Characterization of cytokine secretion in response to soluble and crystalline oxalate in immune cells
Charakterisierung der Zytokinsekretion von Immunzellen nach Stimulierung mit löslichem und kristallinem Oxalat
Der Medizinischen Fakultät der Friedrich-Alexander-Universität
Erlangen-Nürnberg zur
Erlangung des Doktorgrades Dr. med.
vorgelegt von Louise Maida Tonner
Als Dissertation genehmigt von der Medizinischen Fakultät
der Friedrich-Alexander-Universität Erlangen-Nürnberg
Tag der mündlichen Prüfung: 16.12.2020
Vorsitzender des Promotionsorgans: Prof. Dr. med. Markus Neurath Gutachter/in: Prof. Dr. med. Felix Knauf
Prof. Dr. rer. nat. Christoph Becker
Table of content
1. Zusammenfassung 1
1.1 Hintergrund und Ziele 1
1.2 Methoden 1
1.3 Ergebnisse 2
1.4 Schlussfolgerung 2
2. Abstract 4
2.1 Background 4
2.2 Methods 4
2.3 Results 5
2.4 Conclusion 5
3. Introduction 7
3.1 Chronic kidney disease 7
3.1.1 Background 7
3.1.2 Epidemiology and etiology 8
3.2 Oxalate 9
3.2.1 Hyperoxalemia and systemic oxalosis 10
3.2.2 Oxalate transporters 12
3.3 Inflammatory cells and kidney disease 12
3.3.1 Macrophages and kidney disease 12
3.3.2 Dendritic cells and kidney disease 13
3.3.3 Inflammasome in CKD 14
3.4 Aim of the study 16
4. Material 17
4.1 Software 17
4.2 Statistical analysis 17
4.3 Buffers 17
4.3.1 Buffers for cell culturing 17
4.3.2 Buffers used for Western Blot analysis 18
5. Methods 21
5.1 Animal handling and care 21
5.2 Bone marrow derived cell isolation (dendritic cells and macrophages) 21
5.2.1 Dendritic cell differentiation 22
5.2.2 Macrophage differentiation 22
5.3 Bone marrow derived cell stimulation 22
5.4 Peripheral blood mononuclear cell isolation 23
5.5 Enzyme- linked Immunosorbent Assay 24
5.5.1 Interleukin-1 alpha ELISA 24
5.5.2 Interleukin-1 beta ELISA 25
5.6 Protein isolation 26
5.7 Protein concentration measurement 27
5.8 Western Blot 27
5.9 RNA isolation 28
5.10 RNA concentration measurement 29
5.11 Complementary DNA synthesis by reverse transcription 29
5.12 Real-time quantitative polymerase chain reaction 30
5.13 Cell viability assay 31
5.14 Gene deficient Slc26a6-/- mice 31
6. Results 33
6.1. Basic characterization of IL-1α and IL-1β release in murine DCs 33 6.1.1 Dose dependent measurements with crystalline CaOx on murine DCs 33 6.1.2 Time dependent measurements with crystalline CaOx on murine DCs 34 6.2 Characterization of IL-1α and IL-1β release from murine DCs upon stimulation with soluble sodium
oxalate 35
6.2.1 Dose dependent measurements with soluble NaOx on murine DCs 36 6.2.2 Time dependent measurements with soluble NaOx on murine DCs 37 6.3 Does oxalate in concentrations as found in the plasma of ESRD patients induce inflammation? 37
6.4 Long time incubation of murine DCs with NaOx 38
6.5 Characterization of IL-1α and IL-1β release in murine macrophages 39
6.6 Characterization of IL-1α release in human DCs 40
6.7 Ligands experiment 41
6.8 NLRP3 expression after stimulation with NaOx by use of qPCR 42
6.9 Physiological role of oxalate transporter SCL26A6 43
6.9.1 SLC26A6 expression on human DCs 44
6.9.2 Testing a hypothesis of enhanced inflammasome activation by elevated oxalate in Slc26a6-/-
macrophages 44
6.9.4 Comparison of macrophage cell viability of cells from WT and Slc26a6-/- mice following
incubation with soluble oxalate 47
7. Discussion 48
7.1 Discussion of Results 48
7.2 Limitation of this Study 50
7.3 Future prospects 51
8. List of Abbreviations 53
9. Reference List 56
10. Publications 63
11. Acknowledgements 64
1. Zusammenfassung
1.1 Hintergrund und Ziele
Die chronische Nierenerkrankung stellt mit einer globalen Prävalenz von rund 15% eine bedeutende Herausforderung für die betroffenen Patienten und die öffentlichen Gesundheitssysteme dar. Ein genaueres Verständnis der zugrundeliegenden Pathophysiologie wird benötigt, um neue präventive und therapeutische Ansätze zu entwickeln und damit der steigenden Prävalenz entgegen steuern zu können. Eine viel beschriebene Komponente in der Pathophysiologie der chronischen Nierenerkrankung ist die persistierende Entzündung. Patienten mit dialysepflichtiger chronischer Nierenerkrankung, unabhängig von deren Ätiologie, weisen erhöhte Plasmaoxalatkonzentrationen von 30-60 µmol/l auf. Im Vergleich dazu liegen die Werte in gesunden Menschen nur zwischen 1-3 µmol/l. Die Übersättigung von Plasmaoxalat kann zur Ablagerung von Kalziumoxalatkristallen in multiplen Organen führen, dies geschieht ab Konzentrationen von ca. 30 µmol/l. Es ist bekannt, dass die Kalziumoxalatkristalle das NOD- Like Receptor Protein 3 Inflammasom aktivieren können, wodurch es zur Freisetzung von proinflammatorischen Zytokinen und damit zur systemischen Inflammation kommen kann. Der vorliegenden Arbeit liegt die Hypothese zu Grunde, dass erhöhte Plasmaoxalatkonzentrationen, auch ohne vorangehende Präzipitation zu Kristallen, eine Inflammation begünstigen. Ob das erhöhte lösliche Oxalat in den Konzentrationen, in denen es im Plasma von Dialysepatienten gemessen werden konnte, einen Effekt auf das Immunsystem hat, ist nicht geklärt und soll in der vorliegenden Arbeit weiter untersucht werden.
1.2 Methoden
Die Hypothese wurde mittels in-vitro Zellstimulationen überprüft. Dafür wurden sowohl murine als auch humane Immunzellen isoliert und mit steigenden Konzentrationen an Kalziumoxalat und Natriumoxalat stimuliert. Die Konzentrationen wurden dabei den Plasmaoxalatkonzentrationen chronisch Nierenkranker angepasst, um somit die Hyperoxalämie der Patienten zu imitieren.
Nach verschiedenen Zeitpunkten der Stimulation wurde der Zellüberstand abgenommen.
Anschließend wurde die Konzentration an Interleukin-1 alpha und Interleukin-1 beta im
Zellüberstand mithilfe von Enzyme-linked Immunosorbent Assay Messungen bestimmt.
Zusätzlich wurden verschiedene Kofaktoren zur Stimulierung hinzugezogen, um die Vielfalt der urämischen Bedingungen im Plasma eines Patienten mit chronischer Niereninsuffizienz zu simulieren. In weiteren Experimenten wurde der Oxalattransporter Slc26a6 und seine Funktion in Immunzellen untersucht. Dazu wurde zunächst seine Expression in humanen Dendritischen Zellen geprüft. Weiter wurden Vergleiche zwischen der Zytokinausschüttung von Immunzellen aus Wildtyp und aus Slc26a6-/- Mäusen nach Stimulierung mit Oxalat aufgestellt. Zuletzt wurde mithilfe eines Zellviabilitätsassays die Mortalität von Immunzellen aus Wildtyp- und aus Slc26a6-/- Mäusen, nach Stimulierung mit Oxalat, verglichen.
1.3 Ergebnisse
Mittels Enzyme-linked Immunosorbent Assay Messungen konnte nach Stimulierung muriner und humaner Immunzellen mit Calcium- und Natriumoxalat eine Freisetzung von sowohl Interleukin- 1 alpha als auch Interleukin-1 beta gemessen werden. Dabei konnte ein zeit- sowie dosisabhängiger Anstieg der Zytokinfreisetzung aufgezeigt werden.
Eine Zytokinfreisetzung nach Stimulierung mit Oxalatkonzentrationen von ca. 45 µmol/l, wie sie in chronisch Nierenkranken gemessen wurden, konnte in dieser Arbeit nicht gezeigt werden. Ein Vergleich der Prästimulierung zwischen dem Toll like Receptor 4 Liganden Lipopolysaccharid sowie dem Toll like Receptor 3 Liganden Polyinosinic: polycytidylic acid hat ergeben, dass beide Kofaktoren als potente Prästimuli für die Oxalat-abhängige Ausschüttung von Interleukin-1 alpha wirken. In weiteren Experimenten konnte die Expression des Oxalattransporter SLC26A6 in humanen Dendritischen Zellen nachgewiesen werden.
Ein Vergleich der Mortalität von Zellen aus Wildtyp Mäusen im Vergleich zu Zellen aus Slc26a6-/- Mäusen, nach Stimulierung mit Natriumoxalat, konnte eine erhöhte Mortalität der Zellen aus Slc26a6-/- Mäusen aufzeigen.
1.4 Schlussfolgerung
Die pro-inflammatorische Aktivität von Oxalat konnte sowohl an murinen als auch an humanen Zellen bestätigt werden. Diese konnte weiter charakterisiert werden, indem ein dosis- und zeitabhängiger Verlauf der Zytokinausschüttung der Zellen nach Stimulierung mit Calcium- und
Natriumoxalat gezeigt werden konnte. Die in der vorliegenden Arbeit präsentierten Ergebnisse untersuchen damit den potentiellen Einfluss von erhöhten Plasmaoxalatwerten auf die Entstehung der systemischen Entzündungsreaktion in Patienten mit chronischer Nierenerkrankung. Weitere Experimente sind jedoch nötig, um in vivo eine pro-inflammatorische Wirkung von niedrigen Oxalatkonzentrationen, wie sie in chronisch Nierenkranken vorkommen, darzustellen.
Die vorliegende Arbeit zeigt darüber hinaus eine mögliche physiologische Rolle des Slc26a6 Transporters in Makrophagen auf. Durch einen Vergleich der Mortalität von Zellen aus Slc26a6-/- Mäusen mit Zellen aus Wildtyp Mäusen konnte gezeigt werden, dass die Zellen aus Wildtyp Mäusen nach Stimulierung mit Natriumoxalat eine niedrigere Mortalität aufweisen als die Zellen aus Slc26a6-/- Mäusen. Diese Erkenntnis führt zu der Annahme, dass der Slc26a6 Transporter, unter hyperoxalämischen Bedingungen Makrophagen vor der Überfrachtung mit löslichem Oxalat schützt. Die in der Arbeit präsentierten Ergebnisse geben Anstoß zu weiteren Untersuchungen der pro-inflammatorischen Auswirkungen von Oxalat. Ein detaillierteres Verständnis könnte beispielsweise zu einem verbesserten Patientenmonitoring sowie zu neuen anti-inflammatorischen Therapieansätzen führen.
2. Abstract
2.1 Background
With a global prevalence of around 15%, chronic kidney disease is a significant challenge for affected patients and the health care systems worldwide. In order to further develop new therapeutic approaches and to counteract the continuously increasing prevalence, a better understanding of the underlaying pathophysiology is needed. One well known component in the pathophysiology of chronic kidney disease is the persistent inflammation. Patients with end-stage renal disease and on hemodialysis, regardless of their individual pathological background, tend to exhibit highly elevated plasma oxalate levels, ranging around 30-60 µmol/l. In comparison, healthy individuals exhibit levels around 1-3 µmol/l. The resulting supersaturation of oxalate that occurs when the plasma concentration is above 30 µmol/l, leads to crystal deposition in various organs. Oxalate crystals have been shown to activate the NOD-Like Receptor Protein 3 Inflammasome, resulting in the release of proinflammatory cytokines and with that in systemic inflammation. The presented work deals with the hypothesis that elevated plasma oxalate, even without precipitation to crystals, contributes to systemic inflammation. Whether or not the elevated plasma oxalate, in concentrations as measured in dialysis patients’ plasma has an effect on the immune system, remains unknown and shall be clarified in this work.
2.2 Methods
The hypothesis has been tested with use of in vitro cell-stimulations. Murine as well as human cells were isolated and stimulated with various concentrations of calcium oxalate and sodium oxalate. Oxalate concentrations were set in the range of the plasma oxalate concentrations of patients with chronic kidney disease in order to mimic their hyperoxalemic setting. Supernatant was taken at different time points. Interleukin-1 alpha and Interleukin-1 beta concentrations in the supernatant were measured using an Enzyme-linked Immunosorbent Assay. Subsequent to that, cofactors of stimulation were tested in order to further simulate the uremic conditions. In a consecutive series of experiments, the SLC26A6 oxalate transporter was examined for expression in human dendritic cells. Next, cytokine secretion of cells from both wild-type and Slc26a6-/-
mice were compared after stimulation with oxalate. Finally, mortality of immune cells from wild- type and Slc26a6-/- mice after stimulation with oxalate was compared by use of a cell viability assay.
2.3 Results
Using Enzyme-linked Immunosorbent Assay measurements we were able to measure Interleukin- 1 alpha and Interleukin-1 beta in the supernatants of murine and human cells after stimulation with oxalate. As part of these measurements, a dose- as well as a time-dependent secretion of Interleukin-1 alpha and Interleukin-1 beta could be detected. Yet, we were not able to show cytokine release after stimulation of oxalate in a concentration of around 45 µmol/l, as seen in patients with CKD. Through the comparison of prestimulation with the Toll like Receptor 4 ligand Lipopolysaccharid and Toll like Receptor 3 ligand Polyinosinic:polycytidylic acid, we were able to demonstrate both cofactors as potent prestimuli of oxalate-dependent secretion of Interleukin-1 alpha. In a next series of experiments, the expression of oxalate transporter SLC26A6 in human dendritic cells was detected. A comparison of the mortality between cells from wild-type and Slc26a6-/- mice after stimulation with oxalate, was able to demonstrate less cell viability of the cells from the Slc26a6-/- mice.
2.4 Conclusion
This work confirmed the hypothesis that soluble oxalate has a pro-inflammatory effect on murine as well as human cells. It further characterizes the cytokine release of these cells by revealing a dose- and time dependent course of the release in reaction to stimulation with oxalate. Thus, the results presented in this work point out the potential impact of elevated plasma oxalate concentrations on the progression of systemic inflammation in patients with chronic kidney disease. Yet, further investigation is needed to illustrate the proinflammatory activity of slightly elevated oxalate concentration as measured in the plasma of patients with chronic kidney disease.
Furthermore, this work shows a potential physiological role of the Slc26a6 transporter in macrophages. The comparison of the viability of macrophages from wild-type- and Slc26a6-/- mice after stimulation with soluble oxalate, was found to cause significantly greater loss of viability of the macrophages from Slc26a6-/- mice. This may lead to the insight that the
transporter is able to protect macrophages from the overload of soluble oxalate in an hyperoxalemic setting. The results presented in this work calls for further research on the pro- inflammatory effect of oxalate. A better understanding of the underlying pathophysiology could then lead to an improved patient management as well as to new anti-inflammatory therapeutic approaches.
3. Introduction
3.1 Chronic kidney disease
3.1.1 Background
The kidney unites excretory, endocrine and metabolic functions. Besides that, the kidney is primarily responsible for maintaining the composition of the body fluids.
In total both kidneys consist of around two million nephrons that are each composed of one glomerulus and its duct system. The glomeruli filter the blood plasma by removing excessive and metabolic by-products from it, thus regulating the electrolyte- water- mineral and acid-base balance of the body. Each glomerulus contributes to the total glomerular filtration rate (GFR) which eventually serves as an overall index of all kidney functions. The GFR estimates the volume of liquid filtered out by the glomeruli into the urine within the time span of one minute.
In clinical daily routine, the assessment of creatinine clearance can help to estimate the GFR. In healthy individuals the GFR ranges around 90-110 ml/min/1.73m2 depending on gender and body mass index (BMI). With advancing age, the GFR decreases progressively, either physiologically or pathologically, due to kidney injury of any etiology. Chronic kidney disease (CKD) is defined by the Kidney Disease: Improving Global Outcomes (KDIGO) as “abnormalities of kidney structure or function, present for >3 months, with implications for health” [2]. Depending on the GFR, CKD can be classified into 5 stages, outlined in Figure 1 below. During the course of progressive loss of kidney function, the kidney has the ability to maintain GFR through manifestation of hyperfiltration and compensatory hypertrophy within the remaining healthy nephrons. Therefore, the estimated GFR allows reliable detection of kidney damage only at a substantial decline of kidney function of greater than 50%. This regulatory mechanism makes clinical assessment of kidney disease challenging and demands for other markers of early kidney damage. Such as the serum creatinine, blood urea nitrogen as well as the assessment of proteinuria or the examination of urinary sediment.
Stage Description GFR
ml/min/1.73m2
1 Kidney damage with normal or
increased GFR
>90
2 Kidney damage with mild
decrease in GFR
60-89
3 Moderate decrease in GFR 30-59
4 Severe decrease in GFR 15-29
5 Kidney failure <15
Figure 1: Stages of kidney disease (modified after Clinical Practice Guidelines for Chronic Kidney Disease: Evaluation, Classification and Stratification [3]).
Patients with a GFR <60 ml/min/1.73m2 for more than 3 months are defined as having CKD.
3.1.2 Epidemiology and etiology
CKD is a global public health care problem with sustained incidence and prevalence [4].
According to the United States Renal Data System (USRDS) around 15% of the adult general population in the United States is affected by CKD [6, 7]. That translates to approximately 30 million people, making it a severe health care burden. The disease occurs in the entire population but affects disproportionally more people of lower income as well as ethnic minorities [4].
Among the 5 stages of CKD classification, the earlier stages, most notably stage 3, are the most prevalent while stage 5, end-stage renal disease (ESRD), that is renal replacement therapy requiring, has a comparably low prevalence.
CKD can arise from various underlying kidney diseases such as an acute kidney injury or develop from an existing kidney disease. One of the main causes of CKD today is diabetes mellitus (DM).
Up to 30-40% of all CKD patients have an underlying DM of either type I or II [8]. As metabolic diseases are on the rise especially in the developed countries, CKD caused by DM is a major cause for the steady rise of prevalence of CKD [9, 10]. Other common causes for CKD are hypertension, polycystic kidney diseases as well as glomerulonephritis or interstitial nephritis.
Despite the high prevalence of CKD, the public awareness concerning risk factors for developing CKD, its symptoms and its long-term effects are widely unknown. The loss of kidney function in CKD leads to several pathogenic conditions such as: hypertension and arrhythmia, osteomalacia
due to vitamin D3 deficiency, metabolic acidosis, intestinal dysbiosis and accumulation of uremic toxins. Given these facts, it becomes obvious how CKD is immensely lowering quality of life and causing premature morbidity and mortality. Regardless of the etiology of the injury in the kidney that leads to CKD, chronic inflammatory processes can commonly be detected in affected individuals [11-19]. In this setting, multiple factors such as the uremic milieu, elevated levels of proinflammatory cytokines and oxidative stress trigger inflammatory cells to infiltrate the affected tissue [20]. These cells, mainly neutrophils, lymphocytes and monocytes are capable of removing potential cell debris and inducing tissue-repair. By releasing pro-fibrogenic cytokines and growth factors they then enhance the progression of renal failure [21].
3.2 Oxalate
Oxalate is the ion of oxalic acid, a dicarboxylic acid, that has recently been discussed as a potent stimulus of inflammation. Oxalic acid forms salts with different cations. It then occurs in a soluble (sodium, potassium, magnesium salts) and an insoluble form that is formed in the presence of calcium (calcium salt) [22]. 60-80% of the human plasma oxalate originates from the hepatic metabolism as an end product of protein biosynthesis. The endogenous precursor molecule of oxalate is glyoxylate, synthesized through the oxidation of glycolate in the serine pathway. If the glyoxylate supply is oversaturated, it is oxidized to oxalate by either glycolate- oxidase or lactate dehydrogenase. An additional 20-40 % of the plasma oxalate derives from exogenous oxalate that is absorbed from different food such as green leafy vegetables, cocoa seeds or rhubarb in the small intestine. Intestinal oxalate absorption is mainly passive and paracellular through the tight junctions [23]. Studies have shown that the amount of oxalate absorbed in a regular diet is not harmful whereas higher amounts of ingested oxalate can cause illness, organ damage and death [24, 25]. Up to 90% of oxalate is excreted through the kidneys, which explains why plasma oxalate levels rise in declining renal function. Only an extremely low amount of oxalate is excreted via feces.
Figure 2: Oxalate metabolism (modified after Aronson PS [26]).
60-80% of plasma oxalate is generated as an end product of protein biosynthesis in the liver.
20-40% of plasma oxalate stems from dietary sources. Plasma oxalate is mainly excreted by the kidneys.
3.2.1 Hyperoxalemia and systemic oxalosis
In CKD, regardless of its etiology, plasma oxalate concentration rises due to the impairment of the renal clearance [27, 28]. This elevated plasma oxalate concentration range up to 30-60 µmol/l in patients with ESRD on hemodialysis (HD) [29, 30]. Contrary to that, in healthy individuals, plasma oxalate concentration range around 1-3 µmol/l [29, 31-33]. Elevated plasma oxalate concentration is known to cause kidney damage which then again leads to a decreasing GFR, thus creating a vicious cycle of oxalate retention and decreasing kidney function. Highly elevated plasma oxalate concentration of up to 150-200 µmol can be found in patients with either primary or enteric hyperoxaluria [29, 33]. Both are disorders of oxalate homeostasis. Primary Hyperoxaluria is an autosomal recessive disorder in which enzymatic defects lead to a massive hepatic overproduction of oxalate. In over 80% of the Primary Hyperoxaluria cases, a deficiency of alanine-glyoxylate transferase that transaminases glyoxylate to glycine is the cause. The association of disturbances in oxalate homeostasis with human diseases has been widely shown.
In the state of enteric hyperoxaluria, the paracellular and transcellular pathways of oxalate uptake
are modified, leading to an over absorption of oxalate in the intestine. These modifications are commonly described in patients with inflammatory bowel disease, ileac resection and also in patients that have undergone bariatric surgeries [22, 34]. However, there are several causes leading to elevated plasma oxalate concentration. As previously mentioned, oxalic acid is mainly excreted in the kidney where it is filtrated, secreted and reabsorbed [35]. In patients with ESRD oxalate excretion is decreased leading to elevated plasma oxalate concentration and hyperoxaluria. In conditions of hyperoxaluria, oxalate accumulates in the tissues and solid organs, especially the kidney where the supersaturation of calcium oxalate (CaOx) in the urine leads to the formation of calcium oxalate monohydrate (COM) crystals [32, 36-38]. Further, the adhesion of these crystals to the tubular epithelial cells leads to the formation of CaOx kidney stones and subsequently to inflammation and further cellular injury, causing reduced kidney function. The decrease of renal function itself causes the rising of plasma oxalate concentration due to impaired glomerular filtration, thus creating a vicious cycle of rising concentration and decreasing kidney function as mentioned above [1, 39, 40].
Figure 3: Pathway of oxalate synthesis in the liver and its enzymatic defects in primary hyperoxaluria. 80% of all primary hyperoxaluria cases are caused by a defect of the alanin glyoxylate transferase (AGT) [41, 42].
3.2.2 Oxalate transporters
The Slc26a6 transporter belongs to the phylogenetically ancient Slc26 gene family that encodes anion exchangers for HCO3−, sulfate, oxalate, I- and formate [43]. Whereas the Slc26a6 transporter was first identified as a mouse kidney protein with Cl- formate exchange activity, expression of it today is well-known in several other murine tissues such as the placenta, the esophagus, the stomach, the intestine, the pancreas, the heart and muscle tissue [43-45].
However, its renal and intestinal physiology remains the best studied to date. Measurement of epithelial uptake has shown oxalate absorption in the intestine to be mainly passive and paracellular across tight junctions. Further data attests also Slc26a3 mediated oxalate absorption [46]. The Slc26a6 transporter, however, is responsible for Cl-oxalate exchange activity in the proximal tubule where it enhances the oxalate secretion and therewith lowers the plasma oxalate.
Moreover, Slc26a6 is responsible for secretory oxalate back flux in the intestine, thus regulating the net absorption of ingested oxalate. Although research to date is still unable to detect a gene responsible for an Slc26a6 related CKD phenotype, recent data suggests a correlation of weak expression of Slc26a6 with the recurrence of oxalate nephropathy in a patient with a history of bariatric surgery [34, 47]. In addition to that, rodent studies with knockout mice models suggest Slc26a6 deficiency to be highly associated with hyperoxaluria and oxalate urolithiasis [48, 49].
These findings together strengthen the assumption of a Slc26a6 dependent oxalate secretion that serves as a protective mechanism of hyperoxaluria-induced kidney disease.
3.3 Inflammatory cells and kidney disease
3.3.1 Macrophages and kidney disease
Macrophages (MΦs) are immune cells that derive from Monocytes, which represent 5- 10% of all human peripheral-blood leukocytes. Monocytes are produced in the bone marrow and circulate in the blood prior to their differentiation. As part of the mononuclear phagocyte system, MΦs are present throughout all tissues where they are known as Kupffer cells (hepatic MΦs), Microglia (brain MΦs), Alveolar MΦs (pulmonary MΦs), Histiocytes (tissue MΦs). The term
“Macrophage” origins from the Greek language: μακρος (makros) = large, φαγειν (phagein) = to eat, and describes one of their main features: the phagocytosis and digestion of cellular debris, microbes or other exogenous substances, which gives them an important role in host defense and
tissue homeostasis. In addition to that, MΦs are able to perform other functions and responses, subsequent to stimulation by endogenous as well as exogenous triggers [50]. These responses vary from the types of the activating substances and setting and can be of both pro- and anti- inflammatory origin. Depending on these responses, MΦ phenotypes are generally subdivided in M1 (pro-inflammatory) and M2 (anti-inflammatory) MΦs. Yet, it is believed that each phenotype is able to switch into the other depending on the microenvironment. Several findings indicate the predominant heterogenic role of MΦs in the kidney. In the setting of ischemic acute kidney injury, when kidney tissue is damaged, monocytes are recruited from the circulatory system, differentiate into primarily pro-inflammatory M1 MΦs and accumulate in the injured tissue where they exacerbate the tissues’ inflammation and contribute to further tissue injury. After a while, the M1 MΦs are able to switch into M2 MΦs that secrete angiogenic and anti- inflammatory factors, pro-fibrogenetic factors and clear tissue debris.
Rodent studies with the use of fluorescent microscopy have shown that MΦs surround CaOx crystals in the kidney and later engulf and degenerate these crystals, ultimately leading to the induction of an inflammatory cascade. Thus, MΦs numbers and their state of activation have shown to be correlated to both kidney damage and fibrosis. This shows why it is important to thoroughly understand the mechanisms and pathways in order to further develop specific treatment options for patients with kidney diseases [50-59].
3.3.2 Dendritic cells and kidney disease
Dendritic cells (DCs) are stellate immune cells whose main function is to capture and present antigens to other immune cells in order to initiate and modulate the immune response. In this way, DCs can be seen as mediator cells between the innate and the adaptive immune system.
Because of this role, the ubiquitous distribution of DCs to all tissues becomes evident. Once DCs sense and capture antigens in the periphery, they endocytose and process the antigen to present it via their major histocompatibility complex (MHC) on their surface. These matured DCs migrate to lymphoid organs, where they interact with cytotoxic T cells as well as helper T cells.
Subsequent to the activation by the DCs, the T cells swarm into the tissues to either fight the pathogens directly or to interact with other cells, such as B cells for antibody release or with MΦs for cytokine production. Further, DCs produce and secrete cytokines and chemokines to further regulate infiltrating effector T cells. DCs that have not sensed antigens, stay immature and cause T cells to undergo apoptosis [60]. In the state of kidney injury, DCs among other inflammatory
cells, infiltrate the kidney and accumulate in in the renal tissue. Upon having sensed tissue debris, DCs mature and are among the first cells to produce proinflammatory chemokines and cytokines.
Studies on renal DCs have shown that kidney resistant DCs prevent excess ischemic tissue damage by shifting into anti-inflammatory signaling pathways [61, 62].
3.3.3 Inflammasome in CKD
As mentioned above, CaOx crystals can cause inflammation in the renal tissue. This inflammation can be explained by the activation of the NOD-Like Receptor Protein 3 (NLRP-3) inflammasome leading to the release of Interleukin-1 beta (IL-1β), Interleukin-1 alpha (IL-1α ) and Interleukin-18 (IL-18) [63, 64]. Inflammasomes are multi-protein platforms that can be found in the cytoplasm of both immune and non-immune cells. The inflammasome is a tightly controlled regulator of inflammation that maintains the balance between excessive systemic inflammatory processes and insufficient inflammation leading to persistent infections.
The best studied inflammasome today is the NLRP3 inflammasome. Its main components are the NLRP3 sensor protein, the apoptosis-associated speck-like protein (ASC) adaptor protein and the proinflammatory Caspase-1 [65]. Upon activation, conformational changes lead to the oligomerization of inflammasome proteins resulting in cytokine release and an inflammatory form of cell death, pyroptosis. Initially an activating trigger is needed to first upregulate the generally low NLRP3 levels. These triggers are being sensed by Toll like Receptors (TLRs) that belong to the family of pattern recognition receptors (PRRs). The activation of the TLRs leads to the transcription of Nuclear-Factor-kappa-light-chain-enhancer (NF-kB) and therewith the transcription of pro-inflammatory cytokines [66]. Today 13 mammalian TLRs were described [66, 67]. One well studied TLR is TLR-4 which is known to be mediating inflammasome activation. A potent stimulus of TLR-4 is Lipopolysaccharid (LPS), a lipopolysaccharide of the outer membrane of gram-negative bacteria. LPS is commonly used for oxalate-induced Inflammasome studies [68]. Another TLR, TLR-3 is described as part of the intracellular recognition system responding to RNA virus infection. In the event of retroviral nucleic acid replication, the receptor recognizes the intracellular dsRNA and reacts in a cell-specific inflammatory response. Various cell types are known to express TLR3 such as DCs, monocytes and natural killer cells (NK cells). A common TLR-3 ligand is Poly(I:C), a synthetic ds-RNA analogue, that was first synthesized in the 1970s.
In addition to the first signal of inflammasome activation as described above, a second signal is needed for the maturation of the proinflammatory cytokine (35kD) to its active form (17KD) through cleaving by Caspase-1. Several stimuli such as fungal, viral and bacterial substances as well as toxins and crystals can serve as second signals. Given the variety of these stimuli, a direct recognition from NLRP3 becomes unlikely. Therefore, several activation models have been established. In the case of crystals such as CaOx crystals, a recognition by NOD-Like Receptors is well established. Hence, oxalate is suggested to lead to apoptotic changes, by stimulating the production of Reactive-Oxygen-Species (ROS) and by disrupting mitochondrial function, by activating Phospatidylcholin-2-acylhydrase (Phospholipase A2), that generates fatty acids, arachidonic acids as well as lysophospholipids. NRLP3 activation is known to play a significant role in the progression of CKD. Models of Unilateral Ureter Obstruction (UUO) for example have demonstrated decreased tubulointerstitial fibrosis and tissue injury in NLRP3-/- mice.
Moreover, renal biopsies from human nondiabetic kidney disease patients, have shown a highly elevated expression of NLRP3 as compared to healthy controls. This amplified expression was positively correlated with serum creatinine [38, 69-77].
Figure 4: Model of the NLRP3-inflammasome
(modified after Ermer et al.
[1]). Oxalate crystals activate the NLRP3 inflammasome with subsequent oligomerization of the inflammasome proteins.
This activation ultimately leads to the release of IL-1β, IL-1α and IL-18.
3.4 Aim of the study
Patients with ESRD receiving maintenance HD, regardless of their individual pathological background, tend to exhibit highly elevated plasma oxalate concentration, ranging around 45 µmol/l compared to healthy individuals with plasma oxalate concentration around 1-3 µmol/l [29, 31, 32]. It has been shown that the highly elevated plasma oxalate concentration and the following supersaturation of oxalate can lead to crystal deposition in various organs [36, 37]. In recent studies my group and others were able to demonstrate a systemic correlation of oxalate induced CKD with cardiovascular fibrosis, arterial hypertension and even changes in the gut microbiota [78, 79]. Moreover, oxalate crystals have been shown to activate the NLRP3 inflammasome resulting in pyroptosis and the release of proinflammatory cytokines IL-1β and IL-18. However, whether or not and to what extent the elevated soluble oxalate concentration as presented in plasma of patients on HD have a systemic effect on our Immune system has not yet been clarified. Thus, the first aim of the study was to prove if sodium oxalate in a pathological concentration as found in ESRD patients induce cytokine release, promoting systemic inflammation. To do so, a basic characterization of cytokine secretion in response to crystalline CaOx was initially performed.
In an attempt to compare and to transfer the first results into a more physiological setting, the experiments were repeated with soluble sodium oxalate (NaOx). Therefore, concentrations of soluble oxalate, in a concentration as measured in ESRD Patients plasma were incubated with murine DCs, MΦs as well as human DCs (hDCs). Moreover, the physiological role of oxalate transporter Slc26a6 in MΦs was investigated.
Hypothesizing a protective role of Slc26a6 towards oxalate overload, MΦs of both Slc26a6-/- mice and wild-type (WT) mice were incubated with oxalate and their viability was measured by use of a cell viability assay.
4. Material
4.1 Software
EndNote X8 Thomson Reuters, New York City, NY, USA Microsoft Office 2016 Microsoft Corporation, Redmond, WA, USA GraphPad Prism 7 GraphPad Software, Inc., San Diego, CA, USA
4.2 Statistical analysis
All data are expressed as the means ± SEM. Statistical analysis was performed with GraphPad Prism 7.0. If not otherwise stated, unpaired t-tests were performed to compare two different samples. A value of p<0.05 was considered a statistically significant difference.
4.3 Buffers
4.3.1 Buffers for cell culturing R10-DC medium:
Reagent: Amount:
RPMI 89,9%
FBS 10,0%
Pen/Strep 1,0%
β-Mercaptoethanol 0,1%
Macrophage complete medium:
Reagent: Amount:
RPMI 76,5%
FCS 20,0%
Pen/Strep 1,0%
HEPES 2,5%
ACK buffer:
Reagent: Amount:
Water
NH4Cl 0,15M
KHCO3 10mM
Na2EDTA 0,1mM
PBMC cell media:
Reagent: Amount:
RPMI 98,0%
Pen/Strep 1,0%
AB Serum 1,0%
RIPA buffer:
Reagent: Amount:
NaCl 2,19 g
1M TRIS-HCL, pH 7.4 12,5 ml
500mM EDTA 2,5 ml
Sodium deoxycholate 1,25 g
Triton-X-100 2,5 ml
20% SDS 1,25 ml
Ultra pure water 185 ml
4.3.2 Buffers used for Western Blot analysis Separating gel:
Reagent
Acrylamide, 40% 15 ml
4X TRIS base 12,5 ml
10% SDS 0,5 ml
10% APS 0,3 ml
TEMED 0,07 ml
H2O II 21,6 ml
Stacking gel:
Reagent:
Acrylamide, 40% 2 ml
4x TRIS base, pH 6,8 5 ml
10% SDS 0,2 ml
10% APS 0,1 ml
TEMED 20 µl
H2O II 13 ml
Sample buffer:
Reagent:
0,5M TRIS base, pH 6,8 3 ml
Glycerol 4 ml
Bromophenol Blue Pinch
20% SDS 4 ml
H2O II 5 ml
DTT 200µl per 800µl sample buffer
Running buffer:
Reagent:
TRIS base, 250mM 60,6 g
Glycine 288 g
H2O II 1700 ml
20% SDS 5 ml in 1L 1x Running buffer diluted in
H2O II
Transfer buffer:
Reagent:
Methanol, 15% 900 ml
Glycine, 192 mM 86,4 ml
TRIS base, 25 mM 18,18 g
H2O II 5100 ml
Blocking buffer:
Reagent:
10x PBS 95 ml
Tween 20 1 ml
Dry milk 50 g
H2O II 855 ml
5. Methods
5.1 Animal handling and care
All experiments except those on Slc26a6-/- mice were performed on 8-12-week-old male C57BL/6N mice from Charles River Laboratories (Sulzfeld, Germany). The generation of Slc26a6-/- mice on a 129S background was previously described [48]. In order to match the Slc26a6-/- background, 129S6 WT mice were obtained from Taconic Biosciences, Inc. (Rensselar, NY, USA) and bred in the Yale Animal Resources Center (YARC). Mice were kept in a maximum of 5 animals per cage. Environmental conditions were kept stable with a light cycle of 12 light and 12 dark hours. Water and food were accessible at all time.
5.2 Bone marrow derived cell isolation (dendritic cells and macrophages)
Mice were anesthesized with 3% Isoflurane and oxygen flow of 1 l/minutes until stage III anesthesia of Guedel’s classification was reached. Depth of anesthesia was verified by continuous testing of plantar reflex. Mice were sacrificed by cervical dislocation and fixed on a dissecting board. Coat was sterilized using 70% Ethanol. Then the skin and peritoneum were opened with sterile surgical instruments (forceps and scissors). Femur and tibia were harvested, and tissue and muscles were removed by using scissors and scalpel. Bones were transferred in 10 ml of 1x PBS on ice. The surface of the bio safety cabinet and all required material was thoroughly disinfected with Spitacid. Bones were cut on both ends and flushed with a needle and a syringe. The floating bone marrow was resuspended in PBS until it was homogenized. The suspension was then centrifuged for five minutes with 1200 rpm. After centrifuging the supernatant was discarded.
The pellet was resuspended with ACK lysis buffer for one to two minutes in order to lyse the red blood cells. Lysis reaction was then stopped by adding PBS. Suspension was filtrated through a 70µm Pre-Separation Filter (Miltenyi Biotec, Bergisch Gladbach, Germany). Suspension was centrifuged again for five minutes with 1200 rpm. Meanwhile petri dishes (Sarstedt, Nümbrecht, Germany) were prepared with 10 ml of R10 media. Cell suspension was equally distributed into the dishes. Dishes were gently swirled to mix uniformly and checked under microscope.
5.2.1 Dendritic cell differentiation
DCs were incubated in 10 ml of R10 media and 2 ml of granulocyte macrophage colony-
stimulating factor (GMCSF) (Miltenyi Biotec GmbH, Bergisch Gladbach, Germany) for 7 days at 37°C with 6% CO2. Fresh media was added on day 1 and 4, each time by adding 10 ml of R10 and 2 ml of GMCSF. On day 8 cells were counted by use of trypan blue dye and a Neubauer chamber. Depending on the experiment and therewith the size of the wells on the plates, a stable number of cells per well was seeded into the plates. On day 9 cells were stimulated.
5.2.2 Macrophage differentiation
MΦs were incubated in 10 ml of MΦs complete media and 2 µl (20 ng/ml) Macrophage colony- stimulating factor (MCSF) (Sigma Aldrich, Darmstadt, Germany) for 5 days at 37°C with 6%
CO2. Cells started to attach to the ground of the plate shortly after. On day 2 differentiation of the cells could be seen. 5 ml of fresh media (including 20 ng/ml MCSF) was added on day 4. On day 5 dead cells which were floating in the media were taken off by removing 5 ml of medium. In order to loosen the attachment of the cells, 5 ml of PBS + EDTA was added onto the cells.
Afterwards the plates were placed in the fridge for 20 minutes. MΦs were then scratched from the plate by use of a cell scraper (Sarstedt, Nümbrecht, Germany). Cell suspension was then centrifuged and resuspended. Cells were counted with a Neubauer chamber. Depending on the experiment, a stable number of cells per well was seeded into the plates. On day 6 cells were stimulated [80].
5.3 Bone marrow derived cell stimulation
At the beginning of the experiment, all media was removed and refilled with 500 µg of LPS (Sigma Aldrich, Darmstadt, Germany) solution. This LPS solution served as a prestimulus for cell activation. LPS working concentration of (depending on experiment 10 ng/ml and 100 ng/ml per well) was reached when reconstituted in RPMI media (Thermo Fisher Scientific Inc,
Waltham, MA, USA). After three hours of prestimulation, 50 µl of medium was taken out of the well and refilled with oxalate solution. This oxalate solution was either prepared with CaOx- or NaOx (both Sigma Aldrich, St. Louis, MT, USA). Oxalate was weighed with an analytical
balance and dissolved in RPMI medium, thus, creating a stock solution of 1 mg/ml concentration,
which served as a stock for further dilutions. Time of oxalate stimulation ranged between 30 minutes and 24 hours, depending on each experiment. After the stimulation, plates were centrifuged for 5 minutes at 1200 rpm in order to exclude cells from the supernatant. Cell-free supernatant was collected into 1.5 ml Eppendorf tubes. Tubes were then stored at -20°C until use in the protein assay.
5.4 Peripheral blood mononuclear cell isolation
Leukoreduction system chambers (LRSC) from healthy donors were obtained from Transfusionsmedizin Erlangen. All following procedures were performed under a biosafety cabinet. The whole conus was carefully disinfected with Isopropanol. The conus was cut at the conical head and its apex with sterile scissors. Next the conus was put onto a 50 ml tube. After complete evacuation of the chamber into the tube, the chamber was rinsed by inserting a syringe containing 10 ml PBS. The tube was then filled up with PBS to 50 ml. 25 ml of blood was carefully loaded onto 13 ml of Lymphoflot (BioRad, Dreieich, Germany) a medium with a density of 1.077 g/ml. The tube was then centrifuged at 1500 rpm for 30 minutes in 22°C. Since mononuclear cells have a lower buoyant density than erythrocytes and polymorphonuclear leukocytes, they were isolated by centrifugation on the isosmotic medium, in which they remain at the medium interface in form of a white band. This white band was taken by a pipette and washed with 50 ml of PBS, 1mM EDTA. The suspension was again centrifuged at 1100 rpm for 15 minutes at 4°C. Supernatant was discarded, the pellet was washed with 50 ml of PBS, 1 mM EDTA. The suspension was again centrifuged at 900 rpm, 10 minutes, 4°C. Red blood cells were lysed with 2 ml of 1:10 diluted RNASE-free, ammonium chloride-based lysis buffer (BD Pharm Lyse™) (Biosciences BD, Heidelberg, Germany) for 2 minutes. At the same time, a small aliquot was taken to count the cells by use of a Neubauer chamber. After lysis, the suspension was centrifuged again at 700 rpm for 10 minutes at 4°C. In the meantime, cell culture flasks were prepared with 10 ml of DC medium. 200x 10^6 cells were added into each flask. After 1 hour of incubation (37°C with 5% CO2), cells were washed with RPMI (a little amount of RPMI was added into the flask). Flasks were swirled for around 10 times to remove the non-adherent fraction. DC medium was again added into the flasks. Flasks were finally incubated for 6 days.
5.5 Enzyme- linked Immunosorbent Assay
The Enzyme- linked Immunosorbent Assay (ELISA) is a common biochemical assay used for the measurement of specific analytes in liquid samples. It is performed on a microwell plate and consists of various steps that include sequentially added, incubated and washed liquid reagents. In a first step a primary capture antibody is added to the plate and binds specifically overnight. This capture antibody serves as an immobilized scavenger on the plated surface and is able to later adsorb the analyte of interest onto the plate. Nonspecific binding sites of the surface of the wells were then blocked with use of a blocking buffer. Cell free supernatant-samples containing the analyte of interest and standards with a specific content of the analyte were applied to the wells and incubated. During the incubation time, the analyte specifically binds to the capture antibody.
In order to detect this antibody-antigen complex, a second detection antibody conjugated to an enzyme is added. The enzyme converts the chemical substance that is added in a next step into a color signal. In order to stop this reaction a stop solution is finally added to the wells. Subsequent to this last step, the quantitative reading is performed by use of spectrophotometry which involves the reading of transmission of specific wavelengths of light through the liquid. By interpolating these optical density (OD)- values with the values obtained from the reading of the standard row, concentrations of the antigen of interest in the samples can be determined.
5.5.1 Interleukin-1 alpha ELISA
96-well ELISA plate (Thermo Fisher Scientific Inc, Waltham, MA, USA -Nunclon 96 Flat Bottom Transparent Polystyrol Catalog No.: 269620/269787/439454/442404/475094) were coated with 100 µl per well of capture antibody (rat anti-mouse IL-1α) diluted in PBS. The plate was sealed and incubated over night at room temperature (RT). On the next day the plate was washed with 400 µl per well of wash buffer (PBS with 0,05% Tween 20) and was then blocked with 300 µl of reagent diluent (1& BSA in PBS) for a minimum of 1 hour. Supernatants of DCs were applied to the wells and incubated for 2 hours at RT. A mouse IL-1α antibody was used as a standard in 8 different dilutions. Samples were stored on ice at all times. The plate was then washed and again incubated with 100 µl per well of detection antibody diluted in reagent diluent.
After this, 100 µl of enzyme (Streptavidin HRP) was added into each well and incubated for 20 minutes at RT. After washing 100 µl of substrate solution was added into each well. After 20 minutes of incubation, 50 µl of stopping solution (Sulphuric acid (2N)) was added into each well.
IL-1α signal was measured with an ELISA reader at 450 nm (wavelength correction: 570 nm and 450 nm) and ICON program.
5.5.2 Interleukin-1 beta ELISA
96-well ELISA plate (Thermo Fisher Scientific Inc, Waltham, MA, USA) was coated with 100 µl per well of anti-mouse IL-1β capture antibody diluted in coating buffer (0.1M sodium carbonat).
The plate was sealed and incubated over night at 4°. On the next day the plate was washed with 300 µl per well assay diluent (PBS+10% FBS). The plate was then blocked with 200 µl per well assay diluent and incubated for 1 hour. After another washing step, standards and samples were added and incubated for 2 hours. During the whole procedure samples were stored on ice.
Detection antibody diluted in assay diluent was then applied. After 1 hour of incubation and another washing, 100 µl of enzyme reagent diluted in assay diluent (1:250) was added into each well. Then substrate solution (Tetramethylbenzidin+ Hydrogen, 1:1) was added. After 30 minutes of incubation stop solution was added into each well. IL-1β signal was measured with an ELISA reader at 450 nm (wavelength correction: 570 nm and 450 nm) and ICON program.
5.6 Protein isolation
To extract proteins post stimulation, wells were first washed with PBS, after this RIPA Buffer (Sigma Aldrich, St. Louis, MT, USA) and protease inhibitor were added. To take off the sticking cells, wells were thoroughly scraped out with a small spatula. Sample was then pipetted into an Eppendorf vial. By use off sonication for 4 cycles cells in the samples were broken open, and Figure 5: Basic principle of a Sandwich- ELISA. A plate is first coated with a target- specific capture antibody. Blocking with assay diluent in order to prevent non-specific binding. The target protein binds to the immobilized capture antibody. Sandwich is formed by the addition of the detection antibody. Streptavidin horseradish- peroxidase is added and forms a complex with the target protein and the capture antibody. TMB substrate solution is added and reacts with the enzyme-antibody-target complex. The reaction produces a measurable signal.
DNA was destroyed. Eppendorf vials were then centrifuged for 30 minutes at 13000 rpm, 4°C.
Supernatants were collected. Lysates were stored in -80°C freezer.
5.7 Protein concentration measurement
In order to guarantee equal amounts of proteins in each well of the Western blot gel, protein concentration was measured with use of the Lowry Protein Assay (Thermo Fisher Scientific Inc, Waltham, MA, USA -Pierce™ Modified Lowry Protein Assay Kit, Catalog No.: 23240). This assay is based on a colorimetric technique that visualizes the total protein concentration by color changes. These color changes are based on a chemical reaction between copper ions with peptide bonds of proteins under alkaline conditions and the reduction of the phosphomolybdic- phosphotungistic (folin& Ciocalteu’s phenol reagent) reagent. Through the addition of folin &
Ciocalteu’s phenol reagent to the previously copper-treated proteins, chromogens with an increased absorbance are generated and maximal color changes can be seen within a short time.
All samples were referred to a standard curve with different concentrations of BSA diluted in distilled H2O. To every tissue and standard sample 1 ml of the so-called Blue reagent was added and incubated for 10 minutes at RT. The Blue reagent consist of 0,8 NaOH, to provide the needed alkaline pH, 20% SDS, CTC solution and distilled H2O. CTC solution consists of 0,2% copper sulfate pentahydrate, 0,4% potassium sodium tartrate, sodium carbonate and H2O II. Following this step, 500 µl folin reagent was added and incubated at RT for 30 minutes. The folin reagent (Sigma-Aldrich Chemie GmBH, Steinheim, Germany, product number: 1002109607, Lot:
SHBF4284V) consists of 2N phenol diluted in H2O II. The total protein concentration can be deduced from the reduced folin reagent by the aromatic residues of proteins in the solution, which were measured by absorbance at 750 nm using a spectrophotometer (Pharmacia Biotech Ultrospec 3000, GE Healthcare Life Science, Buckinghamshire, UK) [81].
5.8 Western Blot
Western blotting is a common method in molecular biology to identify specific proteins in samples of tissues and cells. Samples were diluted in loading buffer (5x Sodium Dodecyl Sulfate DSD) and a specific concentration of each sample was added onto the lanes of a gel. An electrode with a Voltage of 100 V was then applied to the gel, thus initiating the entirely negatively
charged amino acids in the sample to move through the gel towards the cathode. The gel is made of two different layers. The upper ‘stacking gel’ with a lower concentration of Acrylamide and a lower ‘separating gel’ with a high concentration of acrylamide. Due to the low concentration of the stacking layer, the gel is more porous which accelerates the movement of the proteins in the gel. Whereas the lower ‘separating gel’ is denser and allows a separation of the proteins by their molecular weight, where smaller proteins move faster as compared to larger ones. Page Ruler Plus (Thermo Fisher Scientific Inc, Waltham, MA, USA- PageRuler Plus Prestained Protein Ladder, Catalog No.:26620) was then added onto one lane to be used as a molecular size standard. After this electrophoresis step, a Trans-Blot Turbo Transfer System (BioRad Laboratories Inc.) was used to blot the proteins from the gel onto a Polyvinylidene Difluoride (PVDF) membrane. An additional electric field was set up and placed on the layers of the gel, the membrane and a Whatman paper, for protection reasons. After this step, the membrane was blocked with 2% of dried milk solution diluted in TBST to prevent nonspecific binding of the following antibodies. After blocking, the primary antibody, diluted in dried milk and TBST, was applied onto the membrane and incubated over night at 4°C. The next day, the membrane was washed thoroughly to exclude unbound antibodies. After that the membrane was incubated for one hour at RT with a secondary antibody, which was labeled with horseradish-peroxidase. Next, the plate was washed again for 30 minutes. Pierce ECL Plus Westernblot Substrate (Thermo Fisher Scientific Inc., Waltham, MA, USA, product Number: 32132) was then applied onto the membrane. With help of Amersham Western Blotting System the membrane was detected. To review the successful transfer of the proteins from the gel to the membrane, the membrane was stained with Ponceau S.
5.9 RNA isolation
To extract RNA, a PureLink® RNA Mini Kit (Ambion, Austin, TX, USA- Catalog No.
12183018A, Lot. No. 1677413) was used. Frozen cell pellets were transferred to an RNAse- free tubes and lysis buffer with 2-mercaptoethanol was added. Tubes were vortexed until the cell pellet was completely dispersed. Lysates were then homogenized using a rotor-stator homogenizer. The homogenates were centrifuged for centrifuged at 1500 rpm for 5 minutes and supernatants were transferred to a clean RNase- free tube. 70% ethanol was added in order to provide binding of the RNA to the membrane of the filter cartridge. The tubes were thoroughly
vortexed. 700 µl of each sample was transferred to a Spin Cartridge. Samples were centrifuged at 12000 rpm for 15 seconds at RT. All supernatants were collected and 700 µl of Wash Buffer I was added. Samples were repeatedly centrifuged at 12000 rpm for 15 seconds at RT for RNA processing and the flow-through was discarded after each centrifugation. 500 µl of Wash Buffer II was added each time. Samples were then centrifuged for 1-2 minutes to dry the membrane with bound RNA, therefore spin cartridge was added into a recovery tube. 20 µl of RNA water was added onto the center of the filter and samples were incubated at RT for 1 minute. In a last step, all samples were centrifuged for 2 minutes at RT to elute the RNA from membrane into the recovery tube. RNA samples were stored at -80 °C until further processing.
5.10 RNA concentration measurement
NanoDrop 2000 Spectrophotometer (Thermo Fisher Scientific Inc, Waltham, MA, USA) was used to determine the total RNA concentration. Calibration with 1.5 µl of RNase-free water and light with a wavelength of 340 nm. For each measurement, 1.5 µl RNA sample was loaded and RNA-absorption was measured at an ultraviolet light with a wavelength of 260 nm. An extinction coefficient of 40 ng-cm/μl was chosen for all measurements. RNA concentration was then calculated by use of the Beer-Lambert-Law. RNA samples were stored on ice at all times.
5.11 Complementary DNA synthesis by reverse transcription
Total RNA samples were first diluted with RNase-free water to be transcribed into cDNA with help of Thermo Scientific RevertAid Reverse Transcriptase (Thermo Fisher Scientific Inc., Waltham, MA, USA). For each sample 13.25 µl of diluted RNA, 11.75 µl of cDNA synthesis master mix containing 5 µl 5x reaction buffer, 2 µl deoxyribose containing nucleotide triphosphates (dNTPs), 2.5 µl random hexamer primers, 0.625 µl RiboLock RNase inhibitor and 0,675 μl RNase free water, was used. After that 1 µl of Reverse Transcriptase was added. One negative was prepared using 11.75 µl of cDNA synthesis master mix and 13.25 µl of the highest diluted RNA sample, without Reverse Transcriptase. All samples were then centrifuged and reverse transcription was started using a peqSTAR & Primus advanced thermocycler (VWR International GmbH, Erlangen, Germany). In order to denature the RNA-strands, all samples were heated up to 65°C for 5 minutes, which allows primer binding to single-strands. In order to initiate the elongation of the new cDNA strands, samples were heated up to a temperature of
42°C for 1 hour. In a last step, the samples were subjected to 90°C for 10 minutes. Each sample included a total amount of 25 µl, all samples were stored at -20°C.
5.12 Real-time quantitative polymerase chain reaction
Quantitative Real-time polymerase chain reaction (qRT-PCR) is a common method used to detect and measure gene expression. The method is based on the DNA-replication of a DNA-molecule in repetitive cycles. Each cycle of PCR starts with denaturation of the double stranded DNA into single stranded DNA (ssDNA) at a temperature of 95°C. This is followed by the binding of DNA-primers to the ssDNA, ultimately leading to elongation with help of the thermoresistant polymerase enzyme. The elongation proceeds in 5’-3’ direction, complementary to the template.
At every cycle, the amount of DNA is doubled thus reaching an exponential increase. In order to perform a quantitative detection of the double-stranded complimentary DNA, SYBR Green I (Thermo Fisher Scientific, Waltham, MA, USA) was used. SYBR Green dye binds to the amplified cDNA and forms a complex, which then proportionally increases the intensity of the fluorescence. The exact amount of the targeted gene can then be quantified based on an amplification plot [82, 83]. All cDNA samples and primers were diluted 1:10 with RNAse-free water to supply 10 ng cDNA per well. 18S rRNA, the small unit of ribosomes, is constitutively and ubiquitarily expressed and therefore serves as a housekeeping gene that can be used for normalization to the expression level of the target gene. 8 µl of qPCR-mix, containing 5 µl of Maxima SYBR Green/ROX qPCR Master Mix (2X) (including Taq-polymerase, dUTPs, qPCR buffer and passive reference dye (ROX)), 0,25 µl of a forward and a reverse primer and 2.5 µl RNase-free water were pipetted into each well of a 96-well-plate, each sample in duplicates.
After that, 2 µl of template cDNA, containing 20 ng cDNA, were added to the qPCR-mix. Two negative controls were prepared for each primer, one containing 2 µl RNase-free water and 8 µl of qPCR-Mix, the other one containing 2 µl reverse transcriptase minus controls and 8 µl qPCR- mix. After the centrifugation of the 96-well plate, the gene expression measurement was performed using a StepOnePlus™ Real-Time PCR System (Thermo Fisher Scientific Inc., Waltham, MA, USA). Detection and evaluation of the samples was performed using StepOneTM Software (Version 2.3). In order to prove the specificity of all primers, a melting curve analysis was included. NLRP3 primer was obtained from Sigma Aldrich (Sigma-Aldrich Chemie GmBH,
Steinheim, Germany), forward primer sequence: CCTTGGACCAGGTTCAGTGT. Reverse primer sequence: AGAAGAGACCACGGCAGAAG.
5.13 Cell viability assay
The water-soluble Tetrazolium (WST-1) Cell Proliferation assay (Sigma Aldrich, St. Louis, MT, USA) is a colorimetric assay for the nonradioactive quantification of cell viability. The WST-1 reagent contains WST-1, an electron-coupling-tetrazoliumsalt (4-[3-(4-Iodophenyl)-2-(4-nitro- phenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate). Once added to the medium of living cells, these tetrazolium salts are cleaved to formazan by mitochondrial dehydrogenases. The amount of formazan directly correlates with the number of metabolically active cells in the culture.
Quantification of the formazan dye produced by metabolically active cells can be done by a spectrophotometer. For the experiments, a stable cell number of 1*10^4 cells and 2*10^4 cells per well in 100 µl medium were seeded in a 96-well plate, the day before the experiment. The next morning, a complete medium change was performed, thus creating stimulation conditions with NaOx in two different concentrations in the medium (0,100 µg/ml NaOx). MCSF was present in medium of all conditions. Cells were then incubated for 12 hours in the incubator.
After the stimulation time, 10 µl of WST-1 reagent was added into each well. The plates were again incubated for a total time of 2 hours. The plate was thoroughly moved for 1 minute on a plate shaker. Absorbance was measured against a background control as a blank, using a microplate reader at 450 nm with a wavelength control at 630 nm. All values are expressed in ratios to control values. Controls were set to 1.
5.14 Gene deficient Slc26a6-/- mice
A knockout (k.o.) mouse is a genetically modified animal with altered genes. Experiments with these mice allow a precise understanding of the effect of the gene that has been knocked out. The gene of interest is therefore isolated from a mouse gene library and a new DNA sequence is engineered into a neighboring sequence [84]. In order to generate Slc26a6-/- mice a vector containing part of exon 2 and 5 and all of exon 3 and 4 of the mouse Slc26a6 gene was electroporated into the 129S6/SvEv embryonic stem cell line TC-1. The clones were then
microinjected into C57Bl/6 blastocysts and the resulting chimeras were bred with 129S6/SvEv mice in order to receive congenic mouse lines [48].
