungi, viruses, and protozoa (Kamada
et al., 2013). The intestinal immune system provides a physical and biochemical
barrier that prevents pathogenic microorganisms and toxins from entering the host
system, and also that is vital to establish tolerance to antigens and commensal
bacteria. This barrier is composed of an epithelial layer, the lamina propria where the
immune cells reside, and the gut-associated lymphoid tissues (GALT) including the
mesenteric lymph nodes (MLN), the Peyer’s patch (PP) in the small intestine, and also
isolated lymphoid follicles (ILFs) (Mowat & Agace, 2014).
A diverse and “healthy” microbiota is crucial for the development and maintenance of
host immunity and metabolism (Rooks & Garrett, 2016). A compromise in the
protective intestinal barrier leads to dysregulated immune responses that are
associated with the onset of not only intestinal disorders such as inflammatory bowel
diseases (IBD) and celiac disease but also of cardiovascular diseases, type 1 and 2
diabetes, obesity, cardiovascular disease, infections, allergies, rheumatoid arthritis,
multiple sclerosis, asthma and cancer (D. Kim et al., 2017).
1.1.1 Innate immunity in the intestine
Innate immunity comprises of immune processes that are inherited but not learned or
adapted. Effector molecules involved in innate responses include a limited number of
secreted proteins and receptors encoded by inborn genes that do not undergo
1 Introduction
rearrangements (Medzhitov & Janeway, 1997). Most microorganisms are recognized
and eliminated by the innate immune system rapidly within minutes or hours, that is particularly important in the intestine where microbial and foreign antigens
continuously challenge the host immunity. Therefore, the cells of the innate intestinal
immunity have evolved to prevent the invasion of toxins and pathogens without
initiating a systemic or local inflammation. These cells include epithelial cells,
mononuclear phagocytes, granulocytes, mast cells, innate lymphoid cells, and
intraepithelial lymphocytes.
1.1.1.1 Intestinal epithelium
The gastrointestinal tract is an organ specializing in food digestion, but it also serves
as a mucosal surface barrier between the host and the inhabiting microbiota. The
intestinal epithelial layer is a one-cell internal lining, consisted of epithelial cells,
between the lumen and the lamina propria. Intestinal epithelial cells are an integral
part of innate immunity despite not being considered as bona fide cells of the innate
immune system. They have a crucial role as the physical barrier against external
factors as the first line of defense and also maintains homeostasis with the commensal
bacteria. Intestinal epithelial cells (IECs) allow transport of ions, nutrients, and water
but prevent the invasion of bacterial toxins and pathogens (Rescigno, 2011). The
epithelial tight junction protein complexes, including occludins, claudins, zonula
occludens, tricellulin, cingulin, and junctional adhesion molecules, are the rate-limiting
factors for paracellular permeability termed as “gate and fence function” (Diamond,
1977; Gunzel & Yu, 2013).
The epithelium in the small intestine contains the villi, protrusions of the intestinal wall,
and invaginations called the crypts. On the other hand, villi are absent in the colon,
2 Introduction
where there is a flat epithelial surface with crypts. Intestinal epithelial cells have rapid
cell turnover, with constant replenishment from the stem cells residing at the base of
the crypts (Gehart & Clevers, 2019). Crypt base columnar cells, dispersed between
the Paneth cells, are considered as intestinal stem cells (ISCs) identified by genetic
lineage tracing experiments where Lgr5 was used as an ISC marker (Barker et al.,
2007). These stem cells and the early progenitor cells compete for niche space.
Failure to retain in the niche results in the initiation of the differentiation process. At
this stage, the cells are called transit-amplifying cells as they move upwards towards
the tip of the villus. After 3-5 days, the terminally differentiated cells undergo apoptosis
and are shed into the intestinal lumen (Gehart & Clevers, 2019). There are six different
types of mature intestinal epithelial cells: enterocytes and M cells, which are
absorptive, and Paneth, goblet, enteroendocrine and tuft cells, which are secretory.
Enterocytes, the most abundant IECs, are responsible for the absorption of nutrients
and water, whereas M cells overlying Peyer’s patches, present in the small but not
large intestine, are central to the uptake of antigens for their presentation to the
immune cells (Ohno, 2016). Paneth cells, a component of the stem cell niche in only
the small intestine but not the colon, secret antimicrobial peptides and support the
ISCs e.g. via EGF and Wnt3a (Clevers & Bevins, 2013; Sato et al., 2011). Goblet cells
secret Muc2 forming the protective mucus layer, enteroendocrine cells produce
hormones, and tuft cells are crucial for helminth detection and IL-25 mediated ILC2
expansion (Gribble & Reimann, 2016; Johansson & Hansson, 2016; von Moltke et al.,
2016).
Another function of IECs in mucosal immunity involves TLRs, in particular, TLR2,
TLR3, TLR5, and TLR9 (Abreu, 2010). TLR activation leads to NF-kB signaling
resulting in secretion of cytokines and chemokines (TNF, IL-6, IL-8, IL-18, CCL20),
3 Introduction induction of antimicrobial factors (iNOS, α-defensins, β-defensins, RegIII-β, RegIII-γ, lysozyme) and mucin granule exocytosis (Abreu, 2010; Coulombe et al., 2016; Gong et al., 2010; Johansson & Hansson, 2016).
In addition to TLRs, IECs also express NOD-like receptor proteins e.g. Nlrp6, Nlrc4 to activate inflammasomes and polymeric Ig receptors to transport the secretory IgA into the lumen (Birchenough et al., 2016; Hooper et al., 2001; Rauch et al., 2017).
Figure 1.1 Intestinal epithelial cells in innate immunity
The intestinal mucosa comprises of an intestinal epithelial cell (IEC) layer, the lamina propria (LP), and muscularis mucosae (M). The mucus constitutes a biochemical barrier separating the mucosa and the lumen with food antigens and the microbiota. In the colon (B), the mucus layer consists of an outer mucus layer (OML) and an inner mucus layer (IML) as opposed to the small intestine (A), where there is only one layer of mucus. Intestinal epithelial cells are an integral part of the intestinal immune system with many functions exemplified in the illustration.
4 Introduction
Interestingly, intestinal epithelial cells produce glucocorticoids (GCs), including corticosterone, the most abundantly produced bioactive GC, that exert anti- inflammatory functions (Cima et al., 2004). GCs are lipid hormones that are synthesized from cholesterol. Due to their immunosuppressive properties, they are used as therapy in various inflammatory disorders, including IBD (Rutgeerts, 1998).
Adrenal glands are the major source of GCs; however, extra-adrenal GC synthesis is evident in tissues such as the brain, thymus, skin, and intestine (Noti et al., 2009).
They have been shown to prevent the production of pro-inflammatory cytokines
(Almawi et al., 1996; Kunicka et al., 1993), to induce apoptosis in immune cells
(Amsterdam et al., 2002), and to enhance tight junction proteins and therefore epithelial barrier function (Boivin et al., 2007).
1.1.1.2 Mononuclear phagocytes
The intestinal lamina propria is home to mononuclear phagocytes, including monocytes, tissue-resident macrophages, and dendritic cells.
Mature intestinal macrophages are distinguished from other cell types by the markers CD64, F4/80, and Mer tyrosine kinase (MerTK) (Cerovic et al., 2014;
Tamoutounour et al., 2012). Additionally, all of them express high levels of MHCII and
CX3CR1 (Bain & Mowat, 2014). In contrast to other tissue-resident macrophages, intestinal macrophages do not develop from the embryonic yolk sac or the fetal liver.
Instead, they are constantly replenished by Ly6Chigh monocytes (Bain et al., 2013).
Migration of Ly6Chigh monocytes, both from the bone marrow and into the intestine, is
dependent on CCR2 (Bain et al., 2013; Serbina & Pamer, 2006). After their
extravasation into the mucosa, monocytes undergo differentiation through their
intermediates displaying a “waterfall” development. During the differentiation, Ly6Chigh
5 Introduction
monocytes, also known as population 1 (P1), lose Ly6C expression progressively to give rise to Ly6Cmid monocytes (P2), Ly6Clow monocytes, and Ly6C-CX3CR1+
macrophages. In the colon, this process takes approximately 5-6 days, whereas the
half-life of a mature macrophage is 6-8 weeks (Bain & Mowat, 2014; Zigmond & Jung,
2013). Most intestinal macrophages reside in the lamina propria in the vicinity of
epithelium, whereas rare populations of macrophages are present in the muscularis
externa and serosa where they interact with the enteric nervous system (Gabanyi et
al., 2016; Muller et al., 2014). Unlike the dendritic cells, macrophages in the intestine function poorly as antigen-presenting cells, and cannot migrate to mesenteric lymph nodes since they lack CCR7 (Schulz et al., 2009). Instead, they search the lumen for antigens by extending their processes between the epithelial cells (Niess et al., 2005;
Rossini et al., 2014). They can transfer the taken-up antigens to migratory dendritic cells, which is essential for establishment of oral tolerance (Mazzini et al., 2014).
Intestinal macrophages are specialized in recognizing and phagocytosing apoptotic cells or opsonized organisms (Bain et al., 2014). Accordingly, they are equipped with scavenger receptors, including CD163 (hemoglobin/haptoglobin scavenger receptor),
CD206 (mannose receptor), and scavenger receptor A (Joeris et al., 2017).
Macrophages produce cytokines such as TNF, IL1-b, and IL-10 in the steady-state intestine (Krause et al., 2015; Shaw et al., 2012; Zigmond & Jung, 2013). Importantly, intestinal macrophages exert their phagocytic and bactericidal activity without compromising intestinal homeostasis by triggering inflammation. They do so by negative regulation or inhibition of TLR signaling and activation of TGF-bR or IL-10R pathways to maintain hyporesponsiveness and suppression of inflammatory gene transcription (Maheshwari et al., 2011; Simon et al., 2016; Smythies et al., 2005). In
addition to hyporesponsiveness, macrophages help maintain the homeostasis also by
6 Introduction
promoting renewal and differentiation of intestinal epithelial cells via hepatocyte
growth factor and Wnt signaling (Cosin-Roger et al., 2016; D'Angelo et al., 2013).
Furthermore, IL-10 production in macrophages is crucial for establishing oral tolerance via expansion of induced regulatory T cells (iTreg) (Hadis et al., 2011). The depletion of macrophages causes slower recovery from experimental colitis in mice, indicating that the macrophages are essential for resolving inflammation and tissue repair
(Qualls et al., 2006). In line with this, the production of the cytokines IL-1b and IL-36 by myeloid cells drive epithelial turnover as a reparative mechanism after acute colitis
(Bersudsky et al., 2014; Scheibe et al., 2017). Macrophages also play a protective role in Citrobacter rodentium infection by secreting IL-23 and IL-1b, which activate ILC3s
to produce IL-22, a critical cytokine in pathogen clearance (Longman et al., 2014;
Manta et al., 2013).
Under inflammatory conditions, Ly6Chigh monocytes show enhanced recruitment into the mucosa and arrested differentiation into fully mature macrophages (Bain et al.,
2013). They produce pro-inflammatory factors such as IL-1b, TNF, IL-23, NO, and
reactive oxygen species, and promote Th1 and Th17 cells via IL-12 and IL-6 (Sanders
et al., 2017; Schreiber et al., 2013). Immune cell infiltrates in mice with colitis and
patients with IBD or infection also contain macrophages that are significant sources of
pro-inflammatory cytokines such as IL-6, IL-1b, and TNF, suggesting that they also
contribute to the inflammatory milieu (Bain & Mowat, 2014; MacDonald et al., 2011).
Dendritic cells are a subset of mononuclear phagocytes specializing in antigen-
presenting and therefore serve as the bridge between innate and adaptive immune
systems. In the intestine, they are required for the generation of Tregs for oral
tolerance and recognizing commensals in the steady-state and also for appropriate
immune reactions against pathogens (Stagg, 2018). There are two major subsets of
7 Introduction dendritic cells: conventional DCs (cDCs) and plasmacytoid DCs (pDCs). All DCs derive from a common hematopoietic progenitor, which depends on FLT3L signaling
(Chen et al., 2004). pDCs diverge from the cDCs and fully develop in the bone marrow prior to traveling to the thymus and secondary lymphoid organs via the bloodstream
(Cheong et al., 2010). pDCs produce type I and III IFN and show cytolytic activity
(Soumelis & Liu, 2006; Tel et al., 2012). Once activated, they can acquire an antigen- presenting DC state (Soumelis & Liu, 2006). Although they can effectively migrate into infection sites, they are less efficient in priming naïve T cells and migrating to the draining lymph nodes when compared to cDCs (GeurtsvanKessel et al., 2008;
O'Doherty et al., 1994). cDCs are found within the Peyer’s patches, isolated lymph follicles (ILFs), intestine- draining lymph nodes, and the lamina propria of the small and large intestine (Chirdo et al., 2005; Kelsall & Strober, 1996). Intestinal lamina propria cDCs are identified by their CD11c and MHCII, and lack of CD64 (Luda et al., 2016). They remain in the tissue for a few days and migrate to the draining lymph nodes via afferent lymph in a
CCR7-dependent manner (Jang et al., 2006). As the primary antigen-presenting cells, cDCs continually sample antigens by different mechanisms involving: i. Uptake of apoptotic cells, soluble antigens that cross the epithelium and MUC2- or IgA-coated bacterial antigens, ii. M or goblet cell-mediated antigen passage, and iii. Transfer of antigens from CX3CR1+ macrophages (Jang et al., 2004; Kadaoui & Corthesy, 2007;
Knoop et al., 2015; Knoop et al., 2013; Mazzini et al., 2014; McDole et al., 2012;
Rossini et al., 2014; Shan et al., 2013). Intestinal cDCs are further subdivided by their
CD103 and CD11b expression as three major subsets CD103+CD11b+,
CD103+CD11b-, CD103-CD11b+ and one minor subset CD103-CD11b-.
CD103+CD11b+ cDCs constitute the majority of cDCs in the small intestine, whereas
8 Introduction
CD103+CD11b- are the major subset in the colon (Houston et al., 2016). On the other hand, cDCs across tissues subcategorize as XCR1+ (cDC1) or SIRPa+ (cDC2)
(Guilliams et al., 2014). For their development, cDC1 require transcription factors
IRF8, BATF3, and ID2, whereas cDC2 are dependent on IRF4, NOTCH2, and KLF4
(Sichien et al., 2017). In the intestine, most of the XCR1+ DCs are CD103+CD11b- whereas SIRPa+ DCs can be CD103+CD11b+ and CD103-CD11b+. CD103-CD11b-
DCs are present in both subsets as a minor subpopulation (Joeris et al., 2017).
Intestinal cDC subsets in the intestine exert overlapping, therefore partially redundant functions through different T helper cell types, which will be further discussed in the context of metabolites.
Figure 1.2 Monocytes, macrophages and dendritic cells in the intestinal lamina propria
The newly extravasated Ly6Chigh monocytes constantly replenish tissue-resident macrophages in the intestine. Macrophages are specialized for recognizing luminal contents and eliminating the invading bacteria without generating an immune response. They can transfer the antigens to the dendritic cells,
9 Introduction
which egress to the MLN and initiate tolerance or immune reaction. The figure is adapted from (Joeris
et al., 2017).
1.1.2 Host- and microbiota-derived metabolites
The gut microbiota in the gastrointestinal tract facilitates not only digestion but also
the intestinal immunity (Hooper et al., 2001). Consequently, germ-free animals develop an impaired intestinal immune system with lower IgA levels in the lumen and improper GALT formation (Bauer et al., 1963; Macpherson & Uhr, 2004). Antibiotic treatment of adult mice also leads to reduced Treg, Th17, ILC populations and production of cytokines such as IL-10, IL-17 and IL-22 in the LP (Cording et al., 2013;
Ekmekciu et al., 2017; Ivanov et al., 2009; Kamada & Nunez, 2014). Studies with gnotobiotic mice indicate that segmented filamentous bacteria (SFBs) promote Th17 responses, whereas certain Clostridia and Bacteroides species drive Treg differentiation (Atarashi et al., 2013; Geva-Zatorsky et al., 2017; Tan et al., 2016).
Intestinal immunity can also shape the composition of the microbiota by enhancing or limiting the growth of different microbe species via immune mediators such as IgA, a- defensins, T-bet, TLR5 and NLRP6 (Elinav et al., 2011; Fransen et al., 2015; Garrett et al., 2007; Salzman et al., 2003; Vijay-Kumar et al., 2010). Microbial composition is also shaped by the interspecies competition or syntrophic interactions called metabolic cross-feeding of nutrients (Zelezniak et al., 2015). Likewise, the interaction between the intestinal immune system and the gut microbiota is mediated by bioactive metabolites.
Metabolites can be categorized as diet-dependent linked to digestion such as short- chain fatty acids or diet-independent synthesized only by gut microbiota such as lipopolysaccharides, ATP, and peptidoglycans. Diet-dependent metabolites are further classified as host- or microbiota-derived (G. Wang et al., 2019). Metabolites
10 Introduction
modulate differentiation, proliferation, maturation, and effector functions of various cell
types through signaling cascades and transcriptional regulations. They may act
intracellularly as histone deacetylase inhibitors or extracellularly via metabolite-
sensing G protein-coupled receptors (Rasoamanana et al., 2012).
Fatty acids comprise one of the most prominent metabolite groups in the intestine.
They are classified into short, medium, and long-chain fatty acids according to the
number of carbon atoms. Medium-chain fatty acids (MCFAs) are mainly absorbed
from the duodenum and contribute to cellular metabolism in the liver, whereas long-
chain fatty acids (LCFAs) are taken-up by enterocytes, esterified into complex lipids
that serve as cell membrane components (Melhem et al., 2019). Short-chain fatty
acids (SCFAs) are generated mostly in the colon by fermentation of non-digestible carbohydrates from fiber diet, including plant polysaccharides such as cellulose by the anaerobic bacteria (Flint et al., 2012). Major SCFAs include acetate, propionate, and butyrate, which are recognized by receptors PPARg, GPR41 and GPR43, and
GPR109a (Alex et al., 2013; Brown et al., 2003). Acetate and propionate are produced
mainly by Bacteroidetes, whereas most of the butyrate comes from Firmicutes phylum
(Hoverstad & Midtvedt, 1986; Macfarlane & Macfarlane, 2003). These SCFAs have a
pivotal role in the regulation of immune responses in steady-state and inflammation.
Accordingly, dietary fiber intake correlates with IgA levels due to SCFAs’ ability to promote B cell activation, differentiation, and antibody production via gene regulation and modulation of metabolism (M. Kim et al., 2016; Kudoh et al., 1998; Peterson et al., 2007). SCFAs can also modulate energy metabolism in IECs (Donohoe et al.,
2011). In addition to this, butyrate can suppress proliferation in enterocytes through negative regulators of cell cycle, thus limiting the access of butyrate to intestinal stem cell niche to protect the ISCs from the anti-proliferative effects (Kaiko et al., 2016).
11 Introduction
Butyrate also activates the NLRP3 inflammasome pathway and downstream
production of IL-18 promoting epithelial repair in IECs via its receptors GPR43 and
GPR109A (Levy et al., 2017; Levy et al., 2015). SCFAs inhibit the NF-kB pathway and
lead to decreased IL-12, TNF, and IFN-g production in DCs while activating anti-
inflammatory genes via GPR109A, resulting in DC-mediated induction of Tregs (Liu et al., 2012; Singh et al., 2014; Singh et al., 2010). Butyrate drives Treg differentiation, whereas acetate and propionate promote the expansion of preexisting Tregs (Arpaia et al., 2013).
Other prominent examples of immune regulator metabolites include tryptophan metabolites. Tryptophan can be used by both the host and the microbiota to generate active substances, most of which are ligands for aryl hydrocarbon receptor (AhR), a
ligand-dependent transcription factor (G. Wang et al., 2019). AhR is a crucial player in
ILF genesis, ILC3 expansion, and IL-22 production by ILCs contributing to protection against Citrobacter rodentium and Listeria monocytogenes (Kiss et al., 2011; Qiu et
al., 2012; Shi et al., 2007; Zelante et al., 2013). It also supports the epithelial barrier
and intraepithelial lymphocytes (IELs) and attenuates chemically-induced colitis (Y. Li et al., 2011). In addition to AhR, tryptophan metabolites can act through other receptors such as the pregnane X receptor (PXR) activated by indole, a tryptophan metabolite, leading to reinforcement of tight junctions and therefore improved epithelial barrier (Venkatesh et al., 2014).
Secondary bile acids are bacterial metabolites derived from primary bile acids of the
host metabolism that are transported into the intestinal lumen. Primary bile acids are
produced by cholesterol catabolism and are conjugated into taurine or glycine in the
liver prior to secretion into the intestine by the gall bladder (Postler & Ghosh, 2017).
While 95% of the bile acids are reabsorbed, they can get converted to secondary bile
12 Introduction
acids by the microbiota by deconjugation (Ridlon et al., 2006; Wahlstrom et al., 2016).
Deconjugated taurine enhances activation of the NLRP6 inflammasome in IECs, although NLRP6 prevents cell death in T cells in an inflammasome- and microbiota- independent manner (Levy et al., 2015; Radulovic et al., 2019). Additionally, secondary bile acids deoxycholic (DCA) and lithocholic (LCA) acids enhance epithelial barrier integrity by signaling through farnesoid X receptor (FXR) in IECs (H. Wang et al., 1999).
Retinoid acid (RA), a vitamin A-derived metabolite produced by aldehyde dehydrogenase (ALDH), is a prominent regulator of the intestinal immune system.
Together with TGF-b, RA, produced by cDCs, induces gut-homing molecules a4b7 and CCR9 on T cells in intestine-draining lymph nodes. Although initially thought as
CD103+ DC-specific, all LP cDCs migrate to the lymph nodes to induce Tregs, which
is central to the establishment of oral tolerance. Importantly, tolerance to small
intestinal antigens occurs in mesenteric lymph nodes mainly by CD103+ cDCs via RA
whereas colonic antigens are sampled by CD103-CD11b+ cDCs that migrate to iliac
nodes to induce Tregs independently of RA (Cerovic et al., 2013; Coombes et al.,
2007; Iwata et al., 2004; Johansson-Lindbom et al., 2005; Stagg et al., 2002;
Veenbergen et al., 2016; Welty et al., 2013). In addition to Treg expansion, RA also
contributes to switching from Th1/Th17 to Th2 immunity and enhances IgA production
by facilitating IgA class-switch recombination (Pantazi et al., 2015; Pino-Lagos et al.,
2011; Watanabe et al., 2010). Other vitamins mediating immune responses include vitamin D, important for T cell activation, and vitamin B and K, which the host relies on the microbiota for production (Hill, 1997; von Essen et al., 2010). Vitamin B9 (folic
acid) is involved in Treg survival, whereas vitamin B12 is crucial for NK cell activity
(Kinoshita et al., 2012; Tamura et al., 1999).
13 Introduction
Figure 1.3 Metabolite-sensing GPCRs on intestinal immune cells
Host- or microbial-derived metabolites can bind to their GPCRs (A) and therefore act as signaling molecules that lead to a variety of immune responses (B). The figure is adapted from (Melhem et al.,
2019).
1.2 Inflammatory bowel diseases
Inflammatory bowel disease (IBD) is a chronic relapsing inflammatory condition in the gastrointestinal tract. The main clinical entities of IBD include Crohn’s disease (CD) and ulcerative colitis (UC). CD is characterized by transmural and non-continuous inflammation in any segment of the gastrointestinal tract, whereas UC causes mucosal inflammation and is limited to the colon (Abraham & Cho, 2009). Disease activity of
IBD is diagnosed or assessed by symptoms such as stool consistency or bleeding, histology, endoscopy or radiography, and biomarkers such as serum C-reactive protein and fecal calprotectin (Walsh et al., 2016). It is a complex immune disorder caused by dysregulated immunity and interaction with the microbiota triggered by environmental factors and genetic susceptibility.
14 Introduction
1.2.1 Host immune system and genetics
CD and UC were initially classified as Th1/Th17- and Th2-mediated disorders,
respectively (M. F. Neurath, 2014). However, this classification is now considered as
an oversimplification and has been challenged by failed clinical trial outcomes, after
neutralizing IFN-g in CD patients, and IL-13 in UC patients proved ineffective (Danese
et al., 2015; Reinisch et al., 2006) and targeting IL-17 worsened CD (Hueber et al.,
2012).
IBD is a multigenic complex disease, and family history is a major risk factor for IBD occurrence. Genome-wide association studies have proven useful for providing insights into the critical molecular pathways involved in IBD pathogenesis. At least 99
non-overlapping risk loci associated with IBD have been identified to date. 28 out of
99 risk loci are shared between UC and CD, suggesting the existence of common
mechanistic features (Anderson et al., 2011; Franke et al., 2010). Analysis of the risk
loci revealed that genes implicated in IBD encode for proteins involved in pathways
that are crucial for processes including intestinal epithelial barrier functions, microbial
defense, metabolite-sensing, adaptive immunity, autophagy and reactive oxygen
species (ROS) production (Khor et al., 2011).
1.2.2 Microbiome
The variants associated with IBD can only explain 20-25% of all the cases, collectively.
Furthermore, the concordance rate in monozygotic twins is 10-15% in UC in
comparison to 30-35% in CD, suggesting that host genetics alone is not sufficient for
the pathogenesis. IBD patients have dysbiosis with reduced diversity of the gut
microbiota characterized by enrichment of Proteobacteria and reduction in Firmicutes
bacterial phyla (Ni et al., 2017) (Manichanh et al., 2012)., In addition, anatomical
15 Introduction
regions with fecal stasis and highest abundance of bacteria are relatively more prone
to inflammation in IBD further suggesting a role for the microbiota in the pathogenesis
of IBD (Janowitz et al., 1998). Nevertheless, the interaction between the microbiota
and IBD is dynamic rather than a causal relationship, and it remains unclear whether
dysbiosis is primary or secondary to the chronic inflammation. Regardless, the gut
microbiota is central to the IBD development and propagation and is shaped by
environmental and host factors (de Souza & Fiocchi, 2016). Approximately one-third
of the fecal microbial consortia is heritable, and 58 SNPs are associated with the
abundance of 33 taxa in healthy individuals (Turpin et al., 2016). Environmental factors that have an impact on the microbial composition will be discussed in the next section.
1.2.3 Diet and other environmental factors
Host genetics only partially determine the gut microbial composition, and external exposomes continue to dynamically alter the microbiota (Kahrstrom et al., 2016).
Additionally, the increasing incidence of IBD associated with the westernization of lifestyle and industrialization suggests that environmental factors are key contributors in the etiology of the disease (Rogler & Vavricka, 2015). Environmental factors affecting IBD predisposition include mode of birth, antibiotics use, breast milk, hypoxia due to high altitude, environmental pollution, nonsteroidal anti-inflammatory drugs
(NSAIDs), smoking and diet (Ananthakrishnan et al., 2018). Western diet with a high intake of red meat, sugar, high-fat and refined grains leads to elevated fecal calprotectin linking intestinal inflammation to dietary habits (Poullis et al., 2004). On the contrary, intake of dietary fiber, zinc, potassium, and vitamin D shows inverse association to risk of IBD (Ananthakrishnan et al., 2012; Ananthakrishnan et al., 2013;
Ananthakrishnan et al., 2015). Correlation between the diet and IBD risk can be in part
16 Introduction
explained by diet-mediated alteration in microbial composition. For instance, high
carbohydrate or caloric intake and a Mediterranean diet are associated with reduced
and increased microbiome diversity, respectively (Sokol et al., 2008; Zhernakova et
al., 2016).
Therefore, a dynamic and inter-changeable relationship between host genetics,
microbiota, and environmental factors determine the IBD predisposition and
prognosis.
Figure 1.4 Factors in IBD pathogenesis
IBD is a multifactorial disease driven by host genetics and environmental factors such as the diet and
the microbiota all of which are intertwined reciprocally. The figure is adapted from (Khalili et al., 2018).
1.2.4 Current therapy options
IBD is a progressive disease leading to complications such as stenosis, fistulas,
abscesses, and cancer (Baumgart & Sandborn, 2012; Danese & Fiocchi, 2011).
Clinical management of IBD patients includes anti-inflammatory drugs such as 5-
Aminasalicylates (5-ASAs) and systemic or topical delivery of corticosteroids. 5-ASAs can induce and maintain remission in UC by mechanisms including reducing prostaglandin synthesis, blocking neutrophil chemotaxis, and impairing NF-kB
(Allgayer, 2003). On the other hand, although capable of inducing remission in UC and
CD, corticosteroids are suboptimal for maintaining the remission response (Baumgart
& Sandborn, 2012; Danese & Fiocchi, 2011). Glucocorticoids bind to their receptor, and the receptor-glucocorticoid complexes translocate into the nucleus where they
17 Introduction regulate expression of genes or inhibit transcription factors such as NF-kB to suppress transcription of pro-inflammatory cytokines such as IL-1 or IL-6 (Oakley & Cidlowski,
2013; Rezaie et al., 2015).
In IBD, microbial strategies could be utilized to reestablish the gut microbial homeostasis. Antibiotics help eliminate pathogenic bacteria, whereas probiotics and prebiotics can facilitate replenishment of beneficial microorganisms. Fecal microbiota transplantation (FMT) is a relatively novel therapeutic strategy to replace the dysbiotic microbial composition in IBD patients. To date, FMT has been only used in clinical trials, and the efficacy is highly selective and donor-dependent, causing unpredictable outcomes (Aroniadis & Brandt, 2014). In addition to the microbial interventions, enteral nutritional therapy (ENT) has been used successively to correct dysbiosis in
IBD. Consequently, ENT has been associated with improvement in clinical symptoms, change in microbiota composition, and mucosal healing (Fell et al., 2000; Leach et al.,
2008).
Modulating the immune factors is the most commonly used strategy in IBD treatment.
Current biologic therapies include targeting of T cell trafficking by α4 subunit of α4β7 integrin, the p19 (IL-23) and p40 subunits (shared between IL-12 and IL-23), and
Janus kinases (JAK) to suppress cytokine signaling (M. Neurath, 2017). Among the biologic target therapies, anti-TNF agents were the first to be used in the clinic for the treatment of IBD patients. TNF is first synthesized as a transmembrane precursor protein. After cleavage, it becomes soluble TNF (sTNF), is released into the extracellular environment, and forms trimers (Black et al., 1997). Both membrane- bound and soluble TNF can activate TNF receptor 1 (TNFR1), whereas TNFR2 preferentially binds to the membrane-bound TNF (Grell et al., 1995). Various cell types produce TNF such as monocytes, macrophages, NK and T cells, fibroblast, and
18 Introduction
epithelial cells (Roulis et al., 2011). Depending on the TNFR composition, context, cell, or tissue type, TNF can exert diverse functions, including inducing survival, apoptosis, necroptosis, and pro-inflammatory or compensatory responses (Delgado & Brunner,
2019). In the context of IBD treatment, it is essential to note that full IgG1 monoclonal
anti-TNF antibodies such as infliximab and adalimumab are effective in the treatment
of both CD and UC while etanercept, which has a lower affinity for the membrane-
bound TNF showed no efficacy (Colombel et al., 2014; Colombel et al., 2010; Hanauer
et al., 2002; Sandborn et al., 2001). The co-stimulatory signal that TNFR2 activation provides in T cell survival might partially explain the difference in the efficacy (Van den
Brande et al., 2003). Overall, 30-50% of IBD patients are non-responsive to anti-TNF therapy, and anti-TNF agents may cause paradoxical inflammation such as psoriasis
(Cleynen & Vermeire, 2012; Korzenik et al., 2019; Niess & Danese, 2014).
Interestingly, anti-TNF therapy, particularly etanercept, increased IBD susceptibility in rheumatoid arthritis patients (Korzenik et al., 2019).
1.3 G protein-coupled receptor (GPR) 35
GPR35, a class A, rhodopsin-like GPCR with seven transmembrane domains, was
first discovered by a human genomic DNA screen (O'Dowd et al., 1998). In humans,
GPR35 is localized in chromosome 2q37.3 and is transcribed into two alternatively
spliced variants. GPR35a encodes an open reading frame (ORF) of 309 amino acids,
whereas GPR35b has an N-terminal extension of 31 amino acids (Okumura et al.,
2004). Nevertheless, GPR35a and GPR35b exert similar responses in in vitro assays,
and the function of the N-terminal extension is still not known (Guo et al., 2008).
GPR35 has the closest homology to a cannabinoid receptor GPR55, lysophosphatidic
19 Introduction acid receptors, LPAR4, 5 and 6, and the nicotinic acid receptor HM74 (Fredriksson et al., 2003; Vassilatis et al., 2003).
In humans, GPR35 is highly expressed in the pancreas, small intestine, colon, spleen, and immune cells including monocytes, and dendritic cells (Taniguchi et al., 2006; J.
Wang et al., 2006). It is also expressed to a lower extent in the stomach, adipose tissue, and liver. Northern blot analysis has shown prominent expression in the rat intestine, and similarly, high expression levels were observed in the spleen, small intestine, colon, and stomach in both mice and rats (Ohshiro et al., 2008; Taniguchi et al., 2006; J. Wang et al., 2006).
GPR35 remains an orphan GPCR, although kynurenic acid, CXCL17, and lysophosphatidic acid have been proposed as endogenous ligands. Kynurenic acid, a tryptophan derived metabolite, was the first identified agonist of GPR35 that is found endogenously in various tissues such as the brain, pancreas, intestine, lungs, and spleen (Mackenzie et al., 2011). Kynurenic acid is generated by the kynurenine pathway and is known for its neuroprotective properties by acting as an antagonist of
N-Methyl-D-aspartic acid (NMDA) receptor, which is a glutamate receptor (Stone,
2001). KYNA has a strong species selectivity for GPR35 activation with EC50s 40 to
100-fold higher in humans than rats (Barth et al., 2009; Jenkins et al., 2011).
Therefore, it is critical to assess whether the concentrations of kynurenic acid required for GPR35 activation are comparable to endogenous levels in vivo. The second proposed endogenous ligand for GPR35 is LPA (Oka et al., 2010). LPA is a naturally occurring phospholipid derivate consisted of a glycerol backbone, an acyl chain, and a phosphate group (M. E. Lin et al., 2010). It is a component of the cell membrane, but also can act as a signaling molecule when released into the extracellular environment (Ye & Chun, 2010). It is known to signal through six GPCRs, LPAR1-
20 Introduction
LPAR6, and is implicated in cancer types, including colorectal cancer (Yun, 2019).
LPA was able to induce an increase in Ca2+ response in GPR35-transfected cells, and
also the internalization of the GPR35. However, there have been no follow-up studies on the interaction between GPR35 and LPA to date. Lastly, it has been reported that
CXCL17, a chemokine for alveolar macrophages, can induce calcium flux in GPR35- transfected cells in nanomolar concentrations and that GPR35-expressing human monocytic THP-1 cells migrate towards CXCL17 (Burkhardt et al., 2014; Maravillas-
Montero et al., 2015). However, another study has shown that human peripheral blood monocytes, which are also GPR35+, do not migrate to CXCL17, making it a
controversial GPR35 ligand candidate (Binti Mohd Amir et al., 2018). Notably, the
ligand identification studies revealed that GPR35 transmits function via interaction with
Gαi/o, Gα13, and also β-arrestin (Mackenzie & Milligan, 2017).
In the context of IBD, a GWAS has revealed a GPR35 SNP, rs4676410, associated
with early-onset UC that is an upstream intron variant resulting in a cytosine to thymine
substitution (Imielinski et al., 2009). Following, another GWAS has identified 2 SNPs in GPR35 linked to increased risk of IBD: rs3749171 and the previously identified rs4676410 (Ellinghaus et al., 2013).
In a recent study, Schneditz et al. have shown that the protein-coding variant
rs3749171 in GPR35 is hypermorphic and that GPR35 expression in intestinal
epithelial cells is oncogenic in spontaneous and inflammation-induced colorectal cancer mouse models (Schneditz et al., 2019). Consistently, GPR35 promotes epithelial cell turn over in the wound healing mouse model, suggesting that GPR35
might regulate epithelial cell renewal (Tsukahara et al., 2017). Ablation of GPR35
leads to exacerbated chemically induced colitis in mice (Farooq et al., 2018). However,
21 Introduction the cell types that govern the protective roles of GPR35 in colitis and putative endogenous ligands mediating them remain unknown.
1.4 Aims of the study
GPR35 is expressed by bone marrow-derived macrophages (BMDMs) and peritoneal macrophages (Lattin et al., 2008; Schneditz et al., 2019). In BMDMs, GPR35 regulates the metabolism by interacting with the Na/K-ATPase (Schneditz et al., 2019).
However, whether GPR35 has a crucial role in modulating intestinal macrophage responses to potential endogenous ligand(s) during inflammation is yet to be covered.
To answer this, in this Ph.D. thesis, we aim:
1. To characterize GPR35 expression in the intestine, particularly in the
macrophages under steady-state and inflammatory conditions.
2. To oversee the interspecies differences in GPR35 activation by combining
transgenic zebrafish and mouse models for determining the GPR35-mediated
cellular events led by the putative endogenous ligand(s) in macrophages.
3. To investigate a role for macrophage-expressed GPR35 in cytokine responses
during chemically-induced colonic inflammation.
22 Methods
2 METHODS
2.1 Human samples
For immunofluorescence staining, biopsies from 4 ulcerative colitis and 3 Crohn’s disease patients from the Basel IBD cohort were included (ethics protocol EKBB
139/13 (PB 2016.02242; Ethics Committee for Northwest and Central Switzerland
(EKNZ)). The biopsies were collected from inflamed or non-inflamed intestinal tissues of the same patients during Ileocolonoscopy and were embedded in optimal cutting temperature (OCT) compound (TissueTek). The biopsies were kept at -800C. Patient characteristics are listed in Table 2.1.
Table 2.1 Patient Characteristics of Basel IBD Cohort for Biopsies Obtained for
Immunofluorescence Staining
Location Treatment Patient Age at Smoking Location Gender Age BMI non- at time of DAI ID diagnosis status inflamed inflamed study
Ulcerative colitis
sigma/rec 504 F 50 25.6 34 Unk trans. colon none 5 tum
rectum/si 535 F 71 27.3 56 Unk colon none 6 gmoid
551 F 53 25 37 Unk rectum Unk Unk Unk
None, sigmoid/r ascend. / 619 Unk Unk Unk Unk Unk Salofalk 10 Unk ectum trans. colon days before
Crohn’s disease
non- Quantalan, 558 F 72 19 23 Unk rectum Unk smoker Immodium
Spiricort, 568 M 68 32 52 active sigmoid desc. colon Aldactone, 74 Orfiril
term. ascend. 620 F 69 21.6 56 active Unk 70 Ileum colon
Unk, unknown; DAI, disease activity index
23 Methods
Baseline group characteristics for the IBD patients included in the Swiss IBD Cohort
Study examined for anti-TNF therapy responses are summarized in Table 2.2
(SwissIBD cohort project 2017-13).
Table 2.2 Group characteristics of IBD patients analyzed for the GPR35T108M SNP variant and TNF blocker therapy response
Baseline Group characteristics Crohn’s disease (n=63) Ulcerative colitis (n=28) C (n=21) CT T (n=41) C(n=9) CT T (n=16) (n=1) (n=0) Gender, male/female, n (%) 12 0 (0) 20 6 (66.7) 0 10 (62.5) (57.14) /1 (100) (48.78) /3 (33.3) /6 (37.5) /9 /21 (42.86) (51.22) Median age (range), yr 39 (14- 54 (54) 38 (17- 42 (28- 0 36.5 (23- 63) 80) 68) 72) Mean BMI (SD), kg/m2 22.7 23.5 (.) 23.5 (4.1) 25.7 0 22.6 (3.4) (1.51) (5.24) Median age at diagnosis (range), yr 22 (6-52) 30 (30) 21 (7-51) 25 (15- 0 24.5 (15- 59) 64) Median disease duration (range), yr 13 (3-26) 23 (23) 15 (3-41) 13 (9-24) 0 9.5 (4-24) CD extent, n (%) Ileum isolated 1 (4.76) 0 (0) 4 (9.76) Colon isolated 6 (28.57) 0 (0) 9 (21.95) Ileocolonic 1 (4.76) 1 (100) 8 (19.51) L4 0 (0) 0 (0) 1 (2.44) Unknown 11 0 (0) 16 (39) (52.38) UC extent, n (%) Proctitis 1 (11.1) 0 2 (15.4) Left-sided colitis 4 (44.4) 0 3 (23.1) Pancolitis 3 (33.3) 0 4 (30.8) Unknown 1 (11.1) 0 4 (30.8) Current medical treatment, n (%) No treatment 1 (4.76) 0 (0) 9 (21.95) 1 (11.1) 0 3 (18.75) 5-ASA 2 (9.52) 0 (0) 8 (19.51) 3 (33.3) 0 12 (75) Steroids 11 0 (0) 10 3 (33.3) 0 5 (31.3) (52.38) (24.39) Immunosuppressants 9 (42.86) 0 (0) 11 3 (33.3) 0 3 (18.75) (26.83) Anti-TNF 8 (38.1) 1 (100) 19 1 (11.1) 0 2 (12.5) (46.34) Antibiotics 2 (9.52) 0 (0) 1 (2.44) 1 (11.1) 0 0 (0) Other 9 (42.86) 0 (0) 9 (21.95) 6 (66.6) 0 7 (43.75) Smoking status 17 1 (11.1) 0 3 (18.8) Active smoker, n (%) 7 (33.3) 1 (100) (41.46) 20 7 (77.8) 0 12 (75) Non-smoker, n (%) 13 (61.9) 0 (0) (41.46) Unknown 1 (4.76) 0 (0) 4 (9.76) 1 (11.1) 0 1 (6.2)
24 Methods
2.2 Animals
2.2.1 Mouse lines
C57BL/6, Cx3cr1-GFP (B6.129P-Cx3cr1tm1Litt/J) and Cx3cr1CreER (B6.129P2(Cg)-
Cx3cr1tm2.1(cre/ERT2)Litt/WganJ) mouse strains were bred in the animal facilities of
Department of Biomedicine, University of Basel, Switzerland or the Karolinska
Institutet, Solna, Sweden. Gpr35-tdTomato, Gpr35-/- and Gpr35flox/flox mouse lines
were generated as described below. Gpr35-tdTomato mice were crossed with Cx3cr1-
GFP mice to obtain double reporter mice, and Gpr35flox/flox were crossed with
Cx3cr1CreER to obtain tamoxifen inducible Gpr35ΔCX3CR1 mice. All animals were
maintained under specific pathogen-free (SPF) conditions. Germ-free C57BL/6
animals were used from the Core Facility for Germ-Free Research at the Karolinska
Institutet, Solna, Sweden. At least three 6-12-week-old mice per group were included in the experimental cohorts by randomization. All experiments were conducted in adherence to the Swiss Federal and Cantonal regulations (animal protocol number
2832 (canton Basel-Stadt)) and the Stockholm regional ethics committee (ethical number N89-15).
2.2.2 Generation of Gpr35-IRES-tdTomato knock-in mice
Gpr35-IRES-tdTomato reporter mouse line was constructed under the genetic background C57BL/6J by Beijing Biocytogen (Beijing, China) by introducing IRES- tdTomato between the protein coding sequences and 3'UTR of the Gpr35 gene. The targeting vector consisted of a 4.7-kb left homology arm spanning exon 1 and an FRT- flanked neo cassette inserted 352 base pair (bp) upstream of exon 2; an internal ribosome entry site 2 (IRES2) sequence for initiating translation; and tdTomato reporter and 3.9-kb right homology arm inserted downstream of the stop codon. The
25 Methods
sequence of the constructed targeting vector was confirmed by sequencing analysis.
The linearized targeting vector was transfected into C57BL/6J embryonic stem (ES)
cells by electroporation. Eight positive ES clones identified by Southern blot using 5’
and 3’ probes were injected into BALB/c blastocysts followed by implantation into
pseudo pregnant females. Four chimeric male mice were then crossed with FLP
recombinase female mice to remove the Frt-flanked neo selection cassette from the recombined allele of the obtained F1 offspring. The removal of the neo cassette in the
F1 mice was verified by PCR using the primers Frt-F2 and Frt-R2. The genotyping of the reporter animals was done by PCR using the primers listed in Table 6.1. Following
PCR cycling parameters were used with 35 cycles of amplification: denaturation 950C
for 2 min; amplification 950C 30 sec, 620C 30 sec, 720C 25 sec; final elongation 720C
10 min.
Figure 2.1 Construct of Gpr35-IRES-tdTomato and PCR products from the genotyping
(A) Construct scheme for Gpr35-tdTomato knock-in. UTR (untranscribed region), E (exon), DTA
(diphtheria toxin), tdT (tdTomato)
(B) PCR products from the genotyping protocol for the Gpr35-tdTomato mice. Product sizes: 385 bp
(mutant), 244 bp (WT).
26 Methods
2.2.3 Construction of Gpr35-flox and -knockout (KO) mice
Gpr35flox and Gpr35-/- mice were constructed by Beijing Biocytogen (Beijing, China)
using the CRISPR/Cas9 genome editing system.
For the deletion of Gpr35 to obtain Gpr35-/- mice, two sgRNAs were designed to target
the region spanning upstream of exon 2 and downstream of 3’UTR by the CRISPR
design tool (http://www.sanger.ac.uk/htgt/wge/). Candidate sgRNAs were tested for
on-target activity using the UCA kit (Z. Lin et al., 2016) followed by transcription of
Cas9 mRNA and sgRNAs by T7 RNA polymerase. The T7 promoter sequence was
then incorporated into the Cas9 and sgRNA templates by PCR. Obtained PCR
products, T7-Cas9 and T7-sgRNA, were purified from the gel and used for in vitro
transcription with the MEGAshortscript T7 kit (Life Technologies). Cas9 mRNA and
sgRNAs were purified using the MEGAclear kit. The same genome editing strategy
was used to generate Gpr35-flox constructs. The targeting construct of Gpr35-flox included 1.3 kb of homologous genomic sequence upstream of exon 2 and downstream of 3’UTR flanked by two loxP sites.
After the production of Cas9 mRNA and sgRNAs, the donor vector was isolated using an endotoxin-free plasmid DNA kit. C57BL/6N females were chosen as embryo donors and pseudo pregnant mothers. Super-ovulated C57BL/6N mice (3-4 weeks
old) mated males of the same genetic background, and fertilized embryos were taken
from the ampullae. Cas9 mRNA, sgRNAs, and donor vector were co-injected into the
fertilized eggs at the one-cell stage. The surviving zygotes were then transferred into
the oviducts of pseudo pregnant females.
The genotyping of Gpr35-deficient animals was done by PCR in 2 different reactions
using the listed primers (Table. 6.1) under the following conditions: initial denaturation
27 Methods at 950C for 3 min; 35 cycles of denaturation 950C 30 sec, annealing 640C 30 sec, elongation 720C 45 sec; and final elongation 720C 10 min.
Figure 2.2 Construct of Gpr35-knock out and PCR of the genotyping
(A) Construct scheme for Gpr35-knock-out. UTR (untranscribed region), E (exon)
(B) PCR products from the genotyping protocol for the Gpr35-deficient mice. M: mouse. Product sizes:
First reaction: 387 bp (WT), Second reaction: 387 bp (mutant).
The Gpr35-flox mice were genotyped by PCR (for primers see Table 6.1) by denaturation at 950C for 3 min, amplifying 35 cycles at 950C 30 sec, 620C 30 sec, 720C
35 sec and elongating at 720C for 10 min.
28 Methods
Figure 2.3 Construct of Gpr35-flox and PCR of the genotyping
(A) Construct scheme for Gpr35-flox. UTR (untranscribed region), E (exon)
(B) PCR products from the genotyping for the Gpr35-flox mice. Product sizes: First reaction: 431 bp
(WT), 516 bp (mutant); Second reaction: 387 bp (WT), 473 bp (mutant).
2.2.4 Zebrafish lines
The Tg(mpeg1:mCherry) macrophage reporter line was kindly given by Professor
Georges Luftalla (Montpellier, France). The identified zebrafish G-protein coupled receptor 35-like gene (LOC101882856) (mRNA sequence ID: XM_021466387.1, previous Ensembl ID: ENSDARG00000075877, current Ensembl ID:
ENSDARG00000113303) was modified by the CRISPR-Cas9 system by the Genome
Engineering Zebrafish, Science for Life Laboratory (SciLifeLab), Uppsala, Sweden.
The gRNA was designed to target a region within the exon 2 in the reverse strand and was followed by a protospacer adjacent motif (PAM), (5’ GGT AGG CCA CAC GCT
CAA ACA GG 3’ – PAM sequence is underlined). Eggs from WT AB strain were co-
injected with a mix of Cas9 mRNA and sgRNA at the single-cell stage. Founder
screening was performed by somatic activity test (CRISPR-STAT), and germline
29 Methods
transmission was tested using fluorescence PCR as previously described (M. Li et al.,
2016). Adult injection groups with somatic activity underwent founder screening and
positive founders (F0) were in-crossed to assess germline transmission in F1 embryos.
F1 embryos were fin-clipped for genotyping by fluorescence PCR. The mutation was
further verified by Sanger sequencing. For maintaining, embryos were kept and grown
in circulating, filtered, and 28.50C water. All experiments were done in compliance with
the Uppsala University Ethical Committee for Animal Research (C161.14) and
Karolinska Institutet Ethical Committee for Animal Research (N5756/17).
Figure 2.4 Construction of gpr35buu1892 mutant zebrafish
(A) Scheme of the gpr35b locus and the targeting strategy to generate gpr35buu1892 mutant zebrafish
line.
(B) Resulting gpr35b protein sequences from WT and gpr35buu1892 mutant zebrafish that was
introduced a preliminary stop codon.
30 Methods
2.3 In vivo experiments
2.3.1 Dextran sodium sulfate-induced colitis mouse model
6 to 12-week-old female weight-matched mice were randomly assigned in
experimental cohorts randomly and treated with 1.5-2.5% DSS (MP Biomedicals) in the drinking water for five days. After, mice were given normal drinking water for two days. Animals were monitored daily for colitis score and weight change. Disease activity indexes were calculated using the criteria (Steinert et al., 2017): rectal bleeding: 0 - absent, 1 - bleeding; stool: 0 - normal, 1 - loose stool, 2 - diarrhea; behavior: 0 - normal movement, 1 - reluctant to move, 2 - hunched back; fur: 0 - normal, 1 - ruffled, 2 - spiky; weight loss: 0 – no loss, 1 - weight loss 0-5%, 2 - weight loss >5 - 10%, 3 - weight loss > 10 - 15%, 4 - weight loss > 15%. The animals were sacrificed on day 7 or 8 or when they met the ending criteria: total score of ≥ 6, > 15
% body weight loss, excessive bleeding, or rectal prolapse.
2.3.2 Mouse endoscopy
To determine the colitis severity macroscopically, mice were first anesthetized with an intraperitoneal injection of 100 mg/kg body weight ketamine and 8 mg/kg body weight.
The distal 3 cm of the colon and the rectum were imaged with a Karl Storz Tele Pack
Pal 20043020 (Karl Storz Endoskope, Tuttlingen, Germany) as previously described
(Melhem et al., 2017).
2.3.3 Treatment of zebrafish with 2,4,6-Trinitrobenzenesulfonic acid
Zebrafish larvae were either left untreated or were treated with 2,4,6-
Trinitrobenzenesulfonic acid (TNBS; Sigma Aldrich) between 72 hours post-
31 Methods
fertilization (hpf) and 120 hpf. TNBS was added in the embryo (E3) water at a final concentration of 50 µg/mL, and the TNBS-water was exchanged daily.
2.3.4 Treatment of zebrafish with antibiotics
Zebrafish larvae were treated with antibiotics Ampicillin (100 µg/ml) and Kanamycin (5
µg/ml) between 72 hpf and 120 hpf (Bates et al., 2006). The antibiotics cocktail was added to the E3 water and was exchanged daily.
2.3.5 Treatment of mice with antibiotics
WT mice were treated with antibiotics Ampicillin (1mg/ml), Kanamycin (1mg/ml),
Gentamicin (1mg/ml), Metronidazole (1mg/ml), Neomycin (1mg/ml), and Vancomycin
(0.5mg/ml) daily for 10 days by oral gavage (Reikvam et al., 2011).
2.3.6 Challenging of mice with E. coli-CFP
E. coli DH10B pCFP-OVA was constructed and propagated as described previously
(Rossini et al., 2014). Gpr35-tdTomato x Cx3cr1-GFP double reporter mice were gavaged with 1x108 CFUs of CFP-OVA+ E. coli resuspended in PBS every other day
for 21 days. Control mice were gavaged with only PBS.
2.3.7 Exposure of zebrafish with Vibrio anguillarum
V. anguillarum strain 1669 was incubated in the tryptic soy broth medium. Bacterial
culture was set to OD600 (optical density at 600 nm) value of 1.5 and pelleted (9 ml of
full-grown culture). The bacteria were resuspended in NaCl (9 g/L), 0.35%
formaldehyde, and incubated overnight at 20˚C. The bacterial suspension was washed four times in NaCl (9 g/L) and resuspended in 800 ml of the same solution. V.
32 Methods
anguillarum bacterial extract was mixed in a 1:1 ratio in phenol red (Sigma Aldrich).
One µL of this solution was diluted in two µL PBS, and two nL of the resulting solution
was injected in the intestinal lumen of zebrafish larvae at 110 hpf. Larvae were
anesthetized using 0.0016% Tricaine MS0222 (Sigma Aldrich), and analyzed 6 h post-
injection.
2.3.9 LPA treatment of zebrafish larvae for cytokine expression analysis
WT or gpr35buu1892 mutant zebrafish larvae were either left untreated or treated with
10 μM LPA (Sigma Aldrich) in water between 96-120 hpf. Following the incubation,
zebrafish larvae were collected for RNA isolation.
2.3.8 In vivo macrophage migration assay in zebrafish
LPA (10 µM) mixed with FITC-Dextran (500 µg/ml) in PBS was prepared, and for
control fish, PBS was used. 2 nl of corresponding solutions were injected in the otic
vesicle of Tg(mpeg1:mCherry)WT and Tg(mpeg1:mCherry)gpr35buu1892 macrophage
reporter zebrafish larvae anesthetized with 0.0016% Tricaine MS-222 at 110 hpf.
Macrophage recruitment was imaged 6 h after the injection.
2.4 Histology and imaging
2.4.1 Preparation of RNA probes for Gpr35b in zebrafish
DNA plasmid with Gpr35b cDNA (5 µg) was linearized using the restriction enzymes
SacI and EcoRI to obtain the sense and anti-sense probe templates, respectively. The
linearized plasmid was isolated by the conventional phenol: chloroform method. In
vitro synthesis of the sense and anti-sense DIG-Labeled RNA probes was done by incubating at 37 °C for 2 hours with the transcription mix (20 µL) containing the
33 Methods
template (1-2 µg), DIG-RNA labeling mix, RNase Inhibitor, buffer and RNA
Polymerase T7 and T3, respectively. The DNA template was digested with DNase I
for 30 min at 37 °C and the digestion was inactivated by 2 μl of 0.2 M EDTA. DIG-
Labeled RNA probes were precipitated by the LiCl method followed by resuspension
in 30 μl probe solution consisted of 19 μl sterilized water, 10 μl RNAlater and 1 μl 0.5M
EDTA.
2.4.2 In situ hybridization for gpr35b in zebrafish
In situ hybridization (ISH) was performed on whole zebrafish larvae at 72, 96, and 120
hpf. Larvae were fixed with 4% PFA at 4 °C for 16 hours and PFA residue was removed
by washing three times with PBS. Fixed larvae were dehydrated by incubation in 25%,
50% and 75% methanol 100% methanol for 5 min and a final 15-min immerse in 100% methanol. Prior to dehydration, depigmentation was done if necessary. For this, larvae were kept in 3% H2O2/0.5% KOH at room temperature (RT) until pigmentation disappeared. Larvae were kept at -20 °C for 2 hours followed by rehydration, washing four times with PBST (0.1% Tween-20 in PBS) and proteinase K (10 µg/mL) treatment at RT. Duration of the proteinase K digestion varied according to the developmental stage e.g. 72 hpf – 20 min; 96 hpf – 30 min; or 120 hpf – 40 min. The larvae were incubated in 4% PFA for 20 min, washed with PBST and prehybridized for 5 h at 70
°C with hybridization mix (HM) solution containing 50% deionized formamide
(Millipore); 0.1% Tween-20 (Sigma Aldrich); 5X saline sodium citrate solution (Merck);
50mg/ML of heparin (Sigma Aldrich); 500 mg/mL RNase-free tRNA (Sigma Aldrich).
The solution was then exchanged by HM with 50 ng of antisense/sense DIG-labeled
RNA probe. The larvae were incubated overnight at 70 °C and washed with SSC and
PBST solutions. Blocking was done with blocking buffer for 4h at RT prior to addition
34 Methods
of anti-DIG-AP antibody solution. Samples were kept in the antibody solution overnight
at 4 °C and were washed 6 times for 15 min on a horizontal shaker. Following
incubation with alkaline Tris buffer for 5 min at RT with, the staining solution was
added. The reaction was stopped by stop solution (1mM EDTA and 0.1% Tween-20 in PBS pH 5.5). Stained larvae were transferred to a tube with 100% glycerol and kept for 24 hours before mounting.
2.4.3 GPR35 staining on human biopsies and mouse tissues
Human biopsies embedded in cryoblocks were provided by the Basel IBD cohort.
Extracted mouse tissues were fixed with 4% PFA and kept in 30% sucrose for 16 hours for cryo-embedding. For paraffin embedding, mouse tissue pieces were dehydrated progressively in ethanol solutions.
All tissues were cut into at 5-6 µm- thick sections and fixed with 4% PFA before staining. For paraffin-sections, antigen retrieval to unmask the antigenic epitope was done using the citric buffer in a microwave at 1200C for 15 minutes. PBS containing
0.4% Triton X-100 (cryo-sections) or 0.1% Tween20 (paraffin sections) containing 5% goat serum were used for blocking of unspecific antibody binding and permeabilization
(all Sigma Aldrich).
GPR35 immunostaining was done using rabbit polyclonal anti-human/mouse GPR35 primary and goat anti-rabbit IgG secondary antibodies. NucBlueTM Live Cell Stain
(Thermo Fisher) was used for nuclear staining. Samples were imaged by a Nikon A1R
confocal microscope.
35 Methods
2.4.4 Hematoxylin-eosin (H&E) staining and histological scoring
Tissue sections from the distal colon of mice were fixed in 4% PFA and tissue were embedded in paraffin after dehydration. H&E staining was done on 5 µm paraffin sections from the distal colon for histological scoring of colitis in mouse samples.
Colitis severity was assessed using the following criteria: tissue architecture (0: normal, 1-3: mild-extensive damage); immune cell infiltration (0: normal, 1-3: mild- transmural); goblet cell loss (0:no, 1: yes); crypt abscesses (0:no, 1: yes); muscle thickening (0: normal, 1-3: mild-extensive). Tissues were scored in a blinded fashion for preventing bias.
2.4.5 Autotaxin staining of mouse colon
Mouse tissues were fixed using 4% PFA, dehydrated in ethanol solutions, embedded in paraffin, and sectioned between 5-6 µm. Endogenous peroxidase was blocked using 3% H2O2 solution prepared in methanol, and antigen retrieval was done in EDTA buffer (1mM EDTA, pH 8.0). Blocking was achieved by BLOXALL™ Blocking Solution
(Vector Laboratories). Staining was done using mouse monoclonal anti-mouse/human
ENPP2 (autotaxin) primary (Abcam) and goat anti-rabbit IgG secondary antibodies.
Counterstaining was done by H&E staining.
2.4.6 Ex vivo imaging of mouse tissues
Tissues were extracted, and intestinal segments were washed by flushing with PBS.
All tissues were placed on microscopy slides in desired positions, and intestinal segments were opened longitudinally. Tissues were kept in a drop of PBS to prevent drying and were covered with a coverslip sealed with nail polish. Freshly extracted tissues were then imaged using a Nikon A1R confocal microscope.
36 Methods
2.5 Cell isolation
Intestinal lamina propria cells were isolated as described before (Steinert et al., 2017).
Briefly, small or large intestines were opened longitudinally and washed with PBS
(Sigma Aldrich). IECs were dissociated by incubation in 5 mM EDTA at 37 °C in a
shaking water bath at 200 rpm for 10 minutes. The dissociation was done by repeating
the incubation 3 times with vortexing for 30 sec in between. IECs were collected for
further processing, if needed. The tissues were washed in PBS to get rid of the EDTA
residue and cut into small pieces. Digestion of the tissues was done using Roswell
Park Memorial Institute (RPMI) 1640 (Sigma Aldrich) medium with 0.5 mg/ml
Collagenase type VIII (Sigma Aldrich) and 10 U/mL DNase (Roche) for 15-20 min at
37 °C in shaking water bath with 30 sec vortexing every 5 min. Single cell suspension
was obtained using a 70 µm cell strainer. Cells were pelleted for further use.
Splenocytes and MLN cells were extracted by pressing the tissue with a syringe
plunger against a 70 µm cell strainer. Erythrocyte lysis was done for solenocytes by
incubating in ammonium-chloride-potassium buffer (150 mM NH4Cl, 10 mM KHCO3,
0.1 mM 0.5 M EDTA) for 10 minutes at RT. Cells were pelleted for further use.
2.6 Antibodies, cell staining, and flow cytometry
After centrifugation of single-cell suspensions, cells were resuspended in PBS and
counted using trypan blue and a hemocytometer. Up to 5x106 cells were incubated
with anti-CD16/CD32 (Fc receptor) clone 93 (Invitrogen) for blocking and with fixable viability dye eFluor455UV (eBioscience) for live/dead cell discrimination for 30 min at
4°C. Cells were stained for surface antigens for 20 min at 4 °C and washed using PBS
containing 2% Fecal Bovine Serum (FBS), 0.1% sodium azide, and 10 mM EDTA
(FACS buffer). For intracellular staining, cells were fixed and permeabilized in
37 Methods
Cytofix/Cytoperm solution for 30 minutes at 4 °C according to the manufacturer’s
instructions (BD Biosciences). Additional incubation with antibodies against
intracellular antigens was done for 20 min at 4 °C. At the end of the surface or
intracellular antigen staining, cells were fixed in 4% PFA for 10-15 minutes at RT and
resuspended in FACS buffer until further analysis. Flow cytometric analysis was
performed using a Fortessa flow cytometer (BD Biosciences). Data were analyzed by
FlowJo software version 10.0.7r2 (TreeStar). For gating, doublet discrimination was
done on forward scatter (FSC-H) versus FSC-A plot.
For mononuclear phagocyte analyses, cells were stained using eVolve655-conjugated anti-CD45 clone 30-F11 (eBioscience), biotin-labeled anti- CD3 clone 145-2C11, anti-
CD19 clone 6D5, and anti-NK.1.1 clone PK136, AF700-conjugated anti- I-A/I-E
(MHCII) clone M5/114.15.2, PE/Cy7-conjugated anti- CD64 clone X54-5/7.1,
APC/Cy7-conjugated anti-CD11c clone N418, FITC-conjugated anti- CD11b clone
M1/70, PerCP/Cy5.5- conjugated anti-Ly6C clone HK1.4, and APC-conjugated anti-
Ly6G clone 1A8 (all BioLegend). For lineage exclusion, CD3+, CD19+, and NK1.1+
cells were gated out. eFluor450 conjugated Streptavidin (eBioscience) was used for
biotin-labeled antibodies.
For lymphocyte staining, antibodies for APC/Cy7-conjugated anti-CD45 clone 30-F11,
AF700- conjugated anti-CD3 clone 17A2, BV785-conjugated anti-CD19 clone 6D5,
BV510-conjugated anti-CD4 clone RM4-5 and PerCP-conjugated anti-CD8a clone 53-
6.7 or Biotin-labeled anti-CD8 clone 53-6.7 (all BioLegend) were used.
For innate lymphoid cell panel, antibodies APC/Cy7- conjugated anti-CD90.2 clone
30-H12 (BioLegend), APC-conjugated anti-GATA3 clone 16E10A23 (BioLegend),
PerCP/Cy5.5-conjugated anti-RORgT clone Q31-378 (BD Biosciences), PE/Cy7-
conjugated anti-T-bet clone 4B10 (BioLegend) and FITC-conjugated anti-Eomes clone
38 Methods
WD1928 (Invitrogen) were included whereas biotin-conjugated antibodies anti-CD3
145-2C11, anti-CD8a 53-6.7, anti-CD19 6D5, anti-CD11c N418 (all BioLegend), anti-
B220 RA3-6B2 (BD Biosciences), anti-Gr-1 RB6-8C5, anti-TCRb H57-597, anti-
TCRgd GL3 and anti-Ter119 TER-119 (all BioLegend) were used for lineage exclusion.
Pe/Cy5 conjugated Streptavidin (BioLegend) was used for biotin-labeled antibodies.
Table 2.3 Mouse mononuclear phagocytes panel
Fluorochrome Viability dye: eFluor455UV
extracellular Fluorochrome Clone Anti-mouse CD45 eVolve655 30-F11 Anti-mouse CD3 Biotin 145-2C11 Anti-mouse CD19 Biotin 6D5 Anti-mouse NK1.1 Biotin PK136 Anti-mouse MHCII AF700 M5/114.15.2 Anti-mouse CD64 PE/Cy7 X54-5/7.1 Anti-mouse CD11c APC/Fire750 N418 Anti-mouse CD11b FITC M1/70 Anti-mouse Ly6C PerCP/Cy5.5 HK1.4 Anti-mouse Ly6G APC 1A8
Fluorochrome Streptavidin eF450
intracellular Fluorochrome Clone Anti-mouse TNF APC/Cy7 MP6-XT22
Table 2.4 Mouse lymphocytes panel
Fluorochrome Viability dye: eFluor455UV
extracellular Fluorochrome Clone Anti-mouse CD45 APC-Cy7 30-F11 Anti-mouse CD19 BV785 6D5 Anti-mouse CD3 AF700 17A2 Anti-mouse CD4 BV510 RM4-5 Anti-mouse CD8 PerCP 53-6.7
39 Methods
Table 2.5 Mouse innate lymphoid cells panel
Fluorochrome Viability dye: eFluor455UV
extracellular Fluorochrome Clone Anti-mouse CD90.2 APC/Cy7 30-H12 Anti-mouse CD11c Biotin N418 Anti-mouse B220 Biotin RA3-6B2 Anti-mouse Ly-6G/Ly-6C (Gr-1) Biotin RB6-8C5 Anti-mouse TCR β chain Biotin H57-597 Anti-mouse TCR γ/δ Biotin GL3 Anti-mouse TER-119 Biotin TER-119 Anti-mouse NKp46 BV421 29A1.4
Fluorochrome Streptavidin Pe/Cy5
intracellular Fluorochrome Clone Anti-mouse GATA3 APC 16E10A23 Anti-mouse RORγt PerCP/Cy5.5 Q31-378 Anti-mouse T-bet PE/Cy7 4B10 Anti-human/mouse EOMES FITC WD1928
2.7 RNA extraction and quantitative PCR
The mouse tissues or human biopsies were homogenized in tubes with TRI Reagent
(Zymo Research), or TRIzol (Invitrogen) containing ceramic beads using a
homogenizer at 20 m/s speed for 40 seconds. RNA was isolated from cells, mouse or
zebrafish tissues, whole zebrafish larvae or human biopsies following manufacturer’s
instructions. For DSS-treated mouse tissues, Direct-zol RNA MiniPrep kit (Zymo
Research) was used to remove the remaining DSS since it can block the reverse transcription (Viennois et al., 2013). Any DNA contamination was removed by TURBO
DNase (Invitrogen) kit, and the RNA was reverse transcribed using High Capacity cDNA Reverse Transcription (Applied Biosystems) or iScript cDNA synthesis (Bio-
Rad) kits according to manufacturer’s instructions. Quantitative Real-Time PCR (qRT-
40 Methods
PCR) was done using QuantiNova SYBR Green PCR (Qiagen) or iTaq™ Universal
SYBR® Green Supermix (Bio-Rad) kits and primers listed in Table 6.1. ABI ViiA 7
cycler or a CFX384 Touch Real-Time PCR was used for analysis, and Ct values were
normalized to that of housekeeping genes efa1, Hrpt, Gapdh or Actb and relative expressions were calculated by the formula 2^(-DCt).
2.8 RNA sequencing
For each sample, RNA was extracted from GPR35+ and GPR35- colonic macrophages
sorted from 1 or 2 Gpr35-tdTomato mice. RNA quality control was done using an
Agilent 2100 Bioanalyzer, and the concentration was determined by using the Quanti-
iT RiboGreen RNA assay Kit (Life Technologies). RNA was reverse transcribed into
cDNA by SMART-Seq v4 Ultra Low Input RNA Kit (Tamara). Sequencing libraries
were obtained by Nextera XT DNA Library Preparation Kit (Illumina). cDNA libraries
were pooled in equal amounts and sequenced SR81 using an Illumina NextSeq 500
Sequencing. Reads were aligned to the mouse genome (UCSC version mm10) with
STAR (version 2.5.2a) using the multi-map settings: '—outFilterMultimapNmax 10 --
outSAMmultNmax 1' (Dobin et al., 2013). Reads and alignments were assessed using
the qQCReport function of the R/Bioconductor package QuasR (R version 3.4.2,
Bioconductor version 3.6) (Gaidatzis et al., 2015). Reads were assigned to genes
employed by the UCSC refGene annotation (downloaded on 2015-Dec-18). The
number of reads (5’ends) overlapping with the exons of each gene was determined to
assume an exon union model by using the QuasR function qCount. In the case of un-
stranded SMART-Seq 4, reads mapping both strands were counted, while the True-
Seq data included only reads on the opposite strand. Differential gene expression
analysis was done using the R/Bioconductor package edgeR (McCarthy et al., 2012).
41 Methods
Genes with logCPM>1 in at least 1 sample were filtered. To compare different
genotype and treatment effects, two models were used. A nested design analysis was
done for comparison of treatment groups within each genotype. A non-nested model
with crossed genotype-treatment groups was performed to compare different
genotypes. To examine the model contrasts, the glmQLFit and glmQLFTest functions
of edgeR were used, respectively, for each model. The obtained P-Values were
adjusted to false discovery rate.
RNA-sequencing data in Figure 3.15B was taken from a published dataset
(Czarnewski et al., 2019).
2.9 In vitro assays
2.9.1 3’-5’-Cyclic adenosine monophosphate (cAMP) assay
cAMP HunterTM eXpress assay (Eurofins) was performed by following the manufacturer’s directions to test potential GPR35 ligand candidates. Chinese Hamster
Ovary (CHO)-K1 cells transfected with human GPR35 naturally coupled to an inhibitory G-protein were thawed and seeded on a 96-well plate at 3x105 cells/well.
° Cells were incubated overnight at 37 C, 5% CO2. Cells were given 15 µm Forskolin
and 1:3 serial dilutions of potential ligands with the following starting concentrations:
10 µM recombinant human CXCL17 (R&D Systems), 10 µM lysophosphatidic acid
(LPA) or 10 mM kynurenic acid (KYNA) (both Sigma Aldrich). Zaprinast (Sigma
Aldrich) was used as a positive control, as recommended by the manufacturer. The
cells were incubated at 37 °C for 30 minutes. cAMP concentration was determined by
enzyme-fragment complementation (EFC) technology that uses two separate
fragments of b- galactosidase. Briefly, cAMP labeled with one part of the enzyme is
outcompeted by the intracellular cAMP for binding to the anti-cAMP antibody and is,
42 Methods
therefore, free to join the enzyme complex that is otherwise unfunctional. This enzyme
complex then cleaves the added substrate resulting in a luminescent signal. The signal
was then detected by a Synergy H1 Microplate Reader (Biotek).
2.9.2 Mouse bone marrow-derived macrophages (BMDMs)
For bone marrow isolation, femur and tibias from WT or Gpr35-deficient mice were cut
at both ends, and bone marrow was washed out with PBS. The cells were seeded at
a density of 2x105 cells/mL in RPMI 1640 supplemented with 10% FBS, 0.05 mM 2-
ME, 100 U/mL penicillin and 100 µg/mL streptomycin, and 20 ng/mL M-CSF
° (BioLegend) for macrophage differentiation. The cells were grown at 37 C, 5% CO2,
and the medium was changed on days 3 and 5. On day 7 of the culture, the BMDMs
were treated with 10 µM LPA or 10 ng/mL LPS for 1 hour or 4 hours for western blot
and RNA extraction, respectively.
2.9.3 Trans-well migration assay for mouse BMDMs
On day 7 of the BMDM culture, cells were collected using an ice-cold PBS solution.
5x105 BMDMs were cultivated on trans-well inserts with 5 µm pore size (Corning) in
100 µl of RPMI 1640 containing. The inserts were placed on a 24-well plate in the
corresponding wells with 600 µl of RPMI 1640 medium containing 2% FBS and 1%
P/S supplemented with vehicle or 10 µM LPA. The cells were incubated and allowed
to migrate for 18 hours. Migrated cells in the wells and the remaining cells on the
inserts were collected and resuspended in 200 µl of FACS buffer. 70 µl of each sample
was acquired by BD AccuriTM C6 flow cytometer (BD Biosciences). Percentages of
migrated cells were calculated relative to the sum of cells in upper and lower chambers
of the same well.
43 Methods
2.9.4 Enzyme-linked immunosorbent assay (ELISA) for corticosterone
For quantifying corticosterone, mouse colonic explants were used. After washing, weighing, and opening longitudinally, 0.5 cm colon tissues were incubated for 24 hours
° at 37 C, 5% CO2 on 24-well plates containing 500 µl of DMEM with 2% FBS, and 100
U/mL penicillin and 0.1 mg/mL streptomycin. Corticosterone Competitive ELISA kit
(Invitrogen) was used following the manufacturer’s instructions, and the concentrations calculated were normalized to the weights of colon pieces measured before the assay.
2.9.5 LPA ELISA using ex vivo colonic explants of DSS-treated mice
The colonic explants were obtained and cultured for 24 hours as described above.
General LPA ELISA kit (MyBiosource) was used according to the manufacturer’s instructions, and the values were normalized to the weights of extracted colonic tissues.
2.10 Western blot LPA- or LPS- treated BMDMs
BMDMs were collected and lysed in cold radioimmunoprecipitation assay (RIPA) buffer with sodium orthovanadate, PMSF, protease inhibitor cocktail (all from Santa
Cruz). BCA method was used to quantify the concentration of protein in each sample.
The amount of protein was adjusted to 10 µg before the samples were transferred onto a nitrocellulose membrane by electrophoretic separation. Blocking of the membranes was done in 5% dry milk in Tris Buffered Saline + Tween20 (TBS-T) buffer. The primary antibodies used are: phospho- NF-kB (p65), NF-kB, phospho-
ERK1/2, ERK1/2, phospho- STAT3, STAT3 (all from Cell Signaling Technology) and b-actin (BD Biosciences) prepared at 1:1000 or 1:2000 dilutions. The secondary
44 Methods
antibodies are the horseradish peroxidase-conjugated secondary antibodies anti-
rabbit IgG (H+L) or anti-mouse IgG (H+L) (both Jackson ImmunoResearch) used at
1:30000 dilution. Chemiluminescent detection was done using SuperSignalTM West
Femto or SuperSignal West Pico PLUS kits (both Thermo Fisher).
2.11 Identification of GPR35T108M and TNF blocker response in IBD patients
To detect the GPR35T108M variant among the patients (see Table 2.2), SNPs were genotyped in Sequenom, Hamburg, Germany. Genomic DNA samples were used at a concentration range of 5-10 ng/µl. Genotyping was performed by mass spectrometry by using the products from the primer extension. TNF blocker responses of all the patients were defined as breakthrough/loss of response, primary non-response (never effective) or side effects/intolerance.
2.12 Statistical analysis
Data are presented as dot plots of individual values with medians unless stated otherwise. GraphPad Prism was used to depict the data and to determine statistical significance. p values were determined using unpaired t-test, Mann-Whitney U or two- way ANOVA tests depending on the experiment. Grubbs’ test was used to identify the outliers. p values were indicated as followed: *: p<0.05, **: p<0.01, ***: p<0.001
45 Results
3 RESULTS
3.1 Colonic macrophages express GPR35
3.1.1 Gpr35b is predominant in the intestine of zebrafish
Our interest in studying GPR35 as a risk gene in IBD in zebrafish prompted us to
identify two zebrafish GPR35 paralogs, Gpr35a, and Gpr35b. Sequence alignment
analysis revealed that Gpr35a and Gpr35b share 25.6% and 24% homology with
human GPR35, and 24.4% and 24.3% homology with mouse GPR35, respectively
(Figure 3.1A). In addition, both paralogs were more closely related to human and
mouse GPR35 compared to human or mouse GPR55, a highly similar protein, as
shown in the phylogenetic tree (Figure 3.1B).
Next, we measured gene expression levels of gpr35a and gpr35b in the intestine and
the rest of the body at 72 and 120 hpf. Gpr35a mRNA levels were comparable at 72
and 120 hpf. Likewise, at 72 hpf, gpr35b transcripts were similar while intestinal gpr35b
expression was higher than that of the body at 120 hpf (Figures 3.1C and 3.1D).
The whole-mount in situ hybridization (WISH) confirmed the specificity of gpr35b in
the intestine at 120 hpf and located the gpr35b mRNA in the anterior intestine called the intestinal bulb (Figure 3.1E).
46 Results
Figure 3.1 gpr35b mRNA levels are pronounced in the zebrafish intestine at 120 hpf
(A) Gpr35a and Gpr35b zebrafish paralog protein sequences (red) aligned with mouse and human
GPR35 by ClustalW. Alignment scores between each of the protein pairs are indicated (below).
(B) Phylogenetic tree indicating homology between human, mouse, and zebrafish GPR35 and also
GPR55 orthologs by ClustalW.
(C) Expression of gpr35a and gpr35b mRNA by qRT-PCR normalized to ef1a in the intestine or rest of the body of WT zebrafish larvae at 120 hpf. The data depicts one representative experiment from two different experiments. A.U.; arbitrary units normalized to the lower value.
(D) Expression of gpr35b mRNA by qRT-PCR normalized to ef1a in the body and intestine of zebrafish larvae at 72 hpf and 120 hpf. A.U.; arbitrary units normalized to the lower value.
(E) Expression of gpr35b mRNA by whole mount in situ hybridization (WISH) in zebrafish larvae at 96 hpf and 120 hpf. The dashed line indicates the intestinal tract, and the bar-headed line indicates the intestinal bulb. The image is representative of 40 larvae.
Data are represented as individual values with medians with each dot representing one biological replicate. *p<0.05 by two-way ANOVA with Tukey’s multiple comparisons test.
47 Results
3.1.2 Gpr35 is expressed in the human and mouse gastrointestinal tract
To examine the GPR35 expression in humans, we took advantage of the human
protein atlas database (proteinatlas.org). The analysis revealed that GPR35 is more
pronounced in the gastrointestinal tract, stomach, duodenum, small intestine, colon,
and rectum compared to the other tissues such as skin, kidney, or liver. (Figure 3.2A).
In mice, gene expression analysis showed that Gpr35 is expressed in higher levels in
proximal and distal stomach, segments of the gut, gut-draining mesenteric lymph
nodes (MLN) and gut-associated lymphoid tissue Peyer’s patches (PP) compared to
the lower expression in liver (Figure 3.2B). Notably, Gpr35 had an increased
expression pattern from the duodenum to the distal colon.
Figure 3.2. Gastrointestinal tract shows high levels of Gpr35 transcripts in human and mouse
(A) GPR35 mRNA expression levels across indicated human tissues from the Human Protein Atlas database (proteinatlas.org).
(B) Expression levels of Gpr35 mRNA by qRT-PCR normalized to Gapdh in indicated tissues from WT mice. Data are represented as individual values with medians with each dot representing one biological replicate.
3.1.3 Macrophages in the intestinal lamina propria are GPR35+
To identify the cells expressing GPR35 in the intestine, we generated a Gpr35-
tdTomato reporter mouse line (Figure 2.1). GPR35 immunofluorescence staining of
the small intestine colocalized with the tdTomato confirmed that tdTomato-positive
cells indeed express GPR35 (Figure 3.3A). To monitor GPR35 expression in
48 Results
CX3CR1+ macrophages, we crossed the Gpr35-tdTomato line with Cx3cr1-GFP reporter mice. Ex vivo imaging of the double reporter mice revealed GPR35 in intestinal epithelial cells and lamina propria CX3CR1+ macrophages in the ileum and
distal colon (Figure 3.3B). In the MLN, GPR35+ cells were confined in the subcapsular
sinus and T cell zone and were absent in B cell follicles. GPR35+ cells were also
present in isolated lymph follicles (ILF) and subepithelial dome region of Peyer’s
patches. Similar to the MLN, B cell follicles in the PP were also deprived of GPR35+
cells.
Figure 3.3. IECs and CX3CR1+ macrophages are GPR35+
(A) Immunofluorescence staining of GPR35 (red) in the small intestine of Gpr35-tdTomato reporter mice. NucBlue (blue) was used for nuclear staining. SA: secondary antibody; tdT: tdTomato (orange).
Scale bars represent 50 µm.
(B) Ex vivo imaging of ileum, distal colon, mesenteric lymph node (MLN), isolated lymph follicle (ILF), and Peyer’s patches (PP) of Cx3cr1-GFP (green) x Gpr35-tdTomato (red) double reporter mice. Distal colon from WT was used as background control. LP: lamina propria; IEC: intestinal epithelial cell; TZ:
T cell zone; BF: B cell follicle; SCS: Subcapsular sinus. Arrows indicate CX3CR1+GPR35+ double-
positive cells. Scale bars represent 50 µm.
To examine immune cell types in terms of GPR35 expression, we performed flow cytometry using colonic lamina propria cells from the Gpr35-tdTomato reporter mice.
The analysis showed that 72% of CD64-CD11c+ dendritic cells were GPR35+. On the
49 Results
other hand, the expression was not detectable in B cells, CD4+ T cells, CD8+ T cells, neutrophils, natural killer (NK) cells, or innate lymphoid cells, ILC1, ILC2, and ILC3s
(Figure 3.4).
Figure 3.4. Characterization of immune cells for their GPR35 expression
Histograms showing percentages of GPR35+ B cells, CD4+ T cells, CD8+ T cells, neutrophils, dendritic cells, natural killer (NK) cells, ILC1, ILC2, and ILC3 in the colonic lamina propria of Gpr35-tdTomato reporter mice (blank red) and WT mice (gray) as background controls.
The monocyte precursors continuously replenish the macrophages in the intestine as
they downregulate the Ly6C expression (Bain et al., 2013). Therefore, we analyzed
MHCII+CD64+ macrophages and their MHCII-CD11b+Ly6Chigh to low monocyte
precursors for their GPR35 expression by flow cytometry (Figure 3.5A). Most of the
Ly6Chigh and Ly6Cmid monocytes in the lamina propria of the small and large intestine
were GPR35+ (Figure 3.5B and 3.5C), whereas the percentages decreased in Ly6Clow
monocytes and macrophages. Consequently, approximately 30% and 50% of
macrophages expressed GPR35 in the small intestinal and colonic lamina propria.
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Figure 3.5 Percentages of GPR35+ cells decrease from monocytes to macrophages
(A) Gating strategy for intestinal lamina propria mononuclear phagocytes, including Ly6Chigh to low monocytes and macrophages. Lin: lineage against CD3, CD19, and NK1.1 to exclude T, B, and NK cells.
(B) Representative histograms depicting GPR35 expression by flow cytometry in monocyte subsets
(Ly6Chigh to Ly6Clow) and macrophages (MAC) from small intestinal and colonic lamina propria (si-LP and co-LP) of Gpr35-tdTomato reporter mice (blank red) and WT mice (gray) as the background control.
Numbers in histograms indicate the percentage of GPR35-tdTomato+ cells.
(C) Quantification flow cytometric data from (B) of the percentages of GPR35+ cells in macrophages and their monocyte precursors in the si-LP and co-LP. Data are represented as individual values with medians with each dot representing one biological replicate. *p<0.05, **p<0.01, ***p<0.001 by two-way
ANOVA with Tukey’s multiple comparisons test.
3.1.4 GPR35+ and GPR35- colonic macrophages are transcriptionally distinct
To gain insight into the functional differences between the GPR35+ and GPR35- colonic macrophages under steady-state conditions, we performed bulk RNA- sequencing analysis following fluorescent activated cell sorting (FACS). Principal component analysis segregated the GPR35+ and GPR35- macrophages separately
(Figure 3.6A), and unsupervised hierarchical clustering of the dataset revealed 2798 down- and 2773 upregulated genes (Figures 3.6B). Notably, GPR35+ macrophages
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had higher expression levels of pro-inflammatory cytokines Il1b, Il1a, Tnf, and Il23a in comparison to GPR35- macrophages (Figure 3.6C).
Figure 3.6 Colonic lamina propria macrophages can be distinguished into GPR35+ and GPR35-
populations
(A) Principal component (PC) analysis on RNA sequencing dataset of GPR35+ (GPR35pos) and GPR35-
(GPR35neg) macrophages (MAC) sorted from colon lamina propria (co-LP) of Gpr35-tdTomato reporter
mice.
(B) Unsupervised hierarchical analysis depicted as a heatmap on RNA sequencing dataset of GPR35+
(GPR35pos) and GPR35- (GPR35neg) colonic lamina propria macrophages (co-LP MAC).
(C) Cytokine expression profiles GPR35+ (GPR35pos) and GPR35- (GPR35neg) colonic lamina propria macrophages (co-LP MAC) represented as a heatmap.
3.2 Regulation of GPR35 by the microbiota and intestinal inflammation
3.2.1 Gpr35 expression is microbiota-dependent in zebrafish and mice
To characterize the GPR35 expression pattern under different microbiota conditions,
we induced bacterial perturbation in zebrafish larvae with antibiotics. Antibiotics-
treated zebrafish showed lower mRNA levels in WISH analysis in the intestinal bulb
compared to vehicle-treated fish (Figure 3.7A). The qRT-PCR analysis confirmed the
reduction in Gpr35 transcripts in the zebrafish intestine in response to antibiotics.
There were no significant differences in Gpr35 expression in the rest of the body
(Figure 3.7B).
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To test our findings in mice, we administered a broad-spectrum antibiotics cocktail consisted of vancomycin, neomycin, metronidazole, gentamicin, and ampicillin. mRNA levels of Gpr35 were lower in the colonic lamina propria of antibiotics-treated mice compared to non-treated controls. The expression levels remained comparable in the intestinal epithelial cell compartment between the control and antibiotics-given mice
(Figure 3.7C).
We compared the Gpr35 expression levels also in the colon of germ-free (GF) and specific pathogen-free (SPF) mice (Figure 3.7D). We identified significant downregulation of Gpr35 in the colon of GF animals when compared to SPF mice.
Figure 3.7 Gpr35 is downregulated in antibiotics-treated and GF animals
(A) gpr35b mRNA levels by WISH in zebrafish larvae at 120 hpf treated with antibiotics (ABX) or vehicle.
Arrowheads mark the intestinal bulb. One image is representative of 40 larvae.
(B) gpr35 mRNA levels by qRT-PCR normalized to ef1a in the intestine or rest of the body of zebrafish larvae at 120 hpf treated with antibiotics or vehicle. Each dot represents one replicate pooled from 10 larvae.
(C) Gpr35 mRNA levels by qRT-PCR normalized to Hprt in the intestinal epithelial cell (IEC) and colonic lamina propria (co-LP) compartments of mice treated with antibiotics or vehicle.
(D) Gpr35 mRNA levels by qRT-PCR normalized to Hprt in the whole colon of specific pathogen-free
(SPF) or germ-free (GF) mice.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate unless stated otherwise. *p <0.05, **p < 0.01, ***p < 0.001 by unpaired t-test (D) or by two- way ANOVA with Tukey’s multiple comparisons test (B, C).
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3.2.2 Intestinal inflammation elevates GPR35 expression in zebrafish and mice
To explore the possibility that GPR35 plays a potential role in intestinal inflammation,
we first investigated whether inflammatory conditions regulate GPR35. In zebrafish,
intestinal inflammation induced by TNBS led to higher gpr35b mRNA levels in the
intestinal bulb compared to vehicle-treated controls. TNBS also resulted in an ectopic gpr35b expression in the posterior intestine of zebrafish (Figure 3.8A). In line with the
WISH analysis, TNBS-treated zebrafish larvae displayed increased gpr35b transcript
levels measured by qRT-PCR in the intestine in comparison to vehicle-treated larvae
(Figure 3.8B).
In mice, we induced colitis by treating the mice with DSS to study the GPR35
expression under inflammation. Ex vivo imaging of Cx3cr1-GFP x Gpr35-tdTomato
double reporter mice showed that GPR35-tdTomato signal by CX3CR1+ macrophages
is increased by DSS treatment in the colon (Figure 3.8C). Flow cytometric analysis of
colonic lamina propria cells from double reporter mice revealed that the number of
GPR35+ colonic lamina propria macrophages is higher in the DSS-given mice than
controls. (Figure 3.8D).
Figure 3.8 Inflammation leads to an increase in GPR35-expressing immune cells in intestinal LP
(A) gpr35b mRNA levels by WISH in zebrafish larvae at 120 hpf treated with TNBS or vehicle.
Arrowheads indicate the intestinal bulb and the posterior intestine. One image is representative of 20 larvae.
(B) gpr35b mRNA transcript levels by qRT-PCR normalized to ef1a in the intestine and rest of the body of zebrafish treated with TNBS or vehicle. Each dot is one biological replicate pooled from 10 larvae.
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(C) Ex vivo imaging of colon from untreated (UT) or DSS-treated Cx3cr1-GFP (green) x Gpr35- tdTomato (red) double reporter mice on day 7 of DSS colitis. Scale bars, 50 µm.
(D) Numbers of GPR35-tdTomato+ monocytes (Ly6Chigh to Ly6Clow) and macrophages (MAC) quantified from flow cytometric analysis of co-LP from UT or DSS-treated Gpr35-tdTomato mice.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate unless stated otherwise. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test.
3.2.3 Bacterial infection or colonization induces GPR35 in the intestine
DSS colitis in mice and TNBS-induced intestinal inflammation in zebrafish are triggered by translocation of microbial components as a result of epithelial damage
(Laroui et al., 2012; Oehlers et al., 2011). Therefore, we hypothesized that the bacteria might be a determining factor in enhancing GPR35 expression during intestinal inflammation. To test this hypothesis, we injected Vibrio anguillarum extract into swim bladders of zebrafish. Similar to the TNBS-treatment, bacterial infection with Vibrio anguillarum led to an increase in gpr35b mRNA levels in the intestine in comparison to PBS control (Figure 3.9A).
To recapitulate our findings in mice, we gavaged Cx3cr1-GFP x Gpr35-tdTomato double reporter mice with Escherichia coli DH10B pCFP-OVA (Rossini et al., 2014).
Ex vivo imaging of colon of PBS- and E. coli-given mice revealed that colonization resulted in increased GPR35-expressing CX3CR1+ macrophages (Figure 3.9B).
Consistently, flow cytometric analysis of colonic lamina propria cells showed that percentage of GPR35+ macrophages is higher in E. coli colonized mice compared to controls (Figure 3.9C and 3.9D).
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Figure 3.9 Bacterial inflammation causes increased GPR35
(A) gpr35b transcript levels measured by qRT-PCR normalized to ef1a in PBS- or V. cholerae-given zebrafish. Each dot represents one biological replicate pooled from 10 larvae.
(B) Ex vivo imaging of colon from Cx3cr1-GFP (green) x Gpr35-tdTomato (red) mice gavaged with PBS or E. coli every other day for 21 days. Scale bars, 50 μm.
(C) Representative flow cytometry histograms showing GPR35-tdTomato signal by co-LP macrophages from WT (gray histogram), PBS-gavaged Cx3cr1-GFP x Gpr35-tdTomato (red blank histogram), and E. coli-gavaged Cx3cr1-GFP x Gpr35-tdTomato (red filled histogram) mice.
(D) Quantified flow cytometric data from (C) for percentages of GPR35+ cells in co-LP macrophages in
PBS-or E. coli-gavaged Cx3cr1-GFP x Gpr35-tdTomato mice.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate unless stated otherwise. *p <0.05, **p < 0.01, ***p < 0.001 by one way (A) or Mann-Whitney
(D).
3.2.4 Increased number of GPR35+ cells in patients with ulcerative colitis
To translate our findings on GPR35 regulation in zebrafish and mouse models into human IBD, we performed immunostaining for GPR35 on intestinal biopsies of ulcerative colitis or Crohn’s disease patients taken from non-inflamed or inflamed regions of the corresponding patient. Numbers of GPR35+ cells in the lamina propria were comparable between non-inflamed and inflamed regions of Crohn’s disease patient biopsies. On the other hand, infiltrates of GPR35+ cell clusters in the lamina propria were present in inflamed ulcerative colitis biopsies compared to non-inflamed
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Figure 3.10 GPR35+ lamina propria cells are present in greater numbers in inflamed sites of UC
(A) Representative images of immunofluorescence staining on UC or CD patient biopsies from non- inflamed (left panel) and inflamed regions (right panel). Tissue sections were stained for GPR35 (red) and NucBlue (blue) for counterstaining. Scale bars, 50 µm.
(B) Semi-quantitative analysis of the number of GPR35+ cells in the lamina propria per mm2 from immunofluorescence images shown in (A) of non-inflamed and inflamed regions of UC or CD patient biopsies.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test.
3.3 Lysophosphatidic acid signaling depends on GPR35 in macrophages
3.3.1 LPA inhibits the cAMP release in humanGPR35-transfected cells
To examine the ability of LPA, KYNA, or CXCL17 to activate GPR35 signaling, we have measured intracellular cAMP levels in Chinese hamster ovary (CHO)-K1 cells transfected with human GPR35 naturally coupled to an inhibitory G (Gi) protein in response to each ligand candidate. Zaprinast, a synthetic agonist of GPR35, used as a positive control, inhibited the forskolin-induced conversion of cAMP (Figure 3.11).
Among the tested potential ligand candidates of GPR35, kynurenic acid did not
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significantly reduce the forskolin-initiated cAMP production. The addition of LPA and
CXCL17 triggered dampening of intracellular cAMP levels in forskolin-treated GPR35- transfected cells at a lower concentration of LPA compared to CXCL17 (Figure 3.11).
Figure 3.11 LPA reduces forskolin-induced cAMP accumulation in GPR35-transfected cells
Intracellular cAMP levels quantified as relative luminescence units (RLU) in Gi-coupled human GPR35-
transfected CHO-K1 cells treated with adenylyl cyclase activating-forskolin and serial dilutions of
zaprinast, KYNA, CXCL17, or LPA at indicated concentrations. Data are represented as median ± range
from doublets with nonlinear fit curves.
3.3.2 LPA leads to GPR35-dependent Tnf induction in zebrafish
LPA induces the production of cytokines such as TNF in Jurkat T cells and IL1-b in
mouse and human macrophages (Chang et al., 2008; Zhang et al., 2016). Therefore,
to explore the possibility that LPA signaling depends on GPR35, we examined whether
LPA alters the expression levels of cytokines in presence or absence of GPR35. To answer this at the multispecies level, we first generated a gpr35b mutant zebrafish line, gpr35buu189, by using CRISPR/Cas9-based genome engineering (M. Li et al.,
2016). WT zebrafish larvae treated with LPA or the zaprinast, a synthetic GPR35 agonist, showed an increase in expressions of the pro-inflammatory cytokines tnf, il1b, and il17a/f compared to untreated fish. On the other hand, LPA and zaprinast did not lead to a significant difference in the expression of these cytokines in gpr35buu189
mutant larvae (Figure 3.12).
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Figure 3.12 LPA leads to upregulation of tnf in WT but not in GPR35-deficient zebrafish
mRNA levels of tnf, il1b, and il17a/f by qRT-PCR in untreated (UT) and zaprinast- or LPA-treated WT
and gpr35buu1892 zebrafish larvae at 72 hpf for 48 hours. Each dot represents one biological replicated
pooled from 10 larvae. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple
comparisons test.
3.3.3 GPR35-deficiency leads to altered LPA signaling in BMDMs
Next, we constructed a Gpr35 knock-out (KO) mouse strain by CRISPR/Cas9 genome
editing system. To confirm the lack of GPR35 expression in Gpr35-/- mice, we
performed immunofluorescence staining of GPR35 on colonic tissue. No GPR35
staining was evident in the colon of Gpr35-/- mice (Figure 3.13A). To examine the
impact of GPR35-deficiency on transcriptional changes in response to LPA in macrophages, we stimulated bone marrow-derived macrophages (BMDMs) with LPS
or LPA and performed RNA sequencing. LPS treatment resulted in similar
transcriptomic profiles between WT and GPR35-deficient BMDMs (Figure 3.13B;
upper plot). On the other hand, WT BMDMs displayed a differential transcriptional
profile upon LPA stimulation compared to that of GPR35-deficient BMDMs (Figure
3.13B; lower plot).
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Figure 3.13 GPR35 mediates LPA signaling in mouse BMDMs
(A) Immunofluorescence staining of GPR35 in colonic tissues from WT (left panel) or Gpr35-/- mice
(right panel). Sections were stained against GPR35 (red) and with NucBlue (blue) for nuclear staining.
Scale bars represent 50 µm.
(B) Fold changes in mRNA levels in control and LPS- or LPA- treated BMDMs from WT and Gpr35-/-
mice obtained from the RNAseq dataset. Each dot represents one gene, and red dots indicate genes that are differentially regulated significantly only in WT but not in Gpr35-/- cells in response to
correspondent stimuli compared to control.
To investigate the signaling cascades activated by LPA in WT and GPR35-deficient
BMDMs, we studied the phosphorylation patterns of ERK, NF-KB, and STAT3 by western blot. Phosphorylation of both ERK and NF-KB (p65) was evident in LPA- treated WT but not GPR35-deficient BMDMs, whereas they remained comparable between genotypes in LPS-treated cells (Figures 3.14A, B). LPA-treatment did not result in a significant level of STAT3 phosphorylation in neither WT nor Gpr35-/-
BMDMs as opposed to LPS-treatment that resulted in a similar amount of
phosphorylated STAT3 between the cells (Figures 3.14A, B).
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Figure 3.14 WT but not Gpr35-/- BMDMs show increased p-ERK upon LPA treatment
(A) Western blot of control, LPA- and LPS- given BMDMs from WT and Gpr35-/- mice for phosphorylated and total NF-kB (p65) and ERK1/2 (left) or STAT3 (right). b-actin was chosen as a protein loading control.
(B) Quantitative analysis of band intensity ratios between phosphorylated p65 (NF-kB), ERK1/2, and
STAT3 to the total of each protein measured from (A). Data are represented as dotplots and medians, each dot representing one biological replicate. **p<0.01 by two-way ANOVA with Tukey’s multiple comparisons test.
Similar to zebrafish larvae, we treated bone marrow-derived macrophages (BMDMs) from WT or Gpr35-/- mice with LPA. LPA led to significantly higher mRNA levels of the cytokines Tnf, Il1b, and Il23a in WT BMDMs compared to untreated cells. LPA could not induce higher expressions of these cytokines in GPR35-deficient cells. (Figure
3.13B). Notably, for LPA-treated cells, Tnf was higher expressed in WT BMDMs
61 Results compared to Gpr35-/- cells. Next, we examined the regulation of cytokines TNF, IL-1b, and IL-23 at the protein level by a bead-based immunoassay. LPA treatment of WT
BMDMs led to higher TNF production in comparison to Gpr35-/- cells. An increase in
TNF concentrations was also seen in LPS-treated WT cells compared to Gpr35-/-, although the difference was less prominent than LPA-treatment. LPA did not change the protein levels of IL-1b and IL-23 in BMDMs (Figures 3.15B).
Figure 3.15 LPA mediates GPR35-dependent Tnf induction in mouse BMDMs
(A) mRNA levels of Tnf, Il1b, Il23a, and Il1a by qRT-PCR normalized to Actb in untreated (UT) and
LPA- or LPS-treated bone marrow-derived macrophages (BMDMs) from WT or Gpr35-/- mice. Each dot represents one biological replicate.
(B) Concentrations of the cytokines TNF, IL-1b, and IL-23 in supernatants of UT and LPA or LPS- treated WT or Gpr35-/- BMDMs measured by LEGENDplexTM bead-based immunoassay.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test.
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3.3.4 LPA facilitates GPR35-dependent chemotaxis of macrophages
It has been shown that monocytes, microglia, and ovarian cancer cells migrate
towards LPA (Oh et al., 2017; Plastira et al., 2017; Takeda et al., 2019). Therefore,
we asked whether LPA serves as a chemoattractant for BMDMs in a GPR35-
dependent manner. Consequently, BMDMs from Gpr35-/- mice showed impaired
chemotaxis towards LPA when compared to WT BMDMs that showed a significantly
higher percentage of migrating cells (Figure 3.16A).
To validate our findings in vivo, we first crossed gpr35buu1892 mutant zebrafish with the
macrophage reporter strain Tg(mpeg1:mCherry) (Nguyen-Chi et al., 2015). Next, we injected LPA into the otic vesicle of WT or gpr35buu1892 reporter zebrafish. LPA
injection led to a macrophage infiltration resulting in a higher number of mpeg+
macrophages in the otic vesicles of WT reporter fish compared to PBS injected fish.
No significant macrophage chemotaxis was evident in gpr35buu1892 mutant fish in
response to LPA injection (Figures 3.16B and C).
Figure 3.16 Macrophages show chemotaxis towards LPA in a GPR35-dependent fashion
(A) Percentages of migrated BMDMs quantified by flow cytometry from WT or Gpr35-/- mice towards
untreated (UT) control or LPA. Each dot represents one biological replicate.
(B) Imaging of Tg(mpeg1:mCherry)WT and Tg(mpeg1:mCherry)gpr35buu1892 macrophage (mpeg1+) reporter zebrafish larvae injected with PBS or LPA (10 µM) in the otic vesicle (white dashed line).
(C) Quantification of macrophage numbers from (B) recruited into the otic vesicles Tg(mpeg1:mCherry)-
WT and Tg(mpeg1:mCherry)-gpr35buu1892 zebrafish larvae injected with PBS or LPA.
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Data are shown as dotplots with individual values and medians, each dot representing one biological
replicate. *p <0.05, **p < 0.01, ***p < 0.001) by two-way ANOVA with Tukey’s multiple comparisons
test.
3.4 Intestinal inflammation enhances Autotaxin and LPA production
Our data indicate that GPR35 is upregulated during intestinal inflammation, and LPA
signaling leads to GPR35-mediated modulation of cytokine expression in
macrophages. We, therefore, questioned whether LPA production might also be
regulated in the intestine under inflammatory conditions. To answer this, we first
determined the expression of atx encoding for autotaxin (ATX), an LPA-producing enzyme, in the inflamed zebrafish intestine using the TNBS model. TNBS treatment of zebrafish larvae resulted in a two-fold increase in atx mRNA levels compared to untreated larvae (Figure 3.17A). To recapitulate these findings in mice, we took advantage of a published longitudinal transcriptomic data obtained from DSS colitic mice (Czarnewski et al., 2019). The data revealed that Atx expression gradually increases peaking at d10 of DSS colitis (Figure 3.17B). To examine the expression of
ATX at the protein level and to identify the autotaxin-expressing cells in the intestine, we established an immunohistochemistry staining of ATX on mouse colonic tissue.
Semi-quantitative analysis of the staining showed that ATX+ cells are in the lamina
propria, and their numbers increase with DSS colitis, reaching the highest numbers
on day 10, at the peak of inflammation (Figures 3.17C and D). Of note, the increase
in the number of ATX+ cells by DSS colitis was not affected by GPR35-deficiency as
Gpr35-/- mice showed a similar increase to WT mice in response to DSS (Figure
3.17E).
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Figure 3.17 Intestinal inflammation results in higher Atx expression
(A) Autotaxin (atx) mRNA levels by qRT-PCR normalized to ef1a in the intestine or rest of the body of
TNBS or vehicle-treated zebrafish at 72 hpf for 48 hours. Each dot represents one biological replicate pooled from 10 larvae.
(B) Atx expression from published RNA-sequencing data (Czarnewski et al., 2019) in the colon of mice during DSS colitis and recovery. Each dot represents the mean from three mice per data point.
(C) Immunohistochemistry staining of autotaxin (brown) and H&E (blue) for counter staining on colonic tissue of DSS colitic mice at indicated timepoints. Scale bars represent 50 µm.
(D) Quantification of autotaxin+ cells from (C). One representative experiment is shown from two.
(E) The number of autotaxin+ cells quantified from immunohistochemistry of colonic tissue at day 0 and day 10 of DSS colitis using WT or Gpr35-/- mice.
Data are shown as dotplots with means with each dot representing one biological replicate unless stated otherwise. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test.
ATX+ cells in the lamina propria indicate host cells might produce LPA in the intestine.
We asked whether intestinal microbiota contributes to the LPA production. To answer this, we took advantage of a published metabolomics dataset measuring metabolite
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18 hours post gavage although there was a transient increase in colon tissue at 2 hours indicating that LPA is in equilibrium between host and microbiota in the colon in this model (Figure 3.18A).
Since the inflamed intestine showed higher ATX, we measured the LPA concentration in the colonic explants of DSS-treated mice. LPA levels increased in the colonic explants of DSS-given animals compared to controls in similar levels between the wild- type and GPR35-deficient animals, suggesting that LPA is synthesized in higher levels in the colon under inflammation regardless of GPR35 expression (Figure 3.18B).
Figure 3.18 DSS colitis is associated with increased LPA production independently of GPR35
(A) Log ratio of LPA in indicated organs or fluids coming from 13C labeled bacteria to host, 2 and 18 h after gavage of GF mice. Data is taken from a metabolomics analysis published by Uchimura et al.
(Uchimura et al., 2018).
(B) Amount of LPA in the supernatant of colonic explants from control or DSS-treated WT or Gpr35-/- mice measured by ELISA and normalized to weights of the colonic tissues.
Data are shown as dotplots with medians with each dot representing one biological replicate. *p <0.05,
**p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test.
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3.5 Macrophage-expressed GPR35 is protective in DSS colitis mouse model
3.5.1 GPR35-deficient mice have augmented DSS-induced colitis
Farooq et al. have shown that GPR35-deficiency leads to worsened signs of colitis in
DSS-treated mice (Farooq et al., 2018). To confirm these findings, we induced colitis
in WT and Gpr35-/- mice by DSS. GPR35-deficient mice displayed more body weight loss, higher disease score and more colon shortening. Histological analysis revealed that in colons of DSS-treated Gpr35-/- mice, more extensive immune cell infiltrations, loss of goblet cells, and thickening of muscle layer were evident, resulting in higher histological colitis scores (Figures 3.19A-F). These data suggest that GPR35 might
have a protective role in the DSS colitis mouse model.
Figure 3.19 GPR35-deficient mice show worsened signs of DSS colitis
(A) Percentages of body weight relative to the initial weight of untreated (UT) and DSS-treated WT or
Gpr35-/- mice. Data are represented as mean ± standard deviation (SD) (n=4).
(B) Disease activity scores of UT and DSS-treated WT or Gpr35-/- mice. Data are represented as mean
± SD (n=4).
(C) Representative images of dissected colons from UT and DSS-treated WT or Gpr35-/- mice on day
8 of colitis.
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(D) Colon lengths measured from colon images (C) of UT and DSS-treated WT or Gpr35-/- mice on day
8 of colitis.
(E) Representative images of H&E staining of colonic tissue from UT and DSS-treated WT or Gpr35-/- mice on day 8 of colitis. Scale bars represent 100 µm.
(F) Histological colitis scores from H&E staining of colonic tissues (E).
Data are represented as individual values with medians, each dot representing one biological replicate
(C-F) unless stated otherwise. *p<0.05, **p<0.01, *** p<0.001 by two-way ANOVA with Tukey’s multiple comparisons test (A, B) or Mann-Whitney (D, F).
3.5.2 Macrophage-specific GPR35 deletion leads to aggravated colitis
In the intestine, epithelial cells and CX3CR1+ macrophages express the GPR35. To elucidate the role of GPR35 signaling in intestinal macrophages, we generated
Gpr35flox mice and crossed them with Cx3cr1CreER mice to obtain tamoxifen-inducible conditional deletion of GPR35 in macrophages. The tamoxifen injection led to a reduction in GPR35+ cells among CX3CR1+ lamina propria macrophages in the intestine (Figures 3.20A and B).
Figure 3.20 Tamoxifen reduces the percentage of GPR35+ cells in colonic macrophages of
Gpr35ΔCx3cr1 mice
(A) Representative images of immunofluorescence staining of GPR35 (red) and NucBlue (blue) for nuclear staining on colonic tissues of Gpr35ΔCx3cr1 mice injected i.p. with vehicle (left) or tamoxifen (TMX)
(right). Arrowheads indicate CX3CR1-YFP+ macrophages. Scale bars represent 50 μm.
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(B) Percentages of GPR35+ cells among CX3CR1-YFP+ cells calculated from (A). Data are represented as individual values with medians, each dot representing one biological replicate. *p < 0.05 by Mann-
Whitney.
When given DSS, Gpr35ΔCx3cr1 mice showed a more pronounced progressive body
weight loss and disease activity during colitis compared to other control groups
(Figures 3.21A and B). Specific deletion of GPR35 in macrophages promoted more
severe colitis indicated by enhanced colon shortening, endoscopic and histological
signs of inflammation (Figures 3.21C-E).
Figure 3.21 GPR35-deficiency in macrophages increases DSS colitis severity
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(A) Body weight changes shown in percentages with respect to initial weights during DSS colitis for corn oil-injected (vehicle) or tamoxifen (TMX)-injected Gpr35wt or Gpr35ΔCx3cr1 mice. Data are shown as mean ± SD (n=4).
(B) Daily disease activity scores for vehicle or TMX-injected Gpr35wt or Gpr35ΔCx3cr1 mice during DSS colitis. Data are shown as mean ± SD (n=4).
(C) Representative images of dissected colons and colon lengths of vehicle- and TMX-injected Gpr35wt or Gpr35ΔCx3cr1 mice on day 7 of DSS colitis.
(D) Endoscopic images of distal colons of vehicle- and TMX-injected Gpr35wt or Gpr35ΔCx3cr1 mice on day 7 of DSS colitis.
(E) Representative images of H&E stainings of colons and histology scores of DSS-given vehicle- and
TMX-treated Gpr35wt or Gpr35ΔCx3cr1 mice on day 7. Scale bars represent 100 µm.
Data are shown as dotplots with medians with each dot representing one biological replicate unless stated otherwise. *p <0.05, **p < 0.01, ***p < 0.001 by two-way ANOVA with Tukey’s multiple comparisons test (A, B) or Mann-Whitney (C, E).
3.5.3 LPA attenuates DSS colitis in a macrophage-expressed GPR35 manner
In the context of colitis, the ATX/LPA axis has been shown to play pro- and anti- inflammatory roles. In one study, the administration of ATX inhibitor reduced inflammation in the chronic DSS colitis mouse model by blocking Th17 differentiation
(Dong et al., 2019) while another study has demonstrated that rectal perfusion of LPA in mice alleviates DSS colitis severity (Xu et al., 2014). On the other hand, the deletion of ATX was shown to suppress the severity of DSS-induced colitis (S. Lin et al., 2019).
To test whether LPA influences the colitis severity, we injected WT and Gpr35ΔCx3cr1 daily with 5 mg/kg LPA, a dosage shown to be protective in endotoxin-induced acute kidney injury model (Mirzoyan et al., 2017). LPA led to less body weight loss, lower disease activity, endoscopic and histological colitis scores in WT but not in
Gpr35ΔCx3cr1 mice (Figure 3.22A-E).
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Figure 3.22 LPA alleviates DSS colitis severity in WT but not in Gpr35ΔCx3cr1 mice
(A) Changes in body weights shown as percentages relative to initial body weights of vehicle- or LPA- treated Gpr35wt or Gpr35ΔCx3cr1 mice monitored daily for seven days during DSS colitis
(B) Disease activity scores of vehicle- or LPA-injected Gpr35wt or Gpr35ΔCx3cr1 mice assessed daily for seven days of DSS colitis.
(C) Representative colon images and colon lengths on day 7 of DSS colitis from vehicle- or LPA-injected
Gpr35wt or Gpr35ΔCx3cr1 mice.
(D) Representative endoscopy images and endoscopic colitis scores,
(E) H&E staining of colons and histological colitis scores corresponding day 7 of DSS colitis from vehicle- or LPA-treated Gpr35wt or Gpr35ΔCx3cr1 mice. Scale bars represent 100 µm.
Each dot represents one animal with medians unless stated otherwise. *p <0.05, **p < 0.01, ***p <
0.001 by two-way ANOVA with Tukey’s multiple comparisons test (A, B) or Mann-Whitney (C, D, E).
3.5.4 Gpr35ΔCx3cr1 mice have enhanced neutrophil infiltration during DSS colitis
Our findings suggest that GPR35-deficiency in macrophages leads to aggravated colitis in the DSS model and reduced migration towards LPA in vivo and in vitro.
Therefore, we asked whether macrophage-specific deletion of GPR35 has an impact on the number of immune cells in the lamina propria during colitis. Flow cytometric
71 Results analysis showed that Gpr35ΔCx3cr1 mice have a higher percentage and number of neutrophils in the colonic lamina propria during DSS colitis compared to other control groups. (Figures 3.23A-C). Notably, the percentage and number of colonic macrophages were comparable between Gpr35ΔCx3cr1 and WT mice.
Figure 3.23 Macrophage-specific GPR35 deletion results in a higher number of neutrophils during colitis
(A) Representative dotplots from flow cytometry of mononuclear phagocytes from colonic lamina propria of vehicle- and TMX-treated WT or Gpr35ΔCx3cr1 mice on day 7 of DSS colitis.
(B) Percentages of macrophages (MAC), dendritic cells (DCs), monocytes (Ly6Chigh, Ly6Cmid, and
Ly6Clow subsets) and neutrophils quantified from (A).
(C) The number of indicated mononuclear phagocytes calculated using flow cytometric data (A).
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate. *p <0.05, **p < 0.01, ***p < 0.001 by Mann-Whitney.
72 Results
3.5.5 GPR35 deletion in macrophages causes reduced TNF production in colitis
We have shown that in zebrafish larvae and BMDMs, LPA induces Tnf expression in a GPR35-mediated way. Therefore, we asked whether GPR35 modulates TNF production in macrophages during DSS colitis. To answer this question, we analyzed the TNF-producing cells among macrophages by flow cytometry. The analysis revealed that, in comparison to WT mice, Gpr35ΔCx3cr1 animals have lower frequencies and median fluorescence intensities for TNF+ macrophages (Figures 3.24A and B). In addition to TNF-producing macrophages, we also analyzed overall expression levels of the cytokines Il10, Il1b, Il6, or Tnf in colonic tissue (Figure 3.24C). mRNA levels of these cytokines were similar between WT and Gpr35ΔCx3cr1 mice.
C
Figure 3.24 GPR35-deficiency leads to lower TNF levels in macrophages during DSS colitis
73 Results
(A) Representative plots for TNF+ colonic lamina propria macrophages (co-LP MAC) by flow cytometry
on day 7 of DSS colitis using vehicle- and TMX-treated WT or Gpr35ΔCx3cr1 mice. Numbers on the plots
indicate the percentage of TNF+ cells among macrophages.
(B) Percentages of TNF-producing cells and median fluorescence intensity (MFI) of TNF in
macrophages from (A).
(C) mRNA levels of Il10, Il1b, Il6, and Tnf by qRT-PCR in colonic tissues of vehicle- and TMX-treated
WT or Gpr35ΔCx3cr1 mice on day 7 of DSS colitis.
Data are presented as individual values with medians with each dot representing one biological
replicate. *p < 0.05, Mann-Whitney U test.
3.6 TNF attenuates colitis and induces corticosterone synthesis in Gpr35ΔCx3cr1
mice
3.6.1 TNF is protective in Gpr35ΔCx3cr1 mice during DSS colitis
Although TNF is a pro-inflammatory cytokine, TNF signaling might exert compensatory
functions to dampen colitis depending on the context. TNF-deficiency in mice worsens
the DSS-induced colitis and TNF administration during oxazolone-induced colitis
alleviates the symptoms of inflammation (Naito et al., 2003; Noti et al., 2010).
We administered TNF daily during the DSS course to examine whether TNF can
dampen the exacerbated colitis in Gpr35ΔCx3cr1 mice. TNF treatment led to less severe body weight loss and disease activity and also colon shortening compared to PBS injection. TNF also improved endoscopic signs of colitis and the damaged mucosal
architecture of the colon in Gpr35ΔCx3cr1 animals. The colitis severity in TNF-injected
Gpr35ΔCx3cr1 mice was comparable to that of WT mice (Figures 3.25A-F).
74 Results
Figure 3.25 TNF administration attenuates colitis in Gpr35ΔCx3cr1 mice
(A) Body weight trends depicted as percentages normalized to the initial weight of DSS-given PBS- injected Gpr35wt (WT) and PBS- or TNF-injected Gpr35ΔCx3cr1 mice (left panel). Daily disease activity scores of PBS-injected Gpr35wt and PBS- or TNF-injected Gpr35ΔCx3cr1 mice (right panel). Data are shown as mean ± SD (n=4).
(B) Representative images (left) and colon lengths (right) of colons from DSS-treated PBS-injected
Gpr35wt (WT) and PBS- or TNF-injected Gpr35ΔCx3cr1 mice on day 7 of colitis.
(C-E) Endoscopic images (C), endoscopic colitis scores (D), representative H&E microscopy images of colons (E), and histological colitis scores (F) from PBS-injected Gpr35wt and PBS- or TNF-injected tamoxifen-treated Gpr35ΔCx3cr1 mice on day 7 of DSS colitis. Scale bars represent 100 µm.
Data are shown as dotplots with individual values and medians, each dot representing one biological replicate. *p <0.05, **p < 0.01, ***p < 0.001 by Mann-Whitney.
75 Results
3.6.2 TNF induces Cyp11b1 and corticosterone synthesis in Gpr35ΔCx3cr1 mice
TNF attenuates the oxazolone-induced colitis by triggering corticosterone production
in intestinal epithelial cells via increasing mRNA levels of Cyp11a1 and Cyp11b1,
which encode for steroidogenic enzymes (Noti et al., 2010). Therefore, we assessed
Cyp11a1 and Cyp11b1 expression levels in the colon of PBS- or TNF-treated
Gpr35ΔCx3cr1 mice. We observed no significant perturbations in Cyp11a1 expression
(Figures 3.26A). On the other hand, when compared to WT mice, Gpr35ΔCx3cr1 mice
had a lower expression of Cyp11b1. Furthermore, TNF-treatment led to increased
Cyp11b1 transcripts in these mice restoring the expression to similar levels measured in the colon of WT animals (Figures 3.26B). In line with the Cyp11b1 expression pattern, colonic explants of Gpr35ΔCx3cr1 had reduced concentrations of corticosterone
compared to WT colonic explants, and TNF-treatment resulted in increased corticosterone (Figure 3.26C).
Figure 3.26 TNF restores corticosterone synthesis in colitic Gpr35ΔCx3cr1 mice
(A-B) mRNA levels of Cyp11a1 (A) and Cyp11b1 (B) qRT-PCR of colonic tissue from PBS-injected
Gpr35wt and PBS- or TNF-injected Gpr35ΔCx3cr1 mice on day 7 of DSS colitis.
(C) Corticosterone levels in colonic explant supernatants of PBS-injected Gpr35wt and PBS- or TNF-
injected Gpr35ΔCx3cr1 mice on day 7 of DSS colitis. The values were normalized to weights of colonic
explants.
Data are presented as individual values with medians with each dot representing one biological
replicate. *p < 0.05, **p < 0.01, ***p < 0.001 by Mann- Whitney.
76 Results
3.6.3 TNF does not influence DSS colitis severity in WT mice
Next, we questioned whether TNF treatment influences DSS colitis severity in WT
animals. As opposed to the protective effect observed in Gpr35ΔCx3cr1mice, daily TNF
injection did not have a significant impact on colitis severity. We also measured colonic mRNA expression levels of the corticosterone-synthesizing enzymes Cyp11b1 and
Cyp11a1. Unlike the increase in Cyp11b1 and Cyp11a1 transcript levels in TNF-
treated Gpr35ΔCx3cr1mice, the expressions were not altered by TNF in WT animals
(Figures 3.27A-I).
Figure 3.27 TNF did not show a significant effect on DSS colitis in WT mice
(A) Body weights changes in percentages compared to the body weights on day 0 of PBS- or TNF- injected WT mice assessed for 7 days of DSS colitis. Data are represented as mean ± SD for 4 mice per group.
(B) Disease activity scores of PBS- or TNF-injected WT animals monitored daily for 7 days during DSS colitis. Data are represented as mean ± SD for 4 mice per group.
(C) Colon images,
(D) Colon lengths measured from (C) on day 7 of colitis from PBS- or TNF-treated WT mice.
77 Results
(E) Representative endoscopy images of PBS- or TNF-injected WT mice on day 7 of DSS colitis.
(F) Endoscopy colitis scores assessed from (E).
(G) H&E staining of colonic tissue (H) Histological colitis scores calculated from (G) of PBS- and TNF-
treated WT mice on day 7 of DSS colitis. Scale bars represent 100 µm.
(I) mRNA expression levels of Cyp11a1 and Cyp11b1 by qPCR normalized to Actb in total colonic tissue
taken on day 7 of DSS colitis from WT mice that received PBS- or TNF-injections.
Data are represented as dotplots with medians unless stated otherwise. Each dot represents one
biological replicate.
3.7 GPR35T108M variant might correlate with TNF blocker responses in IBD
GPR35 coding SNP rs3749171, is a C>T replacement that leads to a hyperactive
T108M missense variant (Schneditz et al., 2019). To better understand the function of
the GPR35T108M variant in IBD patients, we took advantage of the Swiss IBD Cohort
Study, including CD and UC patients genotyped for the SNP variant. We classified the
patients for breakthrough/loss of response, primary non-response (never effective)
and/or side effects/intolerance in response to TNF blockers. Although there were a
limited number of patients and the differences did not reach statistical significance, the
hyperactive GPR35T108M variant resulted in an increased percentage of patients that
responded successfully to TNF blockers (Table 3.1) suggesting a possibility that the
hyperactive GPR35T108M variant might be an indication of anti-TNF therapy
responsiveness in IBD.
Table 3.1 Responses to anti-TNF blockers in IBD patients with GPR35T108M variant
Ulcerative Colitis Crohn’s Disease All Response Success Fail Total Success Fail Total Success Fail Total C 2 7 9 13 23 36 15 30 45 (22.22%) (77.78%) (100%) (36.11%) (63.89%) (100%) (33.33%) (66.67%) (100%) CT 0 0 0 0 3 3 0 3 3 (100%) (100%) (100%) (100%) T 7 12 19 25 34 59 33 46 79 (36.84%) (63.16%) (100%) (42.37%) (57.63%) (100%) (41.77%) (58.23%) (100%) Total 9 19 28 38 60 98 48 79 127 (32.14%) (67.86%) (100%) (38.77%) (61.23%) (100%) (37.80%) (62.20%) (100%) Pearson chi2(2) = 0.5985 Pearson chi2(2) = 0.3653 Pearson chi2(2) = 0.8605 Pr = 0.439 Pr = 0.546 Pr = 0.3536
78 Discussion
4 DISCUSSION
4.1 Heterogeneous expression of GPR35 in intestinal lamina propria monocytes
and macrophages
Fate mapping, as well as Ccr2-/- mouse studies, have indicated that the intestinal macrophages are differentiated continuously from the extravasated Ly6Chigh
monocytes, a process that depends on the CCL2/CCR2 axis (Bain et al., 2014).
Because the mature macrophages scavenge commensal bacteria without generating
an inflammatory response owing to their hypo-responsiveness to TLR stimulation, they
are often described as “anti-inflammatory” in contrast to their monocyte precursors
that increase in numbers during inflammation (Rivollier et al., 2012). However, paradoxically, macrophages can contribute to inflammation, and monocytes are found in the steady-state. Interestingly, we found GPR35+ and GPR35- macrophages in the
steady-state lamina propria and GPR35+ macrophages show a rather pro- inflammatory gene expression profile compared to their GPR35- counterparts.
Although GPR35+ colonic macrophages were very distinct from the GPR35-
subpopulation at the transcriptional level, whether these differences correspond to
distinct functions under steady-state and inflammatory conditions remains unknown.
Given that all monocytes are GPR35+, it might be worthwhile to question whether loss
of GPR35 expression in some of the monocyte-derived macrophages is associated
with gaining anti-inflammatory features.
4.2 GPR35-mediated LPA signaling in macrophage cytokine and migration
responses
Previous studies have attempted to deorphanize the GPR35 and proposed several
potential endogenous ligands. Nevertheless, these proposed ligands remain disputed
79 Discussion
to date due to the counter-argumentative studies questioning their physiological
relevance (Binti Mohd Amir et al., 2018; Mackenzie et al., 2011; Maravillas-Montero et
al., 2015; Oka et al., 2010; Southern et al., 2013). The interspecies variation between
the ligand potencies being one of the challenges in GPR35 pharmacology (Milligan,
2011), we have combined an in vitro approach for human GPR35 activation with
GPR35-deficient zebrafish and mouse models in our study. Our results indicated that
GPR35 loss of function intervenes with LPA signaling, which was not investigated in the study where LPA was described as an endogenous ligand of GPR35 (Oka et al.,
2010). Another limitation of GPR35 ligand identification studies, including ours, is the possible existence of multiple putative endogenous ligands activating GPR35.
Likewise, LPA can also bind to other LPA receptors. Notably, Schneditz et al. demonstrated that LPA signaling leads to increased cytoplasmic Ca2+ in WT but not
GPR35-deficient BMDMs (Schneditz et al., 2019). However, the authors discussed
that the blunting effect of GPR35-deficiency on LPA signaling was due to the
interaction between GPR35 and Na/K-ATPase which might intervene with signaling
events via other LPA receptors. Furthermore, they propose that GPR35 exerts its
functions in a constitutive, ligand-independent manner since the baseline intracellular
Ca2+ levels decreased in Gpr35-/- compared to WT BMDMs. However, there is a possibility that GPR35 endogenous ligand(s) might be present in the culture, or a receptor/ligand binding is essential despite the putative baseline ligand-independent signaling events. Therefore, pharmacological screenings are necessary to reveal the direct receptor-ligand interactions at an inter-species level to understand LPA’s relevance to GPR35 better.
Although we have yet to explore whether LPA binds directly to GPR35, in this thesis, we have shown that the LPA signaling is GPR35-dependent in zebrafish and mouse
80 Discussion
macrophages. Of note, similar to zebrafish, mouse BMDMs have shown increased
TNF expression in response to the GPR35-mediated LPA signaling, which was accompanied by increased phosphorylation of ERK1/2 and NF-kB. Indeed, TNF is known to be driven by transcription factors activated as a result of NF-kB, ERK1/2, and JNK signaling cascades (Kishore et al., 2004). Nevertheless, whether TNF upregulation is due to the activation of these pathways remains to be elucidated.
LPA also served as a chemotactic factor in mouse BMDMs in a GPR35-dependent manner. However, although LPA has led to the migration of macrophages in the WT but not GPR35-mutant zebrafish, we cannot exclude the possibility that the LPA signaling might have been activated in other cells which then led to the macrophage infiltration. In addition, whether GPR35-dependent chemotaxis contributes to the intestinal homeostasis or inflammation remains to be investigated. However, our findings do indicate that LPA response is mediated by GPR35 in zebrafish and mice, particularly in macrophages.
4.3 Regulation of ATX/LPA/GPR35 axis in the intestine
Phospholipids are structural components of plasma membranes that can initiate signaling cascades. LPA is a small bioactive glycerophospholipid that can serve as a membrane component and a signaling molecule depending on its site of production
(Yang & Chen, 2018). LPA can be synthesized intracellularly by glycerol-3-phosphate acyltransferase (GPAT) as an intermediate in the biosynthesis of glycerolipids (Pages et al., 2001). In the extracellular compartment, LPA can be synthesized by phospholipase A1 and 2 from phosphatidic acid (Lee et al., 2019). However, membrane phospholipids are the main sources of LPA production. They are metabolized into lysophospholipids such as lysophosphatidylcholine (LPC) and are
81 Discussion
subsequently cleaved by the ATX, a process that determines the LPA plasma levels
(Aoki et al., 2008). Although the deletion of ATX leads to perinatal mortality in mice
due to developmental defects, Enpp2+/- heterozygote mice are viable yet show 50%
reduced LPA plasma levels in comparison to WT animals (Yung et al., 2014).
Our data indicated increased Atx expression in both zebrafish and mice under
inflammatory conditions. Increased Atx expression correlated with LPA levels in the
inflamed intestine. However, the array of metabolic pathways leading to LPA synthesis
in the inflamed, as well as the steady-state intestinal environment is yet to be explored.
To achieve this, LPA measurement in the intestine of Enpp2+/- mice may prove useful to determine to what extend the ATX serves as a determinant of LPA levels in the intestine. In addition to identifying of the enzymes catalyzing the LPA synthesis, studying the LPA release machinery is important to understand how the LPA abundance is regulated in the inflamed intestine. One possibility is that the disruption of epithelial cells may release of LPA or its precursors that get further metabolized into
LPA. Alternatively, phospholipids derived from the commensals may serve as an LPA source during colitis (Cullinane et al., 2005).
Notably, we found an association between GPR35 expression levels and microbiota as well as inflammation, the latter being in line with ATX and LPA expression patterns.
Therefore, to understand whether there is a causative relationship between regulation of ATX/LPA and GPR35, it is of importance to delineate the molecular events that govern the GPR35 expression and the role of this fine-tuning of expression under microbial perturbations and colitis.
82 Discussion
4.4 Reduced TNF and corticosterone levels during DSS colitis upon
macrophage-specific GPR35 deletion
TNF is a pleotropic cytokine that is mostly-recognized as pro-inflammatory in the context of IBD. TNF inhibition ameliorates intestinal inflammation in several mouse models, including spontaneous ileitis (Gunther et al., 2011), TNBS-induced colitis
(Neurath et al., 1997), and transfer colitis (Corazza et al., 1999). In IBD patients, the
TNF-producing mononuclear cells in the intestinal lamina propria correlate with mucosal inflammation (Reinecker et al., 1993), and anti-TNF antibodies revolutionized the treatment of the disease (Hanauer et al., 2002; Targan et al., 1997).
On the other hand, several studies have shown that TNF has context-dependent anti- inflammatory effects. Tnf-deficiency leads to more severe DSS-induced colitis (Naito et al., 2003; Noti et al., 2010), and TNF administration attenuates oxazolone colitis which is a Th2-type model that does not induce increased TNF expression (Noti et al.,
2010). Of note, 35% and 50% of IBD patients respond poorly to infliximab and vedolizumab, respectively (Paramsothy et al., 2018) and rheumatoid arthritis patients that received anti-TNF blockers, in particular etanercept, have an increased risk of IBD
(Korzenik et al., 2019). Here we describe a model where TNF is protective in the DSS colitis model in the absence of GPR35-signaling in macrophages that is accompanied by decreased TNF production. DSS colitis is an experimental model of IBD that is extensively used to study innate immune mechanisms involving macrophages and those activated by TLR or inflammasome signaling pathways (Fukata et al., 2005; Pull et al., 2005; Zaki et al., 2010). Despite being predominantly characterized by Th1 and
Th17-driven immune responses, DSS colitis closely resembles human UC histologically and symptomatically (Randhawa et al., 2014). Indeed, transcriptomic analysis of mice with DSS colitis and UC patients revealed that the inflammatory
83 Discussion
pathways are conserved, suggesting that DSS colitis is an indispensable model to
study IBD pathogenesis (Czarnewski et al., 2019). Our data indicate that the
attenuation of DSS colitis in macrophage-specific GPR35-deficient animals by TNF is associated with the induction of immunosuppressive corticosterone production via
Cyp11b1. On the other hand, in WT mice, TNF administration alters neither colitis severity nor Cyp11b1 expression. Therefore, our study supports that TNF can exert protective or deleterious functions depending on the context and stage of the intestinal inflammation.
In our hands, LPA alleviated DSS-induced colitis in WT mice, while the impact was lost in macrophage-specific GPR35-deficient mice suggesting that GPR35-mediated
LPA signaling in macrophages might be important for reacquiring homeostasis upon chemically-induced colitis. Among the cellular processes initiated by LPA are proliferation, survival, migration, and cytokine secretion (Pages et al., 2001). However, whether IBD patients carrying the GPR35 variants have aberrant LPA signaling needs to be studied. In this Ph.D. thesis, we examined the IBD-associated hyperactive
T108M variant and anti-TNF treatment responses. It is worthwhile to investigate whether the GPR35T108M variant causes enhanced TNF production in IBD patients that contributes to the pathogenesis and therefore a better response to TNF blockers.
4.5 Conclusion and outlook
GPR35 is an IBD risk gene that promotes IEC turnover during wound healing and
tumorigenesis in the intestine (Schneditz et al., 2019; Tsukahara et al., 2017). In this
Ph.D. project, we examined a role for macrophage-expressed GPR35 in intestinal
immunity. We propose that, in the steady-state, GPR35+ and GPR35- macrophages
are present in the intestinal lamina propria, and GPR35 is associated with a pro-
84 Discussion
inflammatory transcriptional signature. GPR35-deficiency in macrophages impairs the
LPA signaling in vitro and prevents the attenuation of DSS-colitis by LPA-treatment.
Macrophage-specific GPR35-deficient mice have lower TNF and corticosterone production. It remains unknown whether the elevated inflammation and lower TNF production in these mice are due to impaired LPA signaling. Therefore, before GPR35 can be considered as a possible target for IBD treatment, pharmacological screenings need to be carried out to reveal the receptor-ligand interactions surrounding LPA and
GPR35. In addition, further studies need to be conducted to describe whether GPR35 signaling in macrophages plays a role in TNF regulation in IBD patients.
Figure 4.1 GPR35 signaling in macrophages during DSS colitis
A working model depicting that DSS-colitis leads to an increase in LPA production independently of
GPR35, whereas LPA-signaling and TNF-responses are diminished in GPR35-deficient intestinal macrophages which are associated with reduced corticosterone synthesis .
85 Appendix
6 APPENDIX
6.1 Supplementary Information
Table 6.1 Primer list
qRT-PCR Primers Target Gene Forward Sequence Reverse Sequence Mouse Gpr35 Qiagen Cat no: QT00495411 Mouse Gapdh CATCAAGAAGGTGGTGAAGC CCTGTTGCTGTAGCCGTATT Mouse Actb TTCTTTGCAGCTCCTTCGTT ATGGAGGGGAATACAGCCC Mouse Tnf CCACCACGCTCTTCTGTCTAC AGGGTCTGGGCCATAGAACT Mouse Il1b TGTGAAATGCCACCTTTTGA GGTCAAAGGTTTGGAAGCAG Mouse Il1a CGCTTGAGTCGGCAAAGAAAT CTTCCCGTTGCTTGACGTTG Mouse Il23a AATGTGCCCCGTATCCAGTG CAAGCAGAACTGGCTGTTGTC Mouse Il10 ATCGATTTCTCCCCTGTGAA TGTCAAATTCATTCATGGCCT Mouse Il6 TCGGAGGCTTAATTACACATGTTC GCATCATCGTTGTTCATACAATC Mouse Cyp11a1 TGGGGTCCTGTTTAAGAGTTCA CTGCTTGATGCGTCTGTGTAA T A Mouse Cyp11b1 Qiagen Cat no: QT01198575 Human GPR35 Qiagen Cat no: QT02403128 Human GAPDH Qiagen Cat no: QT00079247 Zebrafish ef1a ACCTACCCTCCTCTTGGTCG GGAACGGTGTGATTGAGGGAA Zebrafish gpr35.1 TTTGAACAGGGCTTTGCGATG GGTCCATTGGGTTTTGGGAC Zebrafish gpr35.2 TTGCTCCACACAAACCGTCT ATATGAACCGGCGTGAAGCA Zebrafish il17 CGCCTTGGACATACACAACTT AGTAAATGGGTTGGGACTCCA Zebrafish tnfa GGAGAGTTGCCTTTACCGCT TTGCCCTGGGTCTTATGGAG Zebrafish il1b ATCAAACCCCAATCCACAGAGT GGCACTGAAGACACCACGTT Zebrafish il22 CGATGACTGATACAGCACGA TGTGCTCGTCTGATTCCAAG Zebrafish il10 TAAAGCACTCCACAACCCCA GACCCCCTTTTCCTTCATCTTTT ATTACCGCGGCTGCTGGC ACTCCTACGGGAGGCAGCAGT 16S bacterial rRNA C Mouse Gpr35 ACAAGGCAGGAACTGTG CCTAGGGCTCAGGCAGC
(Sweden) Genotyping Primers Gpr35-dtTomato Gpr35-KO GCCTGGATGCCATCTGTTACTACTAC GCAAGGCCCCAACATCTATAGCTCA GATGCAGCCTCTCTAGTCCAACTG CACTGTCTTTTGTTGCTGCTGCTGT GGTCGCTACAGACGTTGTTTGTC TGGGTTTGGCCCTTAGGATGATGTG Gpr35-flox GCAAGGCCCCAACATCTATAGCTCA CACTGTCTTTTGTTGCTGCTGCTGT TGGGTTTGGCCCTTAGGATGATGTG GTGGCAGACCATTCCGAAGCTAGAG
86 Appendix
Swiss IBD Cohort Investigators
Karim Abdelrahman2, Gentiana Ademi3, Patrick Aepli4, Claudia Anderegg5, Anca-Teodora Antonino6, Eva Archanioti26, Eviano Arrigoni7, Diana Bakker de Jong5, Bruno Balsiger8, Polat Bastürk5, Peter Bauerfeind9, Andrea Becocci10, Dominique Belli10, José M. Bengoa7, Luc Biedermann11, Janek Binek12, Mirjam Blattmann11, Stephan Boehm13, Tujana Boldanova5, Jan Borovicka3, Christian P. Braegger14, Stephan Brand3, Lukas Brügger15, Simon Brunner5, Patrick Bühr14, Sabine Burk11, Bernard Burnand16, Emanuel Burri17, Sophie Buyse18, Dahlia-Thao Cao19, Ove Carstens15, Dominique H. Criblez4, Sophie Cunningham7, Fabrizia D’Angelo20, Philippe de Saussure7, Lukas Degen3, Joakim Delarive21, Christopher Doerig22, Barbara Dora11, Susan Drerup23, Mara Egger16, Ali El- Wafa24, Matthias Engelmann25, Jessica Ezri26, Christian Felley27, Markus Fliegner28, Nicolas Fournier16, Montserrat Fraga26, Yannick Franc16, Remus Frei3, Pascal Frei29, Michael Fried11, Florian Froehlich30, Raoul Ivano Furlano31, Luca Garzoni32, Martin Geyer33, Laurent Girard7, Marc Girardin34, Delphine Golay16, Ignaz Good35, Ulrike Graf Bigler15, Beat Gysi36, Johannes Haarer3, Marcel Halama37, Janine Haldemann8, Pius Heer38, Benjamin Heimgartner15, Beat Helbling29, Peter Hengstler12, Denise Herzog39, Cyrill Hess4, Roxane Hessler22, Klaas Heyland40, Thomas Hinterleitner41, Claudia Hirschi25, Petr Hruz1, Pascal Juillerat15, Stephan Kayser42, Céline Keller43, Carolina Khalid-de Bakker1, Christina Knellwolf(-Grieger)3, Christoph Knoblauch44, Henrik Köhler5, Rebekka Koller14, Claudia Krieger(-Grübel)3, Patrizia Künzler3, Rachel Kusche5, Frank Serge Lehmann38, Andrew J. Macpherson15, Michel H. Maillard26,43, Michael Manz1, Astrid Marot16, Rémy Meier45, Christa Meyenberger3, Pamela Meyer3, Pierre Michetti26,43, Benjamin Misselwitz11, Patrick Mosler46, Christian Mottet47, Christoph Müller48, Beat Müllhaupt11, Leilla Musso16, Michaela Neagu49, Cristina Nichita50, Jan H. Niess1, Andreas Nydegger26,43, Nicole Obialo11, Diana Ollo27, Cassandra Oropesa27, Ulrich Peter40, Daniel Peternac51, Laetitia Marie Petit27, Valérie Pittet16, Daniel Pohl11, Marc Porzner52, Claudia Preissler53, Nadia Raschle11, Ronald Rentsch54, Sophie Restellini27, Alexandre Restellini34, Jean-Pierre Richterich4, Frederic Ris27, Branislav Risti55, Marc Alain Ritz56, Gerhard Rogler11, Nina Röhrich3, Jean- Benoît Rossel16, Vanessa Rueger14, Monica Rusticeanu15, Markus Sagmeister58, Gaby Saner8, Bernhard Sauter59, Mikael Sawatzki3, Michael Scharl11, Martin Schelling3, Susanne Schibli60, Hugo Schlauri61, Dominique Schluckebier27, Sybille Schmid(- Uebelhart)15, Daniela Schmid44, Jean-François Schnegg62, Alain Schoepfer26,43, Vivianne Seematter16, Frank Seibold8, Mariam Seirafi63, Gian-Marco Semadeni3, Arne Senning14,
87 Appendix
Christiane Sokollik60, Joachim Sommer16, Johannes Spalinger4,60, Holger Spangenberger64, Philippe Stadler65, Peter Staub66, Dominic Staudenmann4, Volker Stenz67, Michael Steuerwald56, Alex Straumann38, Bruno Strebel15, Andreas Stulz4, Michael Sulz3, Aurora Tatu15, Michela Tempia-Caliera68, Amman Thomas57, Joël Thorens69, Kaspar Truninger70, Radu Tutuian15, Patrick Urfer71, Stephan Vavricka11, Francesco Viani72, Jürg Vögtlin56, Roland Von Känel11, Dominique Vouillamoz73, Rachel Vulliamy16, Paul Wiesel50, Reiner Wiest15, Stefanie Wöhrle4, Tina Wylie74, Samuel Zamora34, Silvan Zander83, Jonas Zeitz11, Dorothee Zimmermann3
1University Center for Gastrointestinal and Liver Diseases, St. Clara Hospital and University Hospital of Basel, 3223 Basel, Switzerland; 2Clinique de Montchoisi, Lausanne, Switzerland; 3Kantonsspital St. Gallen, St. Gallen, Switzerland; 4Kantonsspital Luzern, Luzern, Switzerland; 5Kantonspital Aarau, Klinik für Kinder und Jugendliche, Aarau, Switzerland; 6Hôpital Riviera–Site du Samaritain, Vevey, Vaud, Switzerland; 7GI private practice, Geneva, Switzerland; 8Gastroenterologische Praxis, Bern, Switzerland; 9Department Gastroenterology and Hepatology, Stadtspital Triemli, Zurich, Switzerland; 10Department of Pediatric, Geneva University Hospital, Geneva, Switzerland; 11Department of Gastroenterology and Hepatology, University Hospital Zurich, University of Zurich, Zurich, Switzerland; 12Gastroenterologie am Rosenberg, St. Gallen, Switzerland; 13Spital Bülach, Bülach, Zurich, Switzerland; 14University Children’s Hospital, Zurich, Switzerland; 15Department of Visceral Surgery and Medicine, Bern University Hospital, University of Bern, Bern, Switzerland; 16Institute of Social and Preventive Medicine (IUMSP), Lausanne University Hospital, Lausanne, Switzerland; 17Department Gastroenterology, Kantonsspital Liestal, Liestal, Switzerland; 18GI private practice, Yverdon-les-Bains, Switzerland; 19Hôpital Neuchâtelois, La Chaux-de-fonds, Neuchâtel, Switzerland; 20Department Gastroenterology and Hepatology, Geneva University Hospital, Geneva, Switzerland; 21GI private practice, Lausanne, Switzerland; 22Clinique Cecil, Lausanne, Switzerland; 23Schulthess Clinic, Zurich, Switzerland; 24GI private practice, La Chaux-de-Fonds, Switzerland; 25Gastropraxis Luzern, Luzern, Switzerland; 26Service of Gastroenterology and Hepatology, Department of Medicine, Centre Hospitalier Universitaire Vaudois and University of Lausanne, Lausanne, Switzerland; 27Centre de Gastroentérologie Beaulieu SA, Geneva, Switzerland; 28Medical Center Sihlcity, Zurich, Switzerland; 29Gastroenterologie Bethanien, Zurich, Switzerland; 30Hospital of the Canton of Jura, Porrentruy And Delémont, Jura, Switzerland; 31Universitäts-Kinderspital beider Basel (UKBB), Basel, Switzerland; 32Clinique des
88 Appendix
Grangettes, Geneva University Hospital, Genève, Switzerland; 33GI private practice, Wettingen, Aargau, Switzerland; 34Groupe Médical d’Onex, Onex, Switzerland; 35Spital Walenstadt, Walenstadt, St. Gallen, Switzerland; 36GI private practice, Reinach, Switzerland; 37Aerztehaus Fluntern, Zurich, Switzerland; 38GI private practice, Olten, Switzerland; 39HFR Hôpital fribourgeois–Pédiatrie, Fribourg, Switzerland; 40KSW Kantonsspital Winterthur Kinderklinik, Winterthur, Switzerland; 41GI private practice, Zurich, Switzerland; 42GI private practice, Luzern, Switzerland; 43Gastroenterology La Source-Beaulieu, Lausanne, Switzerland; 44Kantonsspital Nidwalden, Stans, Nidwalden, Switzerland; 45AMB – Arztpraxis MagenDarm Basel, Basel, Switzerland; 46Kantonsspital Graubünden, Chur, Switzerland; 47GI private practice, Sion, Switzerland; 48Division of Experimental Pathology, Institute of Pathology, University of Bern, Bern, Switzerland; 49Spital Tiefenau, Bern, Switzerland; 50Centre médical d’Epalinges, Epalinges, Switzerland; 51Spital Waid, Zurich, Switzerland; 52Spital Lachen, Lachen, Switzerland; 53Kantonsspital Olten, Olten, Switzerland; 54GI private practice, St. Gallen, Switzerland; 55GI practice, Dietikon, Switzerland; 56GI practice, Liestal, Switzerland; 57GI private practice, Waldkirch, St. Gallen, Switzerland; 58GI private practice, Heerbrugg, Switzerland; 59Klinik Hirslanden Zürich, Zurich, Switzerland; 60Kinderklinik Bern, Bern University Hospital, Bern, Switzerland; 61Derby Center, Wil, Switzerland; 62GI private practice, Montreux, Switzerland; 63Clinique La Colline, Geneva, Switzerland; 64Kantonsspital Wolhusen, Wolhusen, Switzerland; 65GI private practice, Payerne, Switzerland; 66Spital Heiden Appenzell Ausserrhoden, Heiden, Switzerland; 67Kantonsspital Münsterlingen, Münsterlingen, Switzerland; 68Clinique des Grangettes, Chêne-Bougeries, Switzerland; 69GI private practice, Yverdon, Switzerland; 70GI private practice, Langenthal, Switzerland; 71Hirslanden Klinik Aarau, Gastro Zentrum, Aarau, Switzerland; 72Private practice, Vevey, Switzerland; 73Private practice, Pully, Switzerland; 74Infirmière de Recherche chez CHUV Lausanne University Hospital, Lausanne, Switzerland; 83Spital Limmattal, Schlieren, Switzerland
89 Appendix
6.2 List of Tables and Figures
6.2.1 List of Tables
Table 2.1 Patient Characteristics of Basel IBD Cohort for Biopsies Obtained for
Immunofluorescence Staining
Table 2.2 Group characteristics of IBD patients analyzed for the GPR35T108M SNP variant and TNF blocker therapy response
Table 2.3 Mouse mononuclear phagocytes panel
Table 2.4 Mouse lymphocytes panel
Table 2.5 Mouse innate lymphoid cells panel
Table 3.1 Responses to anti-TNF blockers in IBD patients with GPR35T108M variant
Table 6.1 Primer list
6.2.2 List of Figures
Figure 1.1 Intestinal epithelial cells in innate immunity
Figure 1.2 Monocytes, macrophages and dendritic cells in the intestinal lamina propria
Figure 1.3 Metabolite-sensing GPCRs on intestinal immune cells
Figure 1.4 Factors in IBD pathogenesis
Figure 2.1 Construct of Gpr35-IRES-tdTomato and PCR products from the genotyping
Figure 2.2 Construct of Gpr35-knock out and PCR of the genotyping
Figure 2.3 Construct of Gpr35-flox and PCR of the genotyping
Figure 2.4 Construction of gpr35buu1892 mutant zebrafish
Figure 3.1 gpr35b mRNA levels are pronounced in the zebrafish intestine at 120 hpf
Figure 3.2 Gastrointestinal tract shows high levels of Gpr35 transcripts in human and mouse
Figure 3.3. IECs and CX3CR1+ macrophages are GPR35+
Figure 3.4. Characterization of immune cells for their GPR35 expression
Figure 3.5 Percentages of GPR35+ cells decrease from monocytes to macrophages
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Figure 3.6 Colonic lamina propria macrophages can be distinguished into GPR35+ and
GPR35- populations
Figure 3.7 Gpr35 is downregulated in antibiotics-treated and GF animals
Figure 3.8 Inflammation leads to an increase in GPR35-expressing immune cells in intestinal LP
Figure 3.9 Bacterial inflammation causes increased GPR35
Figure 3.10 GPR35+ lamina propria cells are present in greater numbers in inflamed sites
of UC
Figure 3.11 LPA reduces forskolin-induced cAMP accumulation in GPR35-transfected cells
Figure 3.12 LPA leads to upregulation of tnf in WT but not in GPR35-deficient zebrafish
Figure 3.13 GPR35 mediates LPA signaling in mouse BMDMs
Figure 3.14 WT but not Gpr35-/- BMDMs show increased p-ERK upon LPA treatment
Figure 3.15 LPA mediates GPR35-dependent Tnf induction in mouse BMDMs
Figure 3.16 Macrophages show chemotaxis towards LPA in a GPR35-dependent fashion
Figure 3.17 Intestinal inflammation results in higher Atx expression
Figure 3.18 DSS colitis is associated with increased LPA production independently of
GPR35
Figure 3.19 GPR35-deficient mice show worsened signs of DSS colitis
Figure 3.20 Tamoxifen reduces the percentage of GPR35+ cells in colonic macrophages of Gpr35ΔCx3cr1 mice
Figure 3.21 GPR35-deficiency in macrophages increases DSS colitis severity
Figure 3.22 LPA alleviates DSS colitis severity in WT but not in Gpr35ΔCx3cr1 mice
Figure 3.23 Macrophage-specific GPR35 deletion results in a higher number of neutrophils during colitis
91 Appendix
Figure 3.24 GPR35-deficiency leads to lower TNF levels in macrophages during DSS
colitis
Figure 3.25 TNF administration attenuates colitis in Gpr35ΔCx3cr1 mice
Figure 3.26 TNF restores corticosterone synthesis in colitic Gpr35ΔCx3cr1 mice
Figure 3.27 TNF did not show a significant effect on DSS colitis in WT mice
Figure 4.1 Potential scheme for a mechanism where corticosterone induction by TNF is partially mediated by GPR35
92 Appendix
6.3 Acknowledgements
First and foremost, I would like to thank my supervisor Jan Niess for giving me the opportunity to work on this exciting and challenging project. Thank you very much for pushing me to always try my best, and for the long discussions, your patience, and endless support.
I would like to express my sincere gratitude to the committee members, Christoph
Hess and Christian Riedel for their guidance and constructive criticism which always encouraged me to improve myself.
Many thanks to Eduardo Villablanca and Cristian Doñas for their most valuable collaboration. I would also like to thank Florian Geier for the bioinformatics analysis,
Philippe Demougin and Christian Beisel for RNA sequencing.
My deepest gratitude for Christine Bernsmeier and my lab colleagues, Hassan,
Philipp, Tanay, Korcan, Anne, Andrijana and Emilio. Thank you for making our workspace so comfortable, fun and sometimes so noisy that I had to go to the computer room or the library to find some peace and quiet for writing. All jokes aside, thanks for being by my side, listening to me, and motivating me.
Special thanks to my family, my mom, dad and my sister for loving me unconditionally and always supporting me.
Through success and failure, thick and thin, shortest to longest experimental protocols, I have enjoyed every sweat, tear and laugh in these 4 years. Thanks to everyone I’ve crossed paths with during this incredible journey.
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