Investigating Poly(Styrene)-B-Poly(Ethylene Oxide) Polymersomes for the Treatment and Diagnosis of Hyperammonemia

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Doctoral Thesis

Investigating Poly(styrene)-b-Poly(ethylene Oxide) Polymersomes for the Treatment and Diagnosis of Hyperammonemia

Author(s): Matoori, Simon Sam

Publication Date: 2019-03-03

Permanent Link: https://doi.org/10.3929/ethz-b-000328703

Rights / License: In Copyright - Non-Commercial Use Permitted

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ETH Library DISS. ETH Nr. 25671

Investigating poly(styrene)-b-poly(ethylene oxide) polymersomes for the treatment and diagnosis of hyperammonemia

Abhandlung zur Erlangung des Titels

DOKTOR DER WISSENSCHAFTEN der ETH ZÜRICH (Dr. sc. ETH Zürich)

vorgelegt von

Simon Sam Matoori

Master of Science ETH in Pharmaceutical Sciences, Master of Science ETH in Medicinal and Industrial Pharmaceutical Sciences, ETH Zürich

geboren am 30.11.1988

von

Zürich (ZH), Schweiz

Referent: Prof. Dr. Jean-Christophe Leroux Korreferent: Prof. Dr. Peter Walde

Table of contents

Summary ...... 4 Zusammenfassung ...... 6 1. Polymersomes: from Science to science ...... 8 1.1 Polymersome definition ...... 8 1.2 Historical perspective ...... 8 1.3 Preparation methods ...... 9 1.4 Drug delivery ...... 10 1.5 Conclusion ...... 15 1.6 Aims of the Ph.D. thesis ...... 16 2. Recent advances in the treatment of hyperammonemia ...... 18 2.1. Introduction ...... 20 2.2. Reducing ammonia generation and uptake ...... 24 2.3. Increasing ammonia removal ...... 32 2.4. Novel treatment approaches ...... 35 2.5. Conclusion ...... 39 3. Investigating PS-b-PEO polymersomes for oral treatment of hyperammonemia ...... 40 3.1. Introduction ...... 40 3.2. Material and Methods ...... 41 PS-b-PEO polymer synthesis ...... 41 Liposome preparation ...... 42 Polymersome preparation...... 42 Polymer quantification ...... 44 Ammonia uptake in side-by-side cells ...... 44 Ammonia uptake with microparticles ...... 45 Dietary hydrogel experiments ...... 45 Lyophilization ...... 46 In vivo experiments ...... 46 Ammonia assays ...... 47 Statistical analysis ...... 48 3.3. Results and Discussion ...... 48 Screening of microparticle and vesicular formulations ...... 48 Optimization and characterization of PS-b-PEO polymersomes ...... 50 Evaluation of PS-b-PEO library in simulated intestinal fluids ...... 53 Stability in dietary fiber-based hydrogels ...... 54 In vivo evaluation ...... 56 2

3.4. Conclusion ...... 59 4. Development of a PS-b-PEO polymersome assay ...... 60 4.1. Introduction ...... 60 4.2. Materials and Methods ...... 62 HPTS fluorescence excitation spectra ...... 62 PS-b-PEO polymer synthesis ...... 62 Polymersome preparation...... 63 Purification and quantification ...... 63 Ammonia sensing ...... 63 Selectivity ...... 64 Healthy and BDL rat plasma samples ...... 64 Statistical analysis ...... 64 4.3. Results and Discussion ...... 65 pH-dependent fluorescence of HPTS ...... 65 Ammonia sensing with HPTS-containing PS-b-PEO polymersomes ...... 65 Selectivity ...... 68 Ammonia sensing in plasma of healthy rats...... 70 Ammonia sensing in plasma of BDL rats ...... 70 3.4 Conclusion ...... 72 5. General discussion and outlook ...... 73 6. List of abbreviations ...... 78 7. Curriculum Vitae ...... 79 7. Scientific contributions ...... 83 8. Acknowledgments ...... 84 9. References ...... 85

Summary

Ammonia is a ubiquitous metabolite in amino acid and protein metabolism. Despite its important roles in metabolism and pH homeostasis, ammonia needs to be rapidly and efficiently detoxified to avoid an accumulation in the brain where it is associated with neurotoxic effects. The main ammonia removal mechanism in the body is the urea cycle in the liver. In case of inborn or acquired liver disease, the ammonia detoxification capacity of the liver can be insufficient such that ammonia accumulates in the blood (hyperammonemia) and subsequently in the brain. In consequence, patients exhibit low grade to serious neuropsychiatric symptoms. This complication is highly prevalent in patients with liver cirrhosis (hepatic encephalopathy, HE).

As systemic ammonia mainly originates from the bacterial urease activity in the intestine, sequestering gut ammonia is a promising strategy for the chronic treatment of HE. The most widely used treatments for this indication, the laxative lactulose and the antibiotic rifaximin, both target the production of colonic ammonia. However, they fail to provide a sufficient therapeutic benefit to a large number of HE patients and can be associated with unpleasant adverse reactions (e.g., persistent diarrhea). Novel oral treatments with high ammonia extraction capacity are therefore needed.

Furthermore, the current methods to determine ammonia in biologic matrices, namely plasma, have important shortcomings (e.g., interferences, narrow linear range, low-throughput). As ammonia levels correlate with neuropsychiatric symptom severity, are indicative of clinical outcome and the risk of recurrence, and used to assess treatment response, ammonia assays are of great clinical relevance.

In the light of these challenges in the treatment and diagnosis of hyperammonemia, this thesis aims to develop a novel ammonia-lowering treatment and ammonia assay, both based on polymeric vesicles (polymersomes).

In chapter 1, an introduction to polymeric vesicles is provided. Preparation and drug-loading methods of polymersomes are described and compared to liposomal systems. The reported applications of polymersomes in vivo are critically discussed with regard to their potential therapeutic value and their proposed advantages over liposomes. The main objectives of this thesis, the development of polymersome-based systems for the oral treatment and diagnostics of hyperammonemia, are subsequently laid out.

Chapter 2 reviews hyperammonemia-related disorders and hyperammonemia-lowering drugs. Hyperammonemia relates to the accumulation of ammonia in the blood due to the reduced capacity of the liver to metabolize this neurotoxic metabolite. This symptom can be associated with inborn (urea cycle disorders) or acquired (liver cirrhosis, HE) liver disease. Established and investigative treatments reducing ammonia generation and/or increasing ammonia removal are described in this chapter and critically discussed.

Chapter 3 proposes the use of orally applied transmembrane pH-gradient polymersomes to sequester ammonia in the colon and thus lower systemic ammonia levels. We identified polymeric vesicles made of poly(styrene)-b-poly(ethylene oxide) which efficiently captured ammonia in bile salt-containing simulated gastrointestinal fluids. However, a subsequent in vivo study in hyperammonemic rats did not 4 show an effect of the polymersomes on plasma ammonia levels. This finding was explained by the limited stability of the polymersomes in colon-simulating environments.

Chapter 4 proposes how the ammonia concentration in biological fluids can be quantified by pH- sensitive transmembrane pH-gradient polymersomes. HPTS, a dye with pH-dependent and isosbestic fluorescence properties, was encapsulated in the core of poly(styrene)-b-poly(ethylene oxide) polymersomes. Coincubating these polymersomes with ammonia-containing solutions led to changes in the fluorescence spectrum of HPTS, which allowed the quantification of ammonia in a pathophysiologically relevant range. Finally, the ammonia concentration of the plasma of healthy and hyperammonemic rats was determined by the polymersome assay and the most widely used commercial ammonia tests.

Chapter 5 reviews the scientific contributions of this thesis in terms of finding relevant biomedical applications of polymersomes in hyperammonemia-lowering treatments and ammonia diagnostics, and proposes future experiments to elucidate the destabilization of polymersomes in colon-simulating environments and to develop the polymersome-based ammonia assay further.

Zusammenfassung

Ammoniak ist ein essenzieller Metabolit im Aminosäuren- und Proteinmetabolismus. Trotz seiner wichtigen Rolle im Metabolismus und der pH Homöostase muss Ammoniak möglich schnell und effizient detoxifiziert werden, um eine Akkumulierung im Gehirn zu verhindern, wo es mit neurotoxischen Effekten assoziiert wird. Der Hauptmechanismus des Körpers zur Ammoniakentfernung ist der Harnstoffzyklus in der Leber. Im Falle von angeborenen oder akquirierten Lebererkrankungen kann die Ammoniakentfernungskapazität der Leber ungenügend sein, sodass Ammoniak im Blut (Hyperammonämie) und darauffolgend im Gehirn akkumuliert. Als Folge davon zeigen Patienten subklinische bis klinisch offenkundige neuropsychiatrische Symptome. Diese Komplikation ist sehr verbreitet in Patienten mit Leberzirrhose (hepatische Enzephalopathie, HE).

Da das systemische Ammoniak hauptsächlich durch ureaseproduzierende Bakterien im Darm generiert wird, ist die Bindung von Ammoniak in situ eine vielversprechende Strategie für die Behandlung der chronischen HE. Die von den aktuellen Leitlinien empfohlenen Behandlungen für diese Indikation, das Laxans Lactulose und das Antibiotikum Rifaximin, zielen auf die Ammoniakproduktion im Darm ab, entfalten aber bei vielen Patienten nur ungenügende Wirkung und können zu Nebenwirkungen (z. B. Durchfall) führen. Neue orale Behandlungen mit hoher Extraktionskapazität werden deshalb benötigt.

Des Weiteren haben die vorhandenen Methoden zur Bestimmung des Ammoniakgehalts in biologischen Proben, besonders im Plasma, gravierende Nachteile (z. B. Interferenzen, ein beschränkter linearer Bereich, eine tiefe Anzahl messbarer Proben pro Zeiteinheit). Zumal Ammoniakspiegel mit der Schwere der neuropsychiatrischen Symptomatik korrelieren, prognostisch bezüglich klinischen Folgen und der Rückfallwahrscheinlichkeit und zur Evaluation des Ansprechens auf Medikamente benötigt werden, sind Ammoniakmessungen von grosser Bedeutung in der Klinik.

Im Angesicht dieser Herausforderungen bei der Behandlung und Diagnose der Hyperammonämie zielt diese Dissertation auf die Entwicklung neuer ammoniaksenkender Therapien und Diagnostika ab.

Das erste Kapitel bietet eine Einführung in Polymersomen. Herstellungsmethoden und Verfahren zur Beladung der Polymersomen mit therapeutischen Substanzen werden beschrieben und mit Liposomen verglichen. Die in der Literatur beschriebenen Anwendungen von Polymersomen in vivo werden daraufhin mit Blick auf mögliche Vorteile gegenüber Liposomen kritisch diskutiert. Eine potenzielle neue Anwendung der Polymersomen in der Hyperammonämie wird schliesslich vorgeschlagen und die Hauptziele der Dissertation werden dargelegt.

Das zweite Kapitel bietet einen Überblick über hyperammonämieverbundene Erkrankungen und hyperammonämiesenkende Medikamente. Hyperammonämie bezeichnet die Akkumulierung von Ammoniak im Blut aufgrund einer reduzierten Kapazität der Leber, diesen neurotoxischen Metaboliten zu entgiften. Das Symptom kann mit angeborenen (Harnstoffzyklusstörungen) oder akquirierten (Leberzirrhose, HE) Lebererkrankungen assoziiert sein. Etablierte und sich in der Forschung befindende Behandlungen, welche die Ammoniakgenerierung vermindern oder die Ammoniakentfernung erhöhen, werden in diesem Kapitel kritisch besprochen.

Das dritte Kapitel untersucht die perorale Anwendung von Polymersomen mit transmembranärem pH Gradienten zur Ammoniakbindung im Colon vor, womit die Ammoniakspiegel im Plasma erniedrigt werden sollen. In der Literatur wurden Liposomen mit pH Gradienten über der Membran beschrieben, welche Ammoniak sequestrierten und die Ammoniakspiegel im Plasma hyperammonämischer Ratten senkten. In dieser Dissertation wurden PS-b-PEO Polymersomen identifiziert, welche Ammoniak in gallensaltzhaltigen Lösungen, welche den Gastrointestinaltrakt simulieren, effizient aufnahmen. Eine darauffolgende in vivo Studie in hyperammonämischen Ratten zeigte jedoch keine Effekte der Polymersomen auf die Ammoniakplasmaspiegel. Diese Erkenntnis wurde durch die limitierte Stabilität der Polymersomen in colonsimulierenden Umgebungen mit tiefem Wassergehalt erklärt.

Das vierte Kapitel präsentiert eine Methode, mit welcher Ammoniakkonzentrationen in biologischen Flüssigkeiten mit Polymersomen mit transmembranärem pH-Gradienten quantifiziert werden können. Pyranine (8-hydroxypyrene-1,3,6-trisulfonate, HPTS), ein Farbstoff mit pH-abhängigen und pH- unabhängigen Fluoreszenzeigenschaften, wurde in die innere Phase von PS-b-PEO Polymersomen eingekapselt. Werden diese Polymersomen einer ammoniakenthaltenden Lösung beigegeben, ändert sich das Fluoreszenzspektrum des Farbstoffs, sodass eine Quantifizierung des Ammoniakgehalts möglich wird. Mit einer Ammoniakstandardkurve in einer gepufferten Lösung konnte die Ammoniakkonzentration in Plasma, der gebräuchlichsten Matrix für Ammoniakmessungen im Spital, von gesunden und hyperammonämischen Ratten bestimmt werden.

Das fünfte Kapitel fasst die wissenschaftlichen Erkenntnisse der Dissertation, welche die biomedizinische Anwendung von Polymersomen in der Senkung und Quantifizierung von Ammoniakspiegeln betreffen, zusammen. Im Übrigen werden zukünftige Experimente vorgeschlagen, um die Destabilisierung von Polymersomen in wasserarmen Milieus zu untersuchen und um die polymersomenbasierte Ammoniakmessmethode weiterzuentwickeln.

Chapter 1

1. Polymersomes: from Science to science

1.1 Polymersome definition

Polymersomes (polymerosomes, polymeric vesicles) are defined as vesicular macromolecular assemblies whose bilayer membrane is composed of amphiphilic polymers (block, dendronized, graft, or alkylated copolymers) [1–4]. The hydrophilic coronas of the membrane face the aqueous core and outer aqueous medium. The hydrophobic layer of the membrane separates the inner from the outer medium. The molecular composition and length of the hydrophobic and hydrophilic blocks determine different polymersome properties such as membrane rigidity, size, and stability [5,6].

1.2 Historical perspective

In 1995, a seminal paper in Science by Zhang and Eisenberg advanced our understanding of polymeric macromolecular assemblies [1]. The study described the morphological diversity of macromolecular assemblies of poly(styrene)-b-poly(acrylic acid) (PS-b-PAA) diblock copolymers in a N,N- dimethylformamide-water mixture. Upon decreasing the PAA block length, transmission electron microscopy images revealed a transition from spherical (PS200-b-PAA21, 26 nm in diameter) to rod-like micelles (PS200-b-PAA15, 23 nm), to vesicles (PS200-b-PAA8, 100 nm), and large spherical aggregates

(PS200-b-PAA4, up to 1.2 µm). This study revealed that amphiphilic block copolymers were able to form vesicular structures in low molecular weight solvents, and that tuning the hydrophilic to hydrophobic ratio of block copolymers allowed the formation of different macromolecular assemblies. A subsequent study revealed that the macromolecular morphology of a diblock copolymer can be altered by modifying the ionic strength of the medium [7]. Moreover, a study by van Hest et al. in 1995 described different macromolecular assemblies formed by amphiphilic polymers composed of a PS block and a poly(propylene imine) dendrimer. While PS-dendr-(NH2)8 formed vesicles with diameters below 100 nm,

PS-dendr-(NH2)16 formed micellar rods (~12 nm in diameter) and PS-dendr-(NH2)32 spherical micelles (~10-20 nm) in aqueous solutions [2].

The notion that polymersome membranes are much tougher than liposomal ones was first put forward by Discher et al. who showed that poly(butadiene) (PBD)-b-poly(ethylene oxide) (PEO, also referred to as poly(ethylene glycol), PEG) polymersomes resisted higher areal strains than liposomes made of the unsaturated lipid 1-stearoyl-2-oleoyl-sn-glycero-3-phosphocholine (SOPC) in micropipette aspiration experiments [8]. While the investigated SOPC liposomes ruptured at an areal expansion of 5%, poly(ethyl ethylene) (PEE)-b-PEO polymersomes ruptured at ~10%, and PBD-b-PEO at 10-30% areal strain [5]. Interestingly, the hydrogenation of PBD to PEE therefore did not increase the stability even though a better membrane packing in the absence of double bonds would be expected [5]. Furthermore,

PBD250-b-PEO150 polymersomes (membrane thickness ~21 nm) ruptured at lower areal strength than

PBD125-b-PEO80 polymersomes (~15 nm), showing that neither the hydrophobic block length nor the membrane thickness are the only determining factors of the membrane strength [5]. Furthermore, the elastic moduli of several polymersomes and liposomes were in a similar range and strongly depended on block copolymer and lipid composition, respectively [5,9–16]. Albeit observed only for a limited

8 number of liposome and polymersome systems, the higher resistance to rupture of polymersomes led to the wide propagation of the notion that polymersomes are much more stable than liposomes [17–22]. However, an in-depth comparison of different polymersomes (e.g., biodegradable vs non-biodegradable polymers) and liposomes of different composition (e.g., saturated vs. unsaturated lipids, cholesterol content) would be needed to determine the generalizability of this finding. The question, which clinical applications necessitate tougher membranes than liposomal ones, also needs to be addressed, especially with regard to the considerable regulatory hurdles in the introduction of novel delivery systems.

1.3 Preparation methods

Polymersomes can be prepared by various methods. In general, the polymer is dissolved in an organic solvent to form an organic phase, and mixed with an aqueous solution (aqueous phase). The mixing allows a fine dispersion of the polymer in the water phase and the subsequent formation of polymersomes. Different techniques may be employed for the mixing step: nanoprecipitation, emulsification, or film rehydration.

1.3.1 Nanoprecipitation

Nanoprecipitation is a widely used method for polymersome production [6,23,24]. The polymers are dissolved in a suitable water-miscible organic solvent, to which the aqueous phase is slowly added under stirring, usually with a syringe pump. The complete removal of organic solvent in the solution is generally difficult to achieve due to its miscibility with water. The resultant solvent residues may limit the usefulness of this method in the production of formulations for (pre)clinical use due to potential toxicity. Furthermore, the solvent has been reported to act as a plasticizer in the membrane, potentially affecting membrane properties such as membrane rigidity [25].

1.3.2 Emulsification

1.3.2.1 Single emulsion

In the o/w emulsion method, the polymer-containing, at least partly water-immiscible organic solvent is mixed with the aqueous phase under sonication, homogenization, or vigorous stirring [26]. This method is promising for the preparation of formulations for in vivo use because the organic solvent can be removed by evaporation or filtration methods to a very high degree (see below). In addition, emulsification-based methods can be scaled up relatively easily and may be performed in continuous processes [27,28].

1.3.2.2 Double emulsion

In the double-emulsion method, polymersomes form in a w/o/w double emulsion containing an aqueous inner phase, a polymer-containing, at least partly water-immiscible organic solvent in the middle phase, and an aqueous outer phase [29–31]. This method can be elegantly carried out with a microfluidic system with which narrow size distributions in the low micrometer size range can be achieved [29–31]. The scale up with microfluidic systems remains challenging, though [32].

1.3.3 Film rehydration 9

In the film rehydration method, the polymer is dissolved in an organic solvent and subsequently dried, allowing the formation of a thin polymer film [33–35]. Upon addition of the aqueous phase and thorough mixing, the polymersomes spontaneously form. While this method can yield highly concentrated vesicular dispersions with low residual solvent amounts for liposomes [36], it may not be applicable to highly hydrophobic block copolymers with high glass transition temperature such as PS-b-PEO (see below). Potentially, PS forms highly ordered and rigid structures upon drying which hinders the rehydration procedure.

1.3.3 Size control

The vesicle size affects the uptake by the mononuclear phagocytic system, which influences circulation time and organ distribution, and extravasation [37]. Membrane extrusion and sonication can be used to control the polymersome size [25,38–42]. Similarly to liposomes, the size of polymersomes composed of polymers with low glass transition temperature hydrophobic fragments (e.g., poly(propylene oxide), PBD) can readily be decreased applying freeze-thaw and/or extrusion cycles, yielding polymersomes of less than 100 nm in diameter [38–42]. However, for polymers with a high glass transition such as PS-b- PEO, decreasing the size of polymersomes with filtration and sonication has only been reported in the presence of considerable volume fractions of solvent [25]. The solvent probably acts as a plasticizer of the semi-crystalline membrane, facilitating macromolecular rearrangements upon filtration or sonication.

1.4 Drug delivery

1.4.1. Drug loading

Polymersomes and liposomes offer the possibility to host hydrophobic and hydrophilic drugs.

Hydrophobic drugs

Hydrophobic molecules partition into the hydrophobic layer of the vesicle membrane provided that their affinity for this environment is sufficiently high [43]. While liposomal membranes are typically ~3-5 nm thick, polymersomes have membrane thicknesses of up to ~50 nm [44,45]. Therefore, polymersomes could in theory accommodate larger and higher amounts of hydrophobic molecules than liposomes.

Using nanoprecipitation at pH 10.5, doxorubicin was loaded into poly(trimethylene carbonate)-b-poly(L- glutamic acid) polymersomes with an encapsulation efficiency of 78% and a drug loading of 47 wt% [46]. Transmission electron microscopy indicated that the high drug loading was related to the formation of doxorubicin nanoparticles at the alkaline pH (pKa 8.3), which were incorporated in the hydrophobic layer of the membrane [46]. Hydrophobically modified iron oxide nanoparticles with a radius of ~3-5 nm were also incorporated into the membrane of poly(trimethylene carbonate)-b-poly(L-glutamic acid) vesicles [23]. Interestingly, ~5 nm-thick liposomal membranes made of the saturated phospholipid 1,2- dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) can also accommodate hydrophobic iron oxide nanoparticles of 5 nm in radius [47]. Therefore, the membrane thickness does not seem to be the only determining factor in the loading of large hydrophobic substrates such that liposomal controls could be relevant even in the case of larger hydrophobic substrates.

When loading substrates into the membrane, the risk of low drug retention due to the low diffusion distance in the membrane needs to be considered. Even though polymersome membranes are generally thicker than liposomal ones, the diffusion distance remains small, resulting in a risk of rapid cargo release. Indeed, paclitaxel-loaded hydrophobic nanoparticles with a diameter of ~180 nm showed a fast release of their hydrophobic cargo due to diffusion [48]. To determine the degree of retention in the membrane, thorough studies on the release kinetics of membrane-loaded substrates are needed in biorelevant media and under sink conditions or, preferably, in vivo. Moreover, the use of biodegradable polymers (e.g., for systemic administration) could further impact the release profile in vivo, as the degradation of the polymer could destabilize the membrane and result in accelerated cargo release.

Hydrophilic drugs

Hydrophilic substrates can be added to the aqueous phase in which the vesicles form or, if they can diffuse across the membrane, to the outer phase for active loading methods. With the latter processes, notably transmembrane pH-gradients, much higher encapsulation efficiencies can be achieved. In this approach, the uncharged species of weakly basic or acidic drugs diffuses across the membrane into the vesicular core. The core bears a low (basic drugs) or high pH (acidic drugs) such that the substrate becomes charged and subsequently trapped in the core as the charge hinders the diffusion across the hydrophobic part of the membrane [49].

Leaky membranes, however, impair the stability of the transmembrane pH-gradient and may result in low loading efficiencies and drug leakage after loading. The thicker membranes of polymersomes could therefore be an advantage over the liposomal ones for this remote loading procedure provided that they remain permeable enough for a sufficiently rapid diffusion of the substrate. The doxorubicin encapsulation efficiency in poly(2,4,6-trimethoxybenzylidenepentaerythritol carbonate) (PTMBPEC)-b- PEO polymersomes was ~25% [50], for mixed polymersomes composed of PBD-b-PEO (75 wt%) blended with poly(L-lactic acid)-b-PEO (25 wt%) ~60% [51] compared with almost 100% for liposomes made of saturated phospholipids [52,53]. The drug loading in terms of doxorubicin weight per weight of polymer or lipid was ~8 wt% for PTMBPEC-b-PEO polymersomes [50], ~5 wt% for polymersomes made of PBD-b-PEO blended with poly(L-lactic acid)-b-PEO [51], ~5 wt% for PEGylated liposomes made of the saturated phospholipid 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) [54], and ~13 wt% for the Food and Drug Administration (FDA)-approved liposomal doxorubicin formulation Doxil© made of fully hydrogenated soy phosphatidylcholine (55 mol%), cholesterol (40 mol%), and mPEO(2000)-1,2- distearoyl-sn-glycero-3-phosphoethanolamine (5 mol%) [55–58]. With regard to doxorubicin leakage under storage conditions, polymersomes made of PBD-b-PEO blended with poly(L-lactic acid)-b-PEO showed a release of ~5% at 4°C after one month [51]. The shelf-life of Doxil© of twenty months at 2-8°C underlines that drug retention in liposomal systems can also be very high [59]. Over thirty years of experience with the transmembrane pH-gradient loading method in liposomes yielded a range of investigative and FDA-approved liposome formulations with high encapsulations efficiencies, drug loadings, and very low drug leakage rates [52,60]. Therefore, optimized liposomal formulations are highly suitable for the transmembrane pH-gradient-based active loading method. Clear advantages of the polymersome formulations reported to date are not obvious for this procedure in terms of encapsulation efficiency, drug loading, and drug leakage. 11

Conjugating small and large molecules to the polymersome-forming block copolymer or to its hydrophobic fragment can be used to encapsulate drugs and decorate polymersomes with targeting ligands [61–63]. Encapsulating drugs with this method (analogously to drug-lipid conjugation for liposomes [64,65]) may provide prolonged release kinetics due to an improved retention in the vesicle [66]. Biodegradable linkers may be employed to release the molecule from the polymer [18]. If a non- biodegradable linker is used, potential interferences of the polymer(some) with the binding of the drug to its target need to be investigated.

1.4.2 In vivo studies

1.4.2.1 Systemic administration

Conceptually, one of the main advantages of polymersomes over liposomes for systemic drug delivery applications is their higher PEO density. Surface-exposed PEO decreased the binding of opsonins to liposomes, reduced their uptake by opsonin-recognizing cells of the mononuclear phagocyte system (MPS), and prolonged circulation time compared with non-PEGylated liposomes [67]. The PEO density on PEO-containing polymersomes is much higher than on liposomes, which generally contain a maximum of 5 mol% PEO-conjugated lipids [68]. Therefore, PEO-containing polymersomes may in theory exhibit a longer circulation time than liposomes, thereby altering the pharmacokinetic profile of the encapsulated drug more than lipid vesicles. However, a prolonged plasma half-life of the drug is only achieved if it is sufficiently slowly released from the delivery system [48]. Furthermore, a higher PEO density can also confer unwanted properties. PEGylated liposomes elicit an immune response with anti- PEO immunoglobulin M (IgM) antibodies. These antibodies led to an accelerated clearance upon repeated injection of PEGylated liposomes and nanoparticles [69,70]. Even though the effect of PEO density on this immune response is not fully understood, polymersomes with a PEO corona could also induce an anti-PEO IgM-associated immune response with a bearing on the circulation time upon repeated administration. Moreover, a PEO layer on the liposome reduced the liposome uptake into cells [71]. When vesicle internalization is required, the higher PEO density of PEO-containing polymersomes may lead to lower therapeutic effects due to reduced cellular uptake.

The number of pharmacokinetic studies on polymersomes is limited. One of the most cited ones investigated the circulation time of four non-biodegradable polymersomes made of PBD-b-PEO or PEE- b-PEO [33]. To assess the circulation time of the polymersomes in vivo, they were stained with the hydrophobic fluorescent dye PKH26 [33]. This dye is conjugated to a C14 and a C22 chain and generally used for cell staining as the aliphatic chains insert into the lipid bilayer [72]. An evaluation of the dye leakage in biorelevant media was unfortunately not performed. The circulation half-lives of the polymersomes (~120 nm) upon intravenous injection in rats were between 16 and 28 h, with longer circulation times observed for PBD54-b-PEO50 and PBD130-b-PEO80 polymersomes (i.e., with longer PEP chains) than for PBD46-b-PEO26 and PEE37-b-PEO40 polymersomes [33]. In another pharmacokinetic study, 111In-labeled PBD-b-PEO polymersomes of 90 nm showed a circulation half-life of ~20 h in mice as determined with single-photon emission computed tomography [42]. Unfortunately, a direct comparison to PEGylated liposomes was not performed in these studies [33,42]. In the literature, PEGylated liposomes of similar size showed half-lives of 15 to 35 h in rodents [69,70,73–76]. Therefore,

12 a clear advantage in terms of circulation time is currently not proven for polymersomes compared to PEGylated liposomes, and potential effects of repeated administrations on circulation time are unknown.

Incorporating doxorubicin in the core and paclitaxel in the membrane of polymersomes (3 mg/kg doxorubicin, 7.5 mg/kg paclitaxel) made of a blend of PBD125-b-PEO80 (75 wt%) and poly(L-lactic acid)56- b-PEO109 block copolymers (25 wt%) led to prolonged tumor doxorubicin exposure and accelerated tumor regression in tumor-bearing nude mice compared with the free drugs (1.5 mg/kg doxorubicin, 1.0 mg/kg paclitaxel) [77]. While paclitaxel levels were neither assessed in the tumor nor in plasma, doxorubicin tumors levels of the polymersome formulation peaked after one day and decreased to ~40% and ~0% after two and four days, respectively (Figure 1.1A) [77]. Unfortunately, liposomal controls were not included in this study. In our opinion, the liposomal doxorubicin formulation Doxil© would be an appropriate positive control due to its high stability (saturated phospholipids, high cholesterol content) and stealth properties (PEGylation) [56–58]. In a study on Doxil© in tumor-bearing mice, the liposomal formulation resulted in a much greater prolonged exposure of the tumor to doxorubicin compared with the free drug (Figure 1.1B) [78]. The maximum concentration was achieved after two days, remained stable until day five, and was decreased to ~70% on day seven after injection [78]. The faster doxorubicin tumor clearance of the polymersome formulation could be due to a shorter plasma half-life or a higher leakage of doxorubicin before reaching the tumor. Unfortunately, the plasma half-life of the polymersomes was not calculated in the study [77]. Comparing the half-life of similar polymersomes

© (PBD130-b-PEO80, ~26 h [33]) with the one of Doxil (~35 h [73], both in rats), strongly different plasma clearance profiles in mice would not be expected. Furthermore, a comparison of the absolute tumor accumulation of the doxorubicin-loaded vesicles cannot be made because the tumor doxorubicin levels were not quantified in terms of amount of drug per amount of tumor tissue in the polymersome study [77]. This example, in which a liposomal formulation led to a longer exposure of the tumor to the chemotherapeutic agent, underlines the importance of liposomal controls in the evaluation of novel polymersome formulations.

Figure 1.1. Comparison of the normalized tumor doxorubicin deposition in mice receiving polymersome or liposome formulations. Polymersome encapsulation led to higher doxorubicin levels compared with the free drug at 24 h (A), while liposomal doxorubicin (Doxil©) led to prolonged tumor exposure to doxorubicin for at least seven days (B). Plot A: free and polymersome-encapsulated doxorubicin administered in nude mice bearing MDA-MB231 human Caucasian breast adenocarcinoma tumors of

0.5 cm2, dose not mentioned, n = 2 per time point [77]. Plot B: free doxorubicin and Doxil at 9 mg/kg administered ten days after 4T1 mouse mammary carcinoma tumor inoculation in female BALB/c mice (tumor size not indicated), n = 3-5 per time point [78]. Data normalized in both plots to the highest mean tumor doxorubicin level in the respective free doxorubicin group. DOX, doxorubicin.

In another study on doxorubicin delivery, poly(γ-benzyl L-glutamate)-b-hyaluronic acid polymersomes loaded with doxorubicin were taken up by CD44 (a hyaluronic acid-binding glycoprotein)-expressing breast cancer cells in vitro [79]. These polymersomes decreased the tumor size of breast cancer-bearing rats in vivo more than free doxorubicin [79]. The encapsulation of doxorubicin into polymersomes further decreased its cardiotoxicity [79] which was reported for liposomal doxorubicin in 1984 [80]. This example shows that the versatility of the polymersome platform in terms of polymer composition can be elegantly exploited as high levels of surface exposed hyaluronic acid are conceptually very useful to improve the uptake by CD44.

Furthermore, oligo(porphyrin) near-infrared dyes were incorporated in the membrane of PBD PEO polymersomes [44]. These highly rigid and hydrophobic dyes were incorporated into the polymer film (film rehydration method) and encapsulated in the membrane during rehydration [44]. While the polymersomes could incorporate up to pentameric oligo(porphyrin), liposomes made of SOPC could only accommodate dimers in their membrane [44]. The authors speculated that the trimeric to pentameric oligo(porphyrin) were too large (~3-5 nm) for the liposome membrane (~3.5 nm) compared to the polymersome (~10 nm) [44]. However, other factors could have influenced the loading procedure as the membrane of DPPC liposomes was capable of incorporating hydrophobic nanoparticles of ~5 nm [47]. Upon intratumoral injection of the oligo(porphyrin)-loaded polymersomes into subcutaneous glioma-bearing rats (tissue depth 1 cm), a strong fluorescence signal was observed in the investigated 20 min post-injection interval [44]. In view of the missing positive and negative controls in the in vivo study, the added value of a polymersome-based delivery system over other platforms cannot be judged.

In conclusion, convincing evidence that polymersomes circulate longer and transport increased amounts of hydrophilic or hydrophobic cargo to the target tissue than liposomes is still lacking for most systems and applications. Head-to-head comparisons of polymersomes with well-performing liposome formulations would be needed to determine if the cumbersome development of alternative drug delivery systems to liposomes is sensible. The liposomal composition used in Doxil© would be a suitable comparator due to its high toughness (high cholesterol content, saturated lipids) and stealth properties (PEGylation) [57]. Moreover, alternative platforms such as micelles or nanoparticles may be considered as suitable controls in the delivery of hydrophobic cargo [48].

1.4.2.2 Oral administration

The gastrointestinal (GI) tract is a harsh environment for vesicles [81]. High bile salts concentrations, strong osmolarity and pH changes, and high enzymatic activity generally lead to rapid destabilization of vesicular structures [81–85]. Bile salts impaired the membrane integrity of liposomes by insertion into the outer leaflet of the membrane and by the partitioning of the lipids into bile salt micelles, which may lead to cargo release [85–90]. Hypo- and hyperosmolar environments, which can range from 100 to 600 mOsmol/kg in the GI tract [91–93], induce morphologic changes (i.e., shrinking and swelling, 14 respectively) of vesicles [94,95], potentially impairing structural integrity of the membrane which could lead to the loss of the cargo. Furthermore, polyester-based polymersomes, surface-exposed peptide- based targeting ligands, and phospholipid-based liposomes may be hydrolyzed at the low pH of the stomach or due to enzymatic action (e.g., pepsin, (chymo)trypsin, phospholipase A2 for phospholipids) [82,96,97]. In the light of these challenges, reports on the use of polymersomes in oral drug delivery are scarce.

To enhance the solubility of the poorly soluble multikinase inhibitor sorafenib, it was encapsulated in the hydrophobic part of PBD-b-PEO polymersomes and orally administered to healthy mice [98]. The area under the plasma concentration vs. time curve of the polymersome formulation was higher compared to the control formulation, a sorafenib suspension [98]. With regard to the instability of PBD-b-PEO polymersomes in surfactant-containing media [99], we would expect highly variable release profiles due to inter- and intraindividual differences in bile salt secretion, notably with regard to food consumption.

A study reported that orally applied insulin-containing dextran-b-poly(D,L-lactide-co-glycolide) polymersomes led to appreciable systemic insulin levels and decreased glucose levels in diabetic mice [100]. In view of the difficulty of reaching the systemic circulation with small peptides (let alone macromolecular assemblies such as vesicles) [101], a more thorough investigation of the translocation from the gut lumen into the systemic compartment is needed to fully understand these results. Furthermore, a more thorough characterization of the polymersome system (e.g., resistance against amphiphilic structures such as bile salts, extreme osmolality) would be desirable to exclude artefactual results.

A potential approach to improve the resistance of polymersomes to the harsh conditions of the GI tract would be the cross-linking of the polymersome membrane. Cross-linking procedures have been reported for PBD-b-PEO, PEO-b-PAA-b-poly(N-isopropylacrylamide), and PEO-b-poly(diethylaminoethyl methacrylate)-b-poly(3,4-dimethyl maleic imido butyl methacrylate) polymersomes [102–104]. However, the removal of toxic residues from cross-linking procedure could be challenging as they may be retained in the membrane. Therefore, there could be a risk of releasing these toxic compounds once the crosslinked polymersomes are in the body. If the drug is already loaded, performing a cross-linking procedure could be risky with regard to potential chemical interactions of the cross-linking reagents and the drug.

1.5 Conclusion

The observation in 1998 that PBD-b-PEO polymersomes have stronger membranes than SOPC liposomes raised high hopes for the development of novel vesicular drug delivery systems that are superior to liposomes in terms of stability, circulation time, and drug loading capacity. After 20 years, there are still open questions about the generalizability of the higher membrane strength beyond the tested formulations. In addition, the very few studies on circulation time pointed to similar half-lives as observed for PEGylated liposomes. Furthermore, a clear superiority in the loading capacity of hydrophilic or hydrophobic drugs has not been established for polymersomes, and the impact of membrane biodegradability on plasma half-life, drug release, and polymersome stability in general remains poorly investigated. Moreover, finding upscalable polymersome preparation procedures leading to very high 15 polymersome concentrations and low amounts of residual solvent remains challenging. In our opinion, more systematic studies on these issues comparing polymersomes of various composition against strong liposome controls are needed to investigate the initially assumed superiority of polymersomes. Only when the advantages of polymersomes are clearly established, will it be possible to find clinical applications where the efforts of developing an alternative vesicular drug delivery platform to liposomes, whose safety profile is well established in the clinics, is warranted.

1.6 Aims of the Ph.D. thesis

The aim of this Ph.D. thesis is to identify relevant biomedical applications for polymersomes in which a vesicular structure is needed and the performance of liposomes is insufficient. We propose to use polymersomes in the treatment and diagnosis of hyperammonemia. In hyperammonemia, the neurotoxic metabolite ammonia accumulates in blood due to insufficient hepatic clearance in patients with liver disease. This disorder and its current treatments are thoroughly discussed in chapter 2.

Building on the laboratory’s development of a transmembrane pH-gradient liposome formulation for ammonia sequestration in the peritoneal space [36,105,106], we aim in chapter 3 to develop an oral transmembrane pH-gradient polymersome formulation to sequester ammonia in the colon (Figure 1.2). The polymersome formulation is first optimized in GI-simulating media in vitro and subsequently tested in bile duct-ligated (BDL) rats to investigate its ammonia-lowering capacity in cirrhotic, hyperammonemic rats in vivo.

Figure 1.2. Intestinal ammonia capture with transmembrane pH-gradient polymersomes. Ammonia is produced by urease-producing bacteria in the gut. After diffusing across the hydrophobic membrane, ammonia is protonated and thus trapped in the polymersome. The ammonia-containing polymersomes are subsequently excreted in the feces.

As currently used ammonia assays are susceptible to interferences, cumbersome to carry out, low- throughput, and/or have a narrow linear range, we aim to develop a novel ammonia assay based on transmembrane pH-gradient polymersomes (Figure 1.3). In chapter 4, the pH-sensitive fluorescent dye pyranine (8-hydroxypyrene-1,3,6-trisulfonate, HPTS) is encapsulated in PS-b-PEO polymersomes. The

16 ammonia assay based on these fluorescent polymersomes is first optimized in simple buffers and subsequently tested in the plasma of healthy and hyperammonemic rats.

Figure 1.3. Ammonia sensing using transmembrane pH-gradient polymersomes. The influx of ammonia leads to an increase in the core pH, which mainly depends on the extravesicular ammonia concentration. If a pH sensitive dye is encapsulated in the core, the increase in core pH can be evaluated based on the alterations in the spectral properties of the dye. Thus, a relationship between the spectral changes of the encapsulated dye and the ammonia concentration in the solution can be established.

Chapter 2

2. Recent advances in the treatment of hyperammonemia

This chapter is an updated version of the following publication:

Simon Matoori1, Jean-Christophe Leroux1

1ETH Zurich, Department of Chemistry and Applied Biosciences, Institute of Pharmaceutical Sciences, Vladimir-Prelog-Weg 3, 8093 Zurich, Switzerland.

Advanced Drug Delivery Reviews 2015; 90:55-68. doi: 10.1016/j.addr.2015.04.009.

Abstract

Ammonia is a neurotoxic agent that is primarily generated in the intestine and detoxified in the liver. Toxic increases in systemic ammonia levels predominantly result from an inherited or acquired impairment in hepatic detoxification and lead to potentially life-threatening neuropsychiatric symptoms. Inborn deficiencies in ammonia detoxification mainly affect the urea cycle (UC), an endogenous metabolic removal system in the liver. Hepatic encephalopathy, on the other hand, is a hyperammonemia-related complication secondary to acquired liver function impairment. A range of therapeutic options is available to target either ammonia generation and absorption or ammonia removal. Therapies for hepatic encephalopathy decrease intestinal ammonia production and uptake. Treatments for UC disorders (UCD) eliminate ammoniagenic amino acids through metabolic transformation, preventing ammonia generation. Therapeutic approaches removing ammonia activate the UC or the second essential endogenous ammonia detoxification system, glutamine synthesis. Recent advances in treating hyperammonemia include using synergistic combination treatments, broadening the indication of orphan drugs, and developing novel approaches to regenerate functional liver tissue. This manuscript reviews the various pharmacological treatments of hyperammonemia and focuses on biopharmaceutical and drug delivery issues.

2.1. Introduction

Ammonia is an important nitrogen source that is involved in amino acid metabolism, protein synthesis, and pH homeostasis[107–109]. At physiological pH, ammonia predominantly exists in its ionized form

+ (ammonium, NH4 , pKa 9.15) with a small gaseous fraction (NH3)[110]. Both the ionized and neutral species cross cellular membranes and the blood brain barrier, via membrane transport proteins and by free diffusion, respectively[111]. Ammonium passes through potassium channels, and it is transported by potassium carriers (e.g., Na+/K+-ATPase, Na+-K+-2Cl– co-transporter isoform 1) because it has a similar ionic radius to potassium[111]. Furthermore, ammonia crosses membranes through aquaporins and human Rhesus glycoproteins A, B, and C[112–114]. An increase in the blood ammonia concentration leads to higher levels of ammonia in the brain[115], eventually resulting in astrocyte swelling, increased blood-brain barrier permeability, altered cerebral metabolism and neurotransmission, and cerebral edema [111,116]. Ammonia also compromises astrocyte potassium buffering[117]. The resultant increase in extracellular potassium concentration and overactivation of the neuronal Na+-K+-2Cl– co-transporter 1 impairs cortical inhibitory neurotransmission and induces neurological dysfunction, which leads to seizures[117].

Most of the systemic ammonia pool is originally generated in the gastrointestinal (GI) tract[118]. The cleavage of urea to ammonia and carbonate by urease-producing bacteria in the colon is a major source of systemic ammonia[118]. The deamidation of glutamine to glutamate by the enzyme phosphate- activated glutaminase in intestinal enterocytes also contributes to intestinal ammoniagenesis, producing one mole of ammonia per mole of glutamine (Figure 2.1)[118]. The glutamine appears to originate rather from the systemic circulation than from the intestinal lumen[118]. After absorption, portal blood ammonia is mainly detoxified by the UC, which is a low-affinity, high-capacity ammonia detoxification pathway in periportal hepatocytes (Figure 2.2)[119]. The remaining ammonia is removed by an enzyme in perivenous hepatocytes, glutamine synthetase, which incorporates an ammonia molecule into glutamate to form glutamine (Figure 2.1)[119]. Skeletal muscles express low levels of glutamine synthetase but upregulate the expression in liver failure, which emphasizes their ammonia-removing and glutamine-producing role in hepatic dysfunction[120,121]. Having both glutaminase and glutamine synthetase activity, the kidney may both increase and decrease blood ammonia[122]. Renal ammoniagenesis due to glutamine breakdown is predominant in patients with liver cirrhosis[123,124]. A protective barrier of the brain against high ammonia levels is the glutamine synthetase activity in astrocytes[123]. However, a limitation of this detoxification system is the osmotic stress which is induced by glutamine accumulation in astrocytes (the so-called “ammonia-glutamine-brain swelling hypothesis”). To reestablish osmotic equilibrium under increasing glutamine production, myo-inostiol and other weak electrolytes are expelled from the cell[123]. Since the brain glutamine levels correlate with arterial ammonia, acute increases in plasma ammonia can generate high glutamine levels in astrocytes which may overwhelm the compensatory mechanisms[123]. Therefore, patients suffering from acute liver failure are more likely to experience glutamine-induced osmotic stress than patients with liver cirrhosis who generally experience more gradual increases in ammonia levels[123].

Figure 2.1. Glutamine synthesis and glutaminolysis. Glutamine synthesis in perivenous hepatocytes and skeletal muscles is an ammonia-detoxifying process that results in the formation of glutamine, a transient metabolic ammonia sink. In contrast, glutaminolysis is an ammoniagenic metabolic pathway that is initiated by renal and intestinal glutaminase. Therapeutic agents with a role in glutamine metabolism (green) either generate glutamate (L-ornithine, BCAAs), the substrate of glutamine synthetase, or prevent glutaminolysis by enhancing glutamine elimination (phenylbutyrate, phenylacetate) or by inhibiting glutaminase. L-ornithine phenylacetate is combination product which increases both glutamine synthesis and elimination.

Figure 2.2. The UC. This cycle is an endogenous detoxification system in periportal hepatocytes. Substrates, intermediates, and products of the UC are used as therapeutic agents (green) in the treatment of certain hyperammonemic conditions to restore the UC’s ammonia-lowering properties.

Hyperammonemia (HA) is defined as plasma ammonia levels of >50 μmol/L in adults or >100 μmol/L in newborns. Toxic increases in ammonia levels result from impaired endogenous hepatic detoxification (e.g., liver cirrhosis, acute liver failure, portosystemic shunting, inborn UCD and UC suppressors such as the antiepileptic drug valproic acid) or increased production of ammonia (e.g., overgrowth of urease- producing bacteria in the intestine or urinary tract)[125]. Factors precipitating acute hyperammonemic crises include GI bleeding, infections and other catabolic state associated with protein degradation, diuretics overuse, and constipation[126,127]. Hyperammonemia is triggered only in the presence of hepatic dysfunction with the exception of the antineoplastic agent L-asparaginase (and its PEGylated version pegaspargase) whose hydrolysis of asparagine and glutamine increases blood ammonia independently of liver function [125–128]. 22

UCDs are rare hereditary disorders with an incidence of approximately 1 in 35,000 births[129], and underdiagnosis is widespread[125]. Primary HA is diagnosed when the resultant HA is related to a dysfunctional UC enzyme or transporter[125]. The inhibition of the UC as a result of metabolite accumulation or substrate deficiencies is referred to as secondary HA[125]. Severe neurological manifestations with the risk of hyperammonemic coma develop soon after birth in newborns with complete deficiencies[125]. The clinical picture of partial deficiencies is dominated by more variable and less severe neuropsychiatric symptoms that may only become evident in adulthood[127]. The reader is kindly referred to the references [125,127] for a detailed discussion of UCDs.

HA is a pivotal factor in the etiology of hepatic encephalopathy (HE), a serious neuropsychiatric and neurocognitive complication in patients suffering from acute liver failure, portosystemic shunting, or liver cirrhosis[126]. The functional inability of the residual liver parenchyma to detoxify ammonia leads to increases in blood ammonia and neurotoxicity[108,126]. Covert HE (“minimal” HE), concomitant with mild increases in ammonia levels and a subclinical to low-grade impairment of cognitive abilities, affects up to 80% of patients suffering from liver cirrhosis[126]. Overt HE is present in up to 21% of cirrhotic patients at the time of diagnosis, and it is associated with confusion, ataxia, and hyperammonemic coma with the risk of progression to irreversible brain damage and death[126]. The reader is referred to the review from Bismuth et al. [130] for a detailed discussion of the pathogenesis of HE and other factors that contribute to HE. Unfortunately, recent advances in the understanding of the pathophysiological mechanisms of HE have not translated into an alleviated disease burden or a substantial improvement in prognosis[126]. In fact, HE continues to lead to considerable and growing costs for health systems with total national expenses increasing from $4.7b in 2005 to $7.2b in 2009 in the United States[131].

HA treatments reduce the generation and uptake of ammonia or improve endogenous detoxification systems (Figure 2.3). The scope of this manuscript is to mainly review approved and investigational pharmaceutical ammonia-lowering treatments with an emphasis on biopharmaceutical aspects and drug formulations. Promising new biomedical therapies including gene and cell therapy as well as novel dialysis technologies will also be covered. Dietary and surgical strategies will not be addressed in this review.

Figure 2.3. Mechanisms of actions of selected ammonia-lowering drugs. In the intestine, the laxative action of non-absorbable disaccharides and poly(ethylene glycol) leads to the expulsion of ammonia (A, B). Non-absorbable disaccharides further acidify the colon, which increases the fraction of the ionized and thus less absorbed ammonium and inhibits the growth of urease-producing bacteria (A). Pro- and antibiotics inhibit bacterial ammonia production by replacing and suppressing urease- producing bacteria, respectively(C). Ammonia production by phosphate-activated glutaminase in enterocytes generates ammonia by degrading systemic glutamine, which can be tackled by glutaminase inhibitors (D). Activated carbon microparticles adsorb ammonium and ammonia in the intestinal lumen, thus preventing ammonia absorption (E). Ammonia scavengers were applied in the peritoneal cavity as well. Transmembrane pH-gradient liposomes ionize ammonia in the acidic core and trap the resultant ammonium (F). In the liver, amino acid and their derivatives (L-ornithine, L- arginine, L-aspartate, BCAAs) support the endogenous ammonia detoxification systems (G), the UC and glutamine synthesis. Phenylacetate is coupled to glutamine and leads to the renal elimination of this transient metabolic ammonia sink (H). Similarly, phenylacetate and benzoate are conjugated to glycine, leading to the excretion of this amino acid and preventing the ammoniagenic degradation by the glycine cleavage system found in the liver, kidney, and brain (I).

2.2. Reducing ammonia generation and uptake

Therapeutic approaches for HE seek to lower the bacterial production and absorption of ammonia in the intestine (non-absorbable disaccharides, antibiotics, probiotics). UCD treatments targeting the metabolic generation of ammonia, on the other hand, reduce the ammonia-producing degradation of glutamine and glycine (Figure 2.1). Therapeutic agents (e.g., sodium benzoate and sodium phenylacetate/phenylbutyrate, glycerol phenylbutyrate) are coupled to these two amino acids, which prevents them from entering ammoniagenic metabolic pathways. The recently identified therapeutic potential of glycerol phenylbutyrate in HE patients interferes with the traditional distinction of treating acquired and inborn HA.

Non-absorbable disaccharides. Lactulose is resistant to digestion in the proximal GI tract, and colonic bacteria convert lactulose to acetic and lactic acid. Lactulose thus acidifies the colonic environment and renders it more hostile to the growth and metabolism of urease-producing bacteria (Figure 2.3) [132– 134]. The decrease in intraluminal colonic pH also shifts the equilibrium between ammonium and ammonia to the considerably less absorbed cation[132]. Finally, the cathartic effect of lactulose, a commonly used laxative, accelerates the excretion of ammonia[135]. The importance of catharsis was recently emphasized in a randomized clinical trial that demonstrated the superior effect of a poly(ethylene glycol) 3350 (PEG(3350))-electrolyte solution, a potent osmotic laxative, over lactulose on clinical and neuropsychological HE symptoms[136]. Notably, lactulose lowered ammonia levels significantly more than the PEG(3350)-electrolyte solution. The dehydrating effect of the osmotic laxative may have decreased the renal elimination of blood ammonia[136]. However, the clinical trial comparing PEG(3350) to lactulose in a larger population of HE patients was withdrawn (NCT01923376). Clinical practice guidelines continue to recommend orally administered non-absorbable disaccharides (lactulose

24 and lactitol) as the first-line drug treatment for the treatment and prevention of overt HE, for which lactulose is approved by the United States Food and Drug Administration (FDA, Table 2.1)[126]. In the initial phase of lactulose therapy, 25 mL of lactulose syrup every 1-2 h are administered until at least two bowel movements per day are induced. The dose is individually titrated thereafter to achieve two to three soft stools per day. The downsides of lactulose treatment include the need to customize the drug dosage, GI adverse effects (abdominal cramping, bloating, and flatulence) and the risk of electrolyte imbalances[126,137]. A meta-analysis has challenged the therapeutic benefit of lactulose despite the widespread use of non-absorbable disaccharides in the clinic[138,139]. This meta-analysis stated that evidence for the beneficial effects of lactulose in HE was absent when low-quality trials were excluded[139]. More recent randomized placebo-controlled trials indicated the usefulness of lactulose therapy for low-grade HE and for the prevention of overt HE[140–142]. However, the therapeutic benefit in the outpatient setting may not be equivalent to that in a clinical trial because of low compliance in lactulose therapy[143]. A meta-analysis of several small studies has demonstrated that the non- absorbable disaccharide lactitol was similarly effective to lactulose, associated with less flatulence, and less sweet[144,145].

Table 2.1 Approved pharmacologic treatments of HA (Drugs@FDA: FDA Approved Drug Products (https://www.accessdata.fda.gov/scripts/cder/daf/), as of November 2018).

International Brand name Formulation Route of Mechanism of Approved Approval Nonproprietary Administration Action Indication Year (FDA) Name Lactulose Generic (e.g., Solution O (R) Intestinal HE (prevention 1988 Enulose) acidification, and treatment) suppression of intestinal bacteria, catharsis Rifaximin Xifaxan Tablet O Suppression of HE (secondary 2010 intestinal prevention) bacteria Neomycin Generic Solution O Suppression of Adjunctive therapy 1964 (e.g., Teva intestinal in hepatic coma Neomycin bacteria (portal-systemic Sulfate) encephalopathy) Sodium Ammonul Solution, to IVcvc Glycine and Acute treatment of 2005 benzoate and be diluted in glutamine UCD sodium 10% dextrose elimination phenylacetate Sodium Buphenyl Tablet, O Glutamine Acute and chronic 1996 phenylbutyrate powder elimination treatment of UCD involving CPS, OTC, and ASS deficiency Sodium Pheburane Granulate O Glutamine UCD involving 2013 phenylbutyrate elimination CPS, OTC, and (European ASS deficiency Medicine (acute and Agency) chronic) Glycerol Ravicti Solution O Glutamine Chronic treatment 2013 phenylbutyrate elimination of UCD for patients >2 years Carglumic acid Carbaglu Dispersible O Restauration of Acute and chronic 2010 tablet NAGS function treatment of NAGS deficiency Arginine Generic Injectable IV Partial Acute N/A solution restauration of management of UC activity UCD ASS argininosuccinic acid synthetase, CPS carbamoylphosphate synthetase, HCl hydrochloride, IVcvc intravenous infusion via central venous catheter, NAGS N-acetylglutamate synthetase, O oral, OTC ornithine transcarbamylase, R rectal, UCD urea cycle deficiency

Antibiotics. Urease-producing bacteria in the large intestine have early been identified as an important source of ammonia production in the body and targeted by antibiotic therapy (Figure 2.3)[146,147]. Enterocytic glutaminase appears to add to intestinal ammonia generation[118], but the predominant role of the intestinal microbiome was recently underlined by two studies linking changes in the microbiome 26 to the risk of HE development and cognitive dysfunction[148,149]. The aminoglycoside neomycin has long been used in the treatment of overt HE episodes[150], an indication for which it is approved by the FDA (Table 2.1), and it is still recommended as a second-line therapy[126,138]. However, its clinical use has waned because of its unfavorable adverse reaction profile, including oto-, neuro-, and nephrotoxicity following systemic exposure[138]. The absorption of orally administered neomycin is merely 3%, but significant systemic exposure and toxic effects have been observed in cirrhotic patients, especially in cases of impaired renal elimination[151]. Similarly, metronidazole, albeit not approved for this indication by the FDA, is recommended as second-line therapy for overt HE[126]. However, a similar adverse event profile as seen for neomycin speaks against continuous long-term use[126]. Since antibiotic action is only needed in the intestinal lumen, an antibiotic with minimal absorption is conceptually preferable. Rifaximin is a semi-synthetic derivative of rifampicin that was originally conceived for topical application[152]. It was designed to have minimal bioavailability while retaining the potential to cross the cell wall of Gram-negative bacteria[153,154]. These properties are associated with rifaximin’s zwitterionic structure, which is related to its pyridoimidazole ring (Figure 2.4A)[153]. Rifaximin is a broad-spectrum antibiotic that acts against diverse Gram-positive and Gram-negative anaerobic and aerobic bacteria[155]. Several clinical trials have found beneficial effects of rifaximin on blood ammonia levels and clinical HE scores[156–160], which resulted in the approval of rifaximin (Xifaxan®, Salix Pharmaceuticals) for the secondary prevention of overt HE in adult patients by the FDA in 2010 (Table 2.1) and a recommendation as an add-on to lactulose in current practice guidelines[126,138]. A recent meta-analysis supported the use of rifaximin in the approved indication and further observed beneficial effects on mortality and full recovery from HE[161]. However, rifaximin monotherapy in HE did not meet cost-effectiveness criteria because of its high cost[162]. Most adverse events of rifaximin affect the GI tract (nausea, bloating, and diarrhea), and serious adverse reactions are rare[161]. However, the question of the risk of rifaximin accumulation was raised for HE patients[163] because the low-level absorbed fraction is usually eliminated by the liver, and markedly increased systemic rifaximin exposure was observed in patients with liver cirrhosis[164]. Interestingly, the administration of generic rifaximin formulations also led to higher blood rifaximin levels in healthy volunteers compared to the originator product[165]. Different crystal polymorphs in the formulations may account for the altered bioavailability as the generics exhibited other polymorphs than the α form, which is the only polymorph in the originator product and associated with the lowest bioavailability among the five crystal polymorphs[166]. Therefore, the risk of higher systemic exposure may increase with the more widespread use of rifaximin generics in the future. With regard to potential systemic rifaximin exposure, the FDA-approved label of rifaximin (as of November 2018) was amended with the advice of exercising caution in patients with highly elevated Model End Stage Liver Disease (MELD) scores (i.e., a score system evaluating liver function in order to assess the need for liver transplantation), as clinical trials were limited to patients with scores below 25. Furthermore, concerns about the risk of developing antibiotic resistance in chronic rifaximin treatment and a possible cross-resistance to other rifamycins, which are used in the treatment of serious infections (e.g. tuberculosis, methicillin-resistant Staphylococcus aureus and Clostridium difficile infections), were voiced[167].

Figure 2.4. Resonance structure and tautomerism of rifaximin and activation of the pro-drugs glycerol phenylbutyrate and phenylbutyrate to phenylacetate. The resonance structure and tautomeric equilibrium of rifaximin (A) result in a zwitterionic character, which is associated with its low bioavailability. Glycerol phenylbutyrate is hydrolyzed by pancreatic lipases to phenylbutyrate (B). After absorption, phenylbutyrate is transformed to the active substance phenylacetate by β-oxidation.

Probiotics. Probiotics are orally administered formulations that contain mono- or mixed cultures of live microbial organisms in a defined number[168]. They are used to promote the growth of non-urease- producing bacteria in cirrhotic patients and reverse the large-intestinal microbial imbalance that is seen in liver cirrhosis[149,169]. Probiotics appear to decrease plasma ammonia levels in HE patients (on average by 7 µmol/L after three months), especially in long-term therapy[170]. However, conclusive evidence of improvements in clinical outcome measures (recovery rate, mortality rate, and hospital stay duration) are lacking[170]. VSL#3 (Sigma-Tau Pharmaceuticals) is a probiotic that is formulated as capsules without enteric coating (Table 2.2)[171,172]. Each hydroxypropyl methylcellulose capsule contains ca. 110 billion live freeze-dried bacteria (Lactobacillus and Bifidobacterium strains, and Streptococcus thermophilus) that are constituents of the normal GI flora of healthy humans[171,172]. A considerable fraction of the bacterial components of VSL#3 seems to remain viable during the passage through the GI tract and reach the colon[172]. VSL#3 was shown to slightly decrease arterial ammonia levels, improve clinical symptoms, and lower the risk of HE episodes compared to placebo in patient populations with low- and high-grade HE in two phase II/III randomized controlled trials[171,173]. Currently, VSL#3 is under investigation in a study with clinical endpoints (phase IV; cognition, risk of falls, and quality of life; NCT01686698). VSL#3 is also being tested in children and young adults with low-grade HE arising from portal vein thrombosis-induced portal hypertension (phase III, NCT01798329). However, the recruitment status of this trial is unknown, as it has not been updated since early 2013. Furthermore, a recent report identified that certain batches of VSL#3 with increased intestinal permeability in rodents, pointing to batch-to-batch variability in the industrial production of this probiotic [174]. Another probiotic consisting of Lactobacillus rhamnosus Gorbach–Goldin, a bacterial strain originally isolated from the GI tract of a healthy human being, was recently found to be safe in 28 patients with low-grade HE in a phase I study, but self-limiting diarrhea was reported[175]. The probiotic exerted beneficial changes in the microbial composition of the intestine, and it had a positive effect on certain inflammatory markers[175], likely because of its ability to adhere to intestinal tissues and its interactions with the host[176]. However, no effects on cognition, venous ammonia levels, or liver disease severity scores were observed with a twice daily regimen[175]. Finally, a trial investigating the effect of co-administering antibiotics on probiotics has not been conducted but may clarify if antibiotics impair the viability of beneficial bacteria.

Table 2.2 Pharmaceutical treatments under investigation for HA (ClinicalTrials.gov registry and Thomson Pharma Integrity database, as of November 2018).

Formulation Route of Mechanism of Action Status Administration

Polyethylene Powder O Osmotic laxative Available; further clinical glycol evidence needed to establish therapeutic benefit in HE VSL#3 Capsule O Probiotic Phase III/IV; further clinical evidence on clinical outcome measures needed L-Ornithine L- Solution O, IV UC and GS activation Authorized in Germany; aspartate phase IV

L-Ornithine Solution IV UC and GS activation; Phase IIB phenylacetate glutamine elimination

Branched-chain Powder, O, IV Multiple mechanisms of Available; further clinical amino acid injectable action evidence needed to solution establish therapeutic Zinc Capsules O benefit in HE

Acetyl-L-carnitine Powder, O, IV injectable solution MARS Albumin- IV Dialysis Phase III dialysis

VS-01 Liposome IP Peritoneal ammonia Preclinical formulation scavenging

AST-120 Suspension O Activated carbon Phase II* adsorbent

Metformin Tablet O Glutaminase inhibitor Available; further clinical evidence needed to establish therapeutic benefit in HE THDP-17 N/A N/A Glutaminase inhibitor Preclinical

ALF-5755 Peptide IV Liver tissue Phase II* solution regeneration

adeno-associated Viral IV Gene therapy Preclinical virus2/8 OTC suspension

Baculovirus-GS Viral IV Gene therapy Preclinical suspension

HepaStem Cellular IV Hepatocyte replacement Phase II suspension therapy GS glutamine synthase, IP intraperitoneal, IV intravenous, O oral, OTC ornithine transcarbamylase, UC urea cycle; *currently no further clinical trials listed under clinicaltrials.gov

Sodium benzoate and sodium phenylacetate/phenylbutyrate. Sodium benzoate and sodium phenylacetate are metabolically coupled to glycine and glutamine, and thus increase the excretion of these two ammoniagenic amino acids (Figure 2.3)[177]. Combination therapy decreases plasma ammonia levels and contributes to the high percentage of survival in acute hyperammonemic UCD patients[177] with an acceptable adverse effect profile (headache, nausea, and impaired mental status)[178]. Benzoate is coupled to coenzyme A (CoA) by benzoyl-CoA ligase to form benzoyl-CoA, which is a substrate for N-acyl CoA:L-glycine N-acyltransferase, in hepatic mitochondria[179,180]. This enzyme transfers the benzoyl moiety of the CoA ester to glycine and yields hippurate, which prevents the degradation of glycine by the glycine cleavage system, an ammonia-forming metabolic pathway situated in the liver, kidney, and the brain[179,181,182]. Similarly, phenylacetate, is activated to phenylacetyl-CoA by phenylacetyl-CoA ligase, and subsequently conjugated to glutamine by the enzyme N-acyl CoA:L-glutamine N-acyltransferase in the liver and the kidney, yielding the renally eliminated phenylacetylglutamine (Figure 2.1)[179,183–185]. While phenylacetate significantly increased the urinary production of phenylacetylglutamine compared to placebo in a porcine model of acute liver failure, a correlation with blood ammonia levels could not be established[186]. Recently, phenylacetate was observed to detoxify ammonia via a novel metabolic pathway[187]. A stoichiometric inverse correlation of plasma ammonia levels and urinary phenylacetylglycine levels pointed to a conjugation of phenylacetate to glycine yielding phenylacetylglycine (Figure 2.1)[187]. Hippurate, phenylacetylglutamine, and phenylacetylglycine are excreted by the kidney and therefore incapable of entering ammoniagenic metabolic pathways which results in the elimination of one (hippurate) and two nitrogen atoms (phenylacetylated conjugates)[181,183]. Ammonul® (Ucyclyd Pharma) is a combination product of 10% sodium benzoate and 10% sodium phenylacetate that is approved by the FDA as an adjunctive therapy for UCD-induced acute HA (Table 2.1). Ammonul® is applied intravenously through a central venous catheter and must be, like sodium benzoate and sodium phenylacetate in monotherapy, diluted in 10% dextrose before infusion to prevent a catabolic state in the patient, which is associated with ammoniagenic protein degradation[127]. An oral liquid of 10% sodium benzoate and 10% sodium phenylacetate marketed under the name Ucephan® (Paco Pharmaceutical Services) was FDA-approved for the chronic management of UCD in 1987, but it was discontinued in 2002[188]. A negative aspect of the oral formulation is the offensive odor of phenylacetate, which was compared to that of a horse stable[189]. Phenylbutyrate is an odorless pro-drug of phenylacetate, which undergoes rapid β-oxidation to phenylacetate in the liver after absorption[179,187]. Sodium phenylbutyrate is formulated as a powder and tablets (Buphenyl®, Hyperion Therapeutics), and it is FDA-approved for adjunctive therapy in the chronic management of several early and late onset UCDs (Table 2.1). One challenging aspect of oral treatment with sodium phenylbutyrate is the salty (sodium) and bitter (phenylbutyrate) taste, which leads to aversive reactions[189,190]. A taste-masked granulate formulation (Pheburane®, Lucane Pharma) was developed by the application of sodium phenylbutyrate onto microgranular sucrose spheres and the addition of a taste-masking outer ethylcellulose layer[190]. This coating layer delayed the dissolution by several minutes, which prevented the interaction of phenylbutyrate with taste receptors[190]. Nevertheless, the formulation was bioequivalent to the comparator compound Buphenyl®[190]. The European Medicines Agency approved Pheburane® in 31

2013 as an adjunctive treatment for the chronic management of UCDs involving deficiencies of carbamoylphosphate synthetase, ornithine transcarbamoylase, or argininosuccinate synthetase (Table 2.1).

Glycerol phenylbutyrate. Glycerol phenylbutyrate (Ravicti®, Hyperion Therapeutics) is an ester pro-drug of phenylbutyrate formulated as a slow-release liquid formulation for oral use. After ingestion, glycerol phenylbutyrate is gradually hydrolyzed by pancreatic lipases (e.g., triglyceride lipase, lipase-related protein 2, carboxyl-ester lipase), which results in delayed release characteristics compared to sodium phenylbutyrate [191]. The maximum approved daily dose of sodium phenylbutyrate exceeds the daily allowance for sodium[191]. Therefore, glycerol phenylbutyrate promises to be a more suitable alternative, especially for patients suffering from hypertension[191]. An added advantage of glycerol phenylbutyrate is its little to absent taste[192]. Currently approved by the FDA for the chronic management of any UCD (with the limitation that safety and efficacy have not been demonstrated in N- acetylglutamate synthetase deficiency) in patients older than 2 years of age (Table 2.1). In a recently published a clinical trial in 17 UCD patients aged between 2 months and 2 years, glycerol phenylbutyrate controlled ammonia and glutamine levels over six months of therapy [193]. Notably, Ravicti costs up to $290,000 per year, roughly three times the amount of Buphenyl[194]. Moreover, glycerol phenylbutyrate is under investigation for HE, where it successfully lowered plasma ammonia and HE events with a similar adverse event profile as the placebo in cirrhotic patients with at least two HE episodes in the six months prior to inclusion in a recent phase II clinical trial[195]. However, no further clinical trials to investigate the therapeutic benefit of glycerol phenylbutyrate in HE are currently listed on clinicaltrials.gov.

2.3. Increasing ammonia removal

Several therapeutic agents support the endogenous ammonia-detoxifying metabolic pathways, notably glutamine synthesis and the UC (Figures 2.1 and 2.2, respectively), lowering systemic ammonia levels once ammonia has been generated or absorbed. Certain treatments have been applied both in UCD and HE (L-arginine, L-carnitine), and others are mostly used in UCD (carglumic acid, L-citrulline) or HE (L-ornithine L-aspartate, L-ornithine phenylacetate, branched chain amino acids, BCAA).

L-ornithine L-aspartate. The salt L-ornithine L-aspartate (Hepa-Merz®, Merz Pharmaceuticals) consists of two substrates of the UC, which fuels this ammonia-detoxifying metabolic pathway in residual hepatocytes (Figure 2.2)[196]. L-ornithine is also transformed to glutamate semialdehyde by ornithine aminotransferase, which is subsequently converted to glutamate (Figure 2.1)[197]. Glutamine synthetase subsequently converts glutamate to glutamine, which detoxifies one ammonia molecule[197]. L-ornithine L-aspartate stimulated muscular glutamine synthetase activity in a rat model of acute liver failure, probably by increasing the concentration of the enzyme’s substrate[198]. Furthermore, oral and intravenous L-ornithine L-aspartate therapy lowers venous ammonia and improves cognitive abilities while being well-tolerated in low-grade and overt HE patients[199–203]. The benficial effect of L-ornithine L-aspartate on the clinical course of HE was recently underlined in a systematic review[204]. Moreover, prophylactic L-ornithine L-aspartate infusion decreased ammonia levels and improved psychometric tests in hyperammonemic patients after transjugular intrahepatic

32 portosystemic shunting[205]. However, a clinical trial in a cohort of patients suffering from acute liver failure did not find any clinical effect of L-ornithine L-aspartate treatment on arterial ammonia, encephalopathy, or survival compared to the current standard of care[206]. This discrepancy may be related to a potentially enhanced efficacy of the standard of care (e.g., preventive administration of antibiotics), which led to a comparatively low mortality in the control group[207]. In general, L-ornithine L-aspartate is applied either as an intravenous infusion (over 4 h, 20-30 g per day) for five to seven days or as a granulate (9-18 g per day split into several doses) supplied in sachets to be prepared as an oral liquid for long-term use[199–203,205,206]. Despite certain encouraging findings, L-ornithine L-aspartate treatment is not widely available and currently not recommended for HE[126,138].

L-ornithine phenylacetate. The concept of combining L-ornithine with phenylacetate in salt form was suggested in 2007 to simultaneously increase the synthesis and excretion of glutamine[120]. As explained above, L-ornithine detoxifies ammonia by activating the UC and glutamine synthetase, while phenylacetate is conjugated to glutamine and glycine to eliminate these ammoniagenic amino acids (Figures 2.1 and 2.2)[186,187,197]. L-ornithine phenylacetate also increased muscular glutamine synthetase expression and activity and hepatic glutamine synthetase expression in a widely used animal model of liver cirrhosis, BDL rats[197]. L-ornithine phenylacetate decreased arterial ammonia levels, extracellular ammonia in the brain, and intracranial pressure after intravenous administration in a porcine model of acute liver failure[186,187]. In the same animal model, L-ornithine phenylacetate increased muscular glutamine synthetase expression[186]. L-ornithine phenylacetate was intravenously administered as a continuous infusion (maximum daily dose of 10 g) in a phase I study in cirrhotic patients with an episode of upper GI bleeding in the preceding 24 h[208]. Adverse events were mild, and average plasma ammonia decreased to half the initial level after the first 36 h, and it remained in that range under infusion[208]. A significant reduction in plasma glutamine and a gradual increase in urinary phenylacetylglutamine were observed, which confirmed the mechanism of action in man[208]. However, in a clinical trial in 38 cirrhotic patients with acute upper GI bleeding, intravenous L-ornithine phenylacetate (10 g/kg) failed to decrease plasma ammonia levels [209]. In a phase IIB study involving 231 cirrhotic patients with HE stage 2 or higher, an infusion of L-ornithine phenylacetate decreased plasma ammonia levels in a dose-dependent manner [210]. Currently, a phase III clinical trial of intravenous L-ornithine phenylacetate in patients with an acute HE episode is in planning [210]. In the BDL rats, oral administration of L-ornithine phenylacetate diminished intestinal glutaminase activity and expression in the intestine, potentially via the enterocytic generation of the glutaminase-inhibitor glutamate [197]. Furthermore, a daily oral dose of 1 g/kg L-ornithine phenylacetate for 5 weeks decreased plasmatic ammonia levels in the same animal model [211]. A phase IIB study is currently in preparation comparing the effects of oral L-ornithine phenylacetate with rifaximin (NCT03712280).

Carglumic acid. Carglumic acid (N-carbamoyl-L-glutamic acid, Carbaglu®, Orphan Europe) is an FDA- approved orphan drug for the adjunctive therapy of acute and maintenance treatment of chronic HA due to N-acetylglutamate synthetase deficiency (Table 2.1), which is a rare inborn UCD[125]. Currently, carglumic acid is under investigation for other inborn metabolic disorders, such as propionic acidemia and methylmalonic academia (NCT01597440). Carglumic acid is a synthetic analogue that replaces N- acetylglutamate, the product of N-acetylglutamate synthetase (Figure 2.2)[212,213]. In contrast to N-

33 acetylglutamate, carglumic acid is resistant to degradation by cytosolic amino acylase[212,213]. Therefore, carglumic acid reaches the mitochondrion, where it acts as an allosteric activator of the first enzyme of the UC, carbamoyl phosphate synthetase 1, reactivates the genetically impaired UC, and lowers systemic ammonia[212,213]. However, the rarity of N-acetylglutamate synthetase deficiency complicates an evidence-based determination of the optimal dose and therapeutic benefit[213]. Furthermore, carglumic acid seems to be effective against HA induced by the anti-epileptic drug valproic acid, which inhibits N-acetylglutamate synthetase[125,214]. Formulated as a dispersible tablet that contains the superdisintegrant croscarmellose sodium, Carbaglu® is applied via the oral route and may also be administered through a syringe via a nasogastric tube. Carglumic acid appears very well- tolerated but the dispersion is associated with a bitter taste[213].

BCAAs. Valine, leucine, and isoleucine are classified as BCAAs because of their aliphatic branched side-chains[215]. BCAAs are substrates of the ubiquitous enzyme branched-chain aminotransferase, which catalyzes the transfer of the amine group of a BCAA to α-ketoglutarate to form a branched-chain α-keto acid and a glutamate (Figure 2.1)[215]. Glutamine synthetase subsequently transforms glutamate to glutamine under the incorporation of an ammonia molecule[215]. However, BCAA administration led to increased blood ammonia in a positron-emission tomography study[216]. Since the release of glutamine from the muscle was much higher than the uptake of systemic ammonia, the ammonia incorporated into glutamine was probably formed during the degradation of some BCAA molecules. The increase in blood ammonia may therefore be related the glutaminolysis of the released glutamine in extramuscular organs such as the kidney and the intestine[215]. Of note is that the ammonia-increasing effect of BCAAs observed in this study may stem from the administration of a single high dose instead of a regimen of multiple lower doses, which is generally practiced in the clinical setting[216]. BCAA lowered plasma ammonia in HE patients in clinical studies using split doses, which had a beneficial effect on clinical HE manifestations but no impact on mortality[217–219]. A recent systematic review pointed to the capacity of BCAA to significantly improve the mental state of HE patients[204]. Furthermore, BCAAs exert anticatabolic effects on skeletal muscles, as they stimulate hepatic protein synthesis, inhibit protein degradation, and fuel oxidative energy production in skeletal muscles[220,221]. A recent study underlined the beneficial effect of BCAA administration and the importance of skeletal muscle tissue on the prognosis of patients with liver cirrhosis[222].

L-arginine and L-citrulline. L-citrulline is an intermediate and L-arginine a product of the UC (Figure 2.2)[125]. L-arginine administration leads to the excretion of nitrogen as L-citrulline and argininosuccinate in argininosuccinate synthetase and argininosuccinate lyase deficiency, respectively, which restores the ammonia-lowering activity of the UC[127]. L-arginine also becomes an essential amino acid in all UCD, except arginase 1 deficiency[127]. The administration of L-citrulline restores L- arginine levels in deficiencies of N-acetylglutamate synthetase, carbamoyl phosphate synthetase 1, ornithine transcarbamoylase, and ornithine translocase[125,127]. Therefore, the administration of L- arginine or L-citrulline is recommended in acute UCD-induced HA and chronic management of UCDs to reactivate the UC and to reduce ammoniagenic protein catabolism because of low L-arginine levels[125,127]. Even though L-arginine infusion significantly lowered ammonia levels and reduced the risk of HE in cirrhotic patients with acute GI bleeding, it is currently not recommended for HE[126,223].

Miscellaneous. A variety of dietary supplements with potential pharmacological effects on the etiology of HA are given to patients suffering from HA[138]. It should be noted that many of these patients still develop hyperammonemic episodes and therefore their role in clinical practice remains of unclear impact and improperly defined[138]. L-carnitine mediates the transport of short-chain fatty acids across the mitochondrial and peroxisomal membranes and was reported to have several effects on HA, ranging from the activation of UC enzymes and interaction with glutamate receptors to the scavenging of free radicals[224]. Acetyl-L-carnitine, a pro-drug of L-carnitine, was observed to lower ammonia levels and improve HE symptoms in patients with mild and severe HE[225]. L-carnitine was also shown to be beneficial in reducing ammonia levels in valproate-induced HA[226], and it is given intravenously in acute HA of unknown origin[127]. Acetyl-L-carnitine is likely to have a higher bioavailability than L- carnitine because of its acetyl moiety, but evidence in human studies is lacking[227]. Moreover, zinc plays a role in ammonia metabolism as it stimulates the UC enzyme ornithine transcarbamoylase and it activates glutamine synthetase in skeletal muscle[228,229]. Several studies observed an ammonia- reducing effect of zinc administration[229–231] but a therapeutic benefit beyond an improvement in a cognitive test was not demonstrated[232] despite a very high prevalence of zinc deficiency in cirrhotic patients[230]. Moreover, the administration of biotin and hydroxycobalamin should be considered in acute HA of unknown origin because deficiencies in these essential micronutrients of the vitamin B family may increase blood ammonia levels[127]. Finally, dehydration diminishes ammonia excretion by the kidney, especially in HA precipitated by diuretics overuse[233]. Intravenous volume expansion using albumin or saline decreased blood ammonia levels in diuretic-induced HE, possibly via enhanced urinary ammonia excretion[233,234].

2.4. Novel treatment approaches

Advances in the knowledge of the pathogenesis of HA and new developments in bio-engineering have led to new strategies to treat HA. Selected promising treatment options are discussed in this section.

Conventional and albumin-based dialysis modalities. In UCD-induced HA, extracorporeal dialysis, especially continuous veno-venous hemodiafiltration, is recommended for neonates and children with severe HA or in cases of inadequate response to medical treatment[127]. Hemodialysis or hemofiltration are first-line treatments in adult UCD patients with acute hyperammonemic decompensation, and these treatments should be initiated swiftly, even if the diagnosis is not certain[127]. However, extracorporeal dialysis is currently not recommended for HE[126]. A dialysis-based treatment modality that was tested for HE is the Molecular Adsorbent Recirculating System® (MARS®, Baxter, Table 2.2), which is an extracorporeal dialysis method used to extract protein-bound compounds[235]. The free and albumin- bound substances in the extracorporeal circuit distribute across a selectively permeable membrane that contains an albumin dialysate on the trans-side[235]. The toxin-enriched albumin dialysate is purified by dialysis and adsorbing cartridges in an intermediate circuit, and then recirculated[235], which differentiates MARS from single pass albumin dialysis[236]. Clinical proof of efficacy in HE treatment of this expensive and not widely available system is lacking, and ammonia removal is not listed as an end point in the most recent clinical study (NCT02310542) [138,237–244]. Hepa Wash® (Hepa Wash) is another albumin-supported dialysis modality, which differs from the MARS® in the purification of the albumin dialysate[245]. After toxin enrichment across a semipermeable membrane, the albumin 35 dialysate enters the Hepa Wash® circuit and is split in two separate lanes[245]. One lane is acidified and the other alkalinized, leading to the dissociation of albumin-bound toxins[245]. Subsequent to the removal of the toxins by filtration, the lanes are joined, which creates a physiological pH, and reintroduced into the albumin dialysate[245]. Hepa Wash® significantly increased ammonia elimination and survival in a portacavally shunted porcine model of HE[245]. In 14 patients with liver failure, this albumin dialysis-based treatment decreased bilirubin, creatinine, and blood urea nitrogen levels compared to pre-dialysis [246]. Changes in ammonia levels were not reported [246]. In the fractionated plasma separation and adsorption system Prometheus® (Fresenius), albumin is separated from the blood using an albumin-permeable filter (cut-off 250,000 Da)[247]. The plasma protein-rich filtrate is then purified by direct exposure to a neutral and an anion-exchange resin, and subsequently reintroduced into the patient[247]. Water-soluble toxins such as ammonia are separated through a high- flux dialyzer downstream of the albumin-permeable filter[247]. Fractionated plasma separation and adsorption significantly lowered blood ammonia in patients with acute liver failure in two non-controlled studies[247,248].

Bioartificial liver support systems. To reflect the metabolic and synthetic activity of the liver, cell-based bioartificial liver support systems contain cells with hepatocytic functions[249]. In the extracorporeal liver assist device (ELAD®, Vital Therapies), conventional hemodialysis is combined with filtration through multiple hollow-fiber cartridges containing human hepatoblastoma cells (VTL C3A cell line)[250]. In a controlled clinical trial, ELAD® treatment resulted in less deterioration in HE grade in patients suffering from acute liver failure but the minor decrease in arterial ammonia was comparable to the control group[251]. A slightly altered ELAD® system showed a stabilizing effect on patients waiting for liver transplantation (acute liver failure and acute on chronic liver failure)[252,253]. Patients who had undergone ELAD® treatment until recovery showed significantly higher survival than controls[250]. However, in a randomized, multicenter clinical trial with 203 patients suffering from acute alcoholic hepatitis, ELAD® treatment had no significant effect on overall survival, thus failing to meet its primary end point [254]. Several further cell-based bioartificial liver support systems (AMC bioartificial liver®, Excorp bioartificial liver support system®, modular extracorporeal liver support system®, HepatAssist®) are under clinical investigation for liver failure and may prove to be beneficial for the treatment of HA in the future[255–258]. However, the ammonia detoxification capacity varies among different cell lines with hepatocytic functions (neoplastic cell lines derived from the liver, human or porcine primary liver cells such as fetal or mature hepatocytes, human liver stem cells)[259,260]. A critical challenge of cell- supported bioartificial liver support systems is the stability of the cells which are exposed to a variety of toxic substances (e.g., ammonia, components of necrotic cells, cytokines generated by an aberrant immune response) in the plasma of patients with acute liver failure[255].

Peritoneal dialysis. An increasingly appreciated intracorporeal dialysis method is peritoneal dialysis, in which the peritoneum acts as a membrane and the peritoneal cavity as a dialysis chamber[261]. The application of the dialysis solution and the drainage of the dialysate is carried out through a stationary abdominal catheter[261]. Even though the efficacy of peritoneal dialysis was inferior to hemodialysis in the rapid clearance of ammonia from the blood in UCD patients and systematic clinical studies are lacking[127,261], peritoneal dialysis was proposed to be a viable alternative if extracorporeal dialysis is

36 not available[262]. A dialysis solution containing transmembrane pH-gradient liposomes (i.e., vesicles bearing an internal acidic compartment, VS-01, Versantis) was recently demonstrated to efficiently capture ammonia in the peritoneal space of rats[105]. The ammonia-scavenging properties of the liposomal formulation are based on the diffusion of ammonia across the liposomal membrane and the subsequent protonation in the acidic core[105]. The positively charged ammonium remains trapped in the aqueous core because of the hydrophobicity of the phospholipid bilayer[105]. This approach increased the concentration of ammonia in the peritoneal fluid more than 20-fold after 3 h of dialysis compared to conventional peritoneal dialysis[105]. In BDL rats, five peritoneal dialysis sessions significantly lowered plasmatic ammonia concentrations and decreased brain water content compared with no treatment [36]. Furthermore, there were no major interferences with weakly basic drugs commonly used in cirrhotic patients and no significant removal of important metabolites compared with standard PD [106].

GI uptake of ammonia. Activated carbon microparticles are capable of binding toxic substances because of their high adsorptive surface area. Oral carbon suspensions are used to treat poisoning and overdoses due to the ingestion of toxic substances (Figure 2.3)[263]. Orally applied carbon adsorbent microspheres (AST-120, Ocera Therapeutics, Table 2.2) were shown to scavenge ammonia in vitro and in BDL rats, decreasing intestinal ammonia absorption and lowering blood ammonia[264]. However, clinical endpoints in patients with low-grade HE were not met despite a reduction in plasma ammonia[265]. Moreover, a W/O microemulsion containing acetic acid as an ammonia-trapping agent was found to halve the ammonia concentration in ammonia-containing artificial colonic fluid defined in the Chinese Pharmacopoeia, but the stability was low in GI-simulating media[266]. The microemulsion was not stable during GI passage in rats; droplet size increased and an emulsion was formed after ingestion[266]. The subsequent decrease in small intestinal pH was likely related to a release of acetic acid from the instable formulation, which might result in intestinal inflammation[266]. A sodium alginate- containing gel based on this microemulsion was more stable in artificial GI fluids, as no pH changes were observed in the rat GI tract[266]. The ammonia-lowering properties were similar to the alginate- free microemulsion in artificial colonic fluid[266].

Glutaminase inhibition. Glutaminase inhibitors are another promising treatment approach because they prevent ammonia-forming glutaminolysis (Figures 2.1 and 2.3). The overactivity of duodenal glutaminase and the association of a genetic marker in the glutaminase gene with HE development underline the importance of this enzyme in the etiology of HE in cirrhotic patients[118,267]. The antidiabetic biguanide metformin (Table 2.2) has glutaminase-inhibiting activity and protects against HE in diabetic patients with cirrhosis but a potential effect on ammonia levels was not investigated[268]. Metformin inhibited glutaminase only partially, reducing the risk of a potentially toxic glutamine depletion in the enterocyte[268]. Furthermore, the thiourea derivative THDP-17 lowered glutamine production in Caco-2 cells by partial glutaminase inhibition[269].

Liver tissue regeneration. A recombinant form of the human hepatocarcinoma-intestine- pancreas/pancreatitis-associated protein (ALF-5755, Alfact Innovation, Table 2.2) was demonstrated to protect hepatocytes in a mouse model of acute liver failure through its reactive oxygen species- scavenging activity[270]. The tissue regenerative functions are promising in end-stage liver disease. 37

The clinical trial listed for this indication (NCT01318525) has not been updated since 2011, though. This treatment approach may conserve and potentially increase residual functional liver tissue in cirrhosis, with possible positive effects on the ammonia detoxification capacity of the liver.

Gene therapy. The introduction of a gene coding for a functional enzyme promises to restore UC activity in patients with primary UCD. A liver-specific promoter is generally used to restrict enzyme expression to liver tissue[271,272]. The delivery of the ornithine transcarbamoylase gene via an adeno-associated virus2/8 vector (Table 2.2) was shown to restore enzymatic activity and control ammonia levels in a mouse model of ornithine transcarbamoylase deficiency[271,273]. Adeno-associated virus gene delivery of arginase 1 resulted in ammonia control and long-term survival after neonatal virus administration in arginase 1 knockout mice[272,274]. A decrease in viral copy number due to hepatocyte proliferation in the first weeks of life and a low level of ureagenesis (3.3% of controls) was reported in transfected mice[274]. Therefore, minimal enzymatic activity seems to be sufficient to achieve a therapeutic effect[274]. Another approach to lower ammonia levels consists in transfecting skeletal muscle tissue with glutamine synthetase. Intramuscular injections of a baculovirus vector carrying a glutamine synthetase gene (Table 2.2) led to a significant overexpression of this gene and lowered ammonia levels in rats with acute HE[275]. However, gene therapy faces several challenges, such as the need for stable gene expression over long periods of time, which is a prerequisite to achieve a long- term therapeutic benefit. Demonstrating the safety of the viral delivery system, upscaling the production process under good manufacturing practice-compliant conditions, and proving cost-efficiency are further issues to address.

Cell therapy. Populating the liver of UCD patients with functional hepatocytes is a promising approach to restore the activity of the UC[276]. Unfortunately, differentiated hepatocytes have several disadvantages, including poor cultivation properties, low resistance to cryopreservation, and short duration of clinical benefit after transplantation[276]. In contrast, liver stem and progenitor cells have the capability to proliferate and differentiate into hepatocyte-like cells in vivo[276–278]. They are obtained by plating hepatocytes isolated from the liver parenchymal cell fraction (i.e., from primary cell culture) of a donor, and they can be expanded in vitro on a large scale under good manufacturing practice conditions[276]. These cells are also resistant to cryopreservation[276]. Two infusions of adult-derived liver human stem cells (0.9 billion cells in total, Table 2.2) into the portal vein resulted in an implantation and proliferation of the donor cells, which grew from 0.75% to 3-5% of the liver mass in 100 days in a 3- year-old patient suffering from ornithine carbamoyltransferase deficiency[279]. After an intraportal infusion of 3 billion liver stem cells into a 17-year-old patient suffering from a glycogen storage disease, the stem cells almost exclusively homed to the liver where they remained during the course of a five- day observation period[280]. In a mouse model of an advanced nonalcoholic steatohepatitis, intravenous injections of clinical-grade human liver mesenchymal stem cells dose-dependently reduced the nonalcoholic liver disease activity score, which was concomitant with a significant decrease in inflammation[281]. These studies were part of the development of HepaStem® (Promethera), which is a medicinal product composed of heterologous human adult liver-derived progenitor cells. HepaStem® is currently in a phase II clinical trial (NCT02946554) for the treatment of acute-on-chronic liver failure.

2.5. Conclusion

Toxic increases in blood ammonia levels are associated with potentially life-threatening consequences for the patient. HA results from hepatic impairment secondary to inborn metabolic disorders or acquired conditions such as acute liver failure or liver cirrhosis (HE). Therefore, treatments should reflect the differences in the underlying diseases and precipitating factors. Large-intestinal ammonia generation in HE is targeted by non-absorbable disaccharides and antibiotics. The therapeutic benefit of non- absorbable disaccharides, which is the recommended first-line treatment, is disputed, and adverse effects are common. A new member of the antibiotic group, rifaximin, is an alternative treatment or add- on with a good efficacy and safety profile, but currently associated with higher costs. Unfortunately, a considerable fraction of treated cirrhotic patients still experience exacerbations. An acute HA episode is a medically challenging emergency irrespective of its origin and an effective therapy remains a currently unmet clinical need. Liposome-supported peritoneal dialysis is a promising versatile treatment approach for acute HA of different etiologies, especially in view of the limited data on hard clinical endpoints of other dialysis-based technologies (albumin-supported dialysis, cell-based bioartificial liver support systems). Furthermore, no therapeutic agents are currently recommended for covert HE, although this condition impairs the quality of life and bears the risk of progression to overt HE. A promising oral treatment under clinical investigation for HE is glycerol phenylbutyrate which reduces the ammoniagenic metabolic degradation of glutamine and is approved for UCDs. An interesting incentive of developing HA treatments is the potential translatability from an orphan drug setting to one of a highly prevalent condition, namely liver cirrhosis. A challenging aspect of acute and chronic UCD treatments is the extreme rarity of the disorders, which complicates an evidence-based assessment of their safety and efficacy. The functional regeneration of liver tissue using gene or cell therapy is an exciting upcoming approach to tackle liver disease and promises to bring long-term relief to patients with inborn metabolic disorders and an acquired loss of functionally viable liver tissue. However, proving clinical safety and efficacy, especially in the long-term, as well as cost-effectiveness, will be challenging.

Chapter 3

3. Investigating PS-b-PEO polymersomes for oral treatment of hyperammonemia

3.1. Introduction

Ammonia is a naturally occurring metabolite in amino acid and protein metabolism [282,283]. While it has an important role in protein synthesis and pH homeostasis, its neurotoxicity necessitates efficient ammonia-removing metabolic pathways in the body, namely the UC in the liver [282]. In subjects with impaired liver function, however, this metabolic pathway can be insufficient, resulting in a pathologic accumulation of ammonia in the blood (hyperammonemia; >50 µM and >100 µM in adults and newborns, respectively) [126,127]. Hyperammonemia is a crucial symptom in HE, a highly prevalent complication in liver disease patients (liver cirrhosis, acute-on-chronic liver failure, portosystemic shunting), and in patients with inherited UCD [126,127]. It manifests with low grade (cognitive impairments) to severe (ataxia, confusion, hepatic coma, risk of death) neuropsychiatric symptoms whose severity correlates with plasmatic ammonia levels [126,284]. The risk of exacerbation to acute hyperammonemic episodes is partly related to the severity of hyperammonemia at baseline [285].

Urease-producing bacteria in the intestine, which convert urea to ammonia and carbon dioxide, are the primary source of ammonia in the body [282,283]. Hence, ammonia-lowering treatment strategies targeting the gut are essential tools in the treatment of chronic HE [126]. Current practice guidelines recommend lactulose, a widely used laxative, as first-line treatment [126]. This non-absorbable disaccharide is metabolized by gut bacteria to short-chain fatty acids [282]. Its mechanism of action relies on its laxative action (cathartic effect), acidification of the colon, and probiotic effects which increase the excretion of ammonia and decrease ammonia generation by gut bacteria [282]. Despite its wide use in HE patients, the number of non-responding patients is high and its laxative properties negatively affect patient compliance [126,282,286]. The currently recommended second-line treatment is rifaximin (Xifaxan®, Salix Pharmaceuticals), a poorly absorbed antibiotic, which inhibits the growth and metabolism of gut bacteria [126]. However, the prolonged treatment with antibiotics bears the risk of inducing bacterial resistance [282]. Furthermore, rifaximin should be used with caution in patients with severe hepatic impairment because he low amounts of absorbed drug may only be eliminated very slowly by its main metabolizing organ, the liver [164].

In the past, the laboratory of Professor Jean-Christophe Leroux developed a treatment for acute hyperammonemia based on a transmembrane pH-gradient liposome-supported peritoneal dialysis dispersion (VS-01, Versantis) [36,105,106,287]. In the peritoneal space, these liposomes selectively sequester ammonia in their acidic cores, resulting in an efficient removal of ammonia upon extracting the peritoneal dialysis solution [36]. In BDL (i.e., cirrhotic and hyperammonemic) rats, four liposome- supported peritoneal dialysis sessions decreased systemic ammonia levels [36]. However, the invasiveness of this treatment limits its application to acute crises of hyperammonemia.

In light of the limitations of the current treatments for hyperammonemia, the aim of this chapter is to find an oral treatment for chronic HE and to test its ammonia-lowering properties in vivo in an established model of HE-associated hyperammonemia.

3.2. Material and Methods

PS-b-PEO polymer synthesis

PS-b-PEO(2000) synthesis was carried out by atom transfer radical polymerization (ATRP) [6]. PEO(2000) monomethyl ether (Sigma-Aldrich Chemie, Buchs, Switzerland) was converted to an ATRP macroinitiator by reacting it with 2-bromopropionyl bromide (Sigma-Aldrich Chemie) in dry THF (Acros Organics, New Jersey, NJ) and further used to polymerize styrene in bulk. Briefly, the ATRP macroinitiator (2.0 mmol) was loaded in a flame dried Schlenk flask, along with copper bromide (CuBr, 3.0 mmol Alfa Aesar, Ward Hill, MA) and 4,4’-dinoyl-2,2’-dipyridyl (2.64 mmol, TCI, Tokyo, Japan) as the catalyst and ligand, respectively. The Schlenk flask was evacuated and refilled with argon through several cycles to remove oxygen. In a separate flask, 11.5 mL of styrene (100 mmol, Sigma-Aldrich Chemie) was deoxygenated by bubbling argon through it for 0.5 h, and then loaded in the Schlenk flask. The mixture was then heated at 115 °C during 16 h and the brown product solution was dissolved in THF, filtered through a basic alumina column and precipitated twice in hexane. The precipitate was filtered and dried under vacuum. The feeding molar ratio of [monomer]/[initiator] was 50. The PS/PEO composition was determined in deuterated acetone by 1H nuclear magnetic resonance spectroscopy (Bruker AV-400, Billerica, MA) at room temperature. The polymer synthesis was carried out by Dr. Yinyin Bao (Laboratory of Drug Formulation and Delivery, ETH Zurich).

The PS-b-PEO polymers used in this chapter are characterized in Tables 3.1 and 3.2.

Table 3.1 Properties of PS-b-PEO diblock co-polymers purchased from Advanced Polymer Materials (Dorval, QC, Canada).

Exact molecular weight Number-average molecular Dispersity (Đ)1 composition, determined by weight (Mn) in g/mol1 NMR (PS - PEO)1

PS(1000)-b-PEO(2000) 1040 - 2000 3000 1.08

PS(1500)-b-PEO(2000) 1560 - 2000 3500 1.09

PS(2000)-b-PEO(2000) 1970 - 2000 4000 1.07

PS(2500)-b-PEO(2000) 2600 - 2000 4600 1.09

2770 - 2000 4700 1.09

PS(3000)-b-PEO(2000) 3150 - 2000 5000 1.19

PS(3500)-b-PEO(2000) 3570 - 2000 6500 1.13

PS(4000)-b-PEO(2000) 3900 - 2000 6000 1.15

PS(5000)-b-PEO(2000) 5150 - 2060 7200 1.07

PS(6000)-b-PEO(2000) 6000 - 2180 8200 1.09

1According to manufacturer

Table 3.2 Properties of synthesized PS-b-PEO diblock co-polymers.

Polymersome batch ID Exact molecular weight Number-average molecular Dispersity (Đ) composition, determined by weight (Mn)1 NMR (PS - PEO)1

F2 5500 - 2000 7400 1.45

F3 3900 - 2000 6900 1.45

F4 4300 - 2000 7200 1.53

FF1 4500 - 2000 7500 1.45

FF2 4500 - 2000 7200 1.55

N1 2800 - 2000 7200 1.48

N2 3800 - 2000 6500 1.43

N3 4300 - 2000 6300 1.43

N5 3300 - 2000 5900 1.38

N6 4100 - 2000 7200 1.44

A16 3300 - 2000 6300 1.40

Liposome preparation

Liposomes composed of (i) 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC, LIPOID, Ludwigshafen, Germany) or 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC, LIPOID, Ludwigshafen, Germany), (ii) cholesterol (Sigma Aldrich Chemie) and (iii) 1,2-distearoyl-sn-glycero-3- phosphoethanol-amine-N-[methoxy(PEO)-2000] (DSPE-PEO, LIPOID, Ludwigshafen, Germany) at 54:45:1 mol% were prepared by the film hydration method. Stock solutions of DPPC 36.7 mg/mL, DSPC 39.5 mg/mL, cholesterol 19.3 mg/mL, and DSPE-PEO 13.9 mg/mL were prepared in chloroform.

For the DSPC liposomes, 8.2 mg cholesterol, 20.2 mg DSPC, and 1.3 mg DSPE-PEO were added as chloroform stock solutions to a glass vial. For the DPPC liposomes, 19.7 mg DPPC, 8.7 mg cholesterol, and 1.4 mg DSPE-PEO, were added as chloroform stock solutions to a glass vial. The organic solvent was subsequently removed by nitrogen flow for 2 h and storage in a vacuum desiccator overnight. The dried film was hydrated with 1 mL of citric acid solution 250 mM at pH 2.0 at 300 mOsmol/kg (lipid concentration = 29.8 mg/mL) while heating to 55°C and slowly mixing until the lipid film was not visible any more.

Polymersome preparation

PBD(2500)-b-PEO(1500) (number-averaged molecular weights, Đ 1.04, Polymer Source, Dorval, Canada) and PS-b-PEO polymersomes were produced using an oil-in-water (o/w) emulsification method. All polymers (60 mg) except PS(6000)-b-PEO(2000) were dissolved in 100 µL of dichloromethane or chloroform (both Sigma Aldrich Chemie). PS(6000)-b-PEO(2000) (20 mg) was dissolved in 100 µL of toluene (Sigma Aldrich Chemie). The polymer solution was added dropwise to 1 mL citric acid (Sigma Aldrich Chemie) solution 250 mM at pH 2.0 at 300 mOsmol/kg (unless stated 42 otherwise) under sonication in an ice bath using the following parameters: amplitude 70, cycle 0.75 (UP200H, 200W, 24 kHz equipped with sonotrode S1, Hielscher Ultrasonics, Teltow, Germany) for 3 min or amplitude 10 (3.1 mm sonotrode, Fisher Scientific Model 705 Sonic Dismembrator, 700W, 50/60 Hz, Fisher Scientific, Reinach, Switzerland) for 2 min. The organic solvent was evaporated using a rotary evaporator for at least 5 min at 40°C at 70 kPa.

In the scaled-up method, 1.84 g PS-b-PEO polymer (diblock copolymers from Supplementary Table S6; PS-b-PEO polymer batches with different PS fragment length were never pooled prior to polymersome preparation) were dissolved in 1.84 mL dichloromethane. The polymer solution was added dropwise to 16 mL citric acid solution 250 mM at pH 2.0 at 300 mOsmol/kg under sonication in an ice bath using the following parameters: initial amplitude 50 (6 mm sonotrode, Fisher Scientific Model 705 Sonic Dismembrator) during the addition of the oil phase (approx. 1 min) and subsequent sonication at amplitude 60 for another 3 min. The organic solvent was evaporated using a rotary evaporator for at least 10 min at 40 °C at 60 kPa. The polymersome batches were subsequently pooled. The concentration of dichloromethane in the pooled polymersome batches was further reduced by rotary evaporation at 40 °C and 22 kPa for 8 to 13 h. The residual dichloromethane content as determined by head-space gas chromatography (see section 3.2) ranged between 0.13 and 0.33 mg/mL.

The dichloromethane content was determined using headspace gas chromatography coupled to a flame ionization detector. The samples were analyzed by headspace injection (TurboMatrix HS 40, Perkin Elmer, Waltham, MA) and measured by HP6890 (Hewlett Packard, Palo Alto, CA) equipped with a TR- FFAP column (50 m x 0.32 mm x 0.5 μm, Thermo Fisher Scientific). The samples were heated for 20 min at 35 °C and the HS needle and transfer line were set to 90 and 120 °C, respectively. Samples were injected for 0.04 min at a total flow of 40.3 mL/min using a split flow of 36.5 mL/min (split ratio 20:1, pressure 90 kPa). Helium (Pangas, Dagmarsellen, Switzerland) was selected as the carrier gas. The pressure in the column was constant (90 kPa) and the flow 1.8 mL/min. The oven temperature was set to 45°C for the first 2 min and gradually increased to 130°C over the next 6 min. The FID detector was heated to 250°C with a hydrogen flow of 55 mL/min and an airflow of 300 mL/min.

In the sonication-enhanced nanoprecipitation method, PS(2500)-b-PEO(2000) (60 mg) was dissolved in 100 µL of THF. The polymer solution was added dropwise to 1 mL citric acid solution 250 mM at pH 2.0 at 300 mOsmol/kg under sonication in an ice bath at amplitude 70, cycle 0.75 (UP200H, 200W, 24 kHz equipped with sonotrode S1, Hielscher Ultrasonics, Teltow, Germany) for 3 min. The organic solvent was evaporated using a rotary evaporator for at least 5 min at 40°C at 70 kPa.

PS-b-PEO polymersomes were also produced using a film hydration method. PS(2500)-b-PEO(2000) (30 mg) was dissolved in 100 µL of dichloromethane and added to a glass vial. The organic solvent was subsequently removed by nitrogen flow for 2 h and storage in a vacuum desiccator overnight. The dried film was hydrated with 1 mL of citric acid solution 250 mM at pH 2.0 at 300 mOsmol/kg under heating to 65°C and sonication for 1.25 h.

The hydrodynamic diameter of the PS(3500)-b-PEO(2000) polymersomes was determined using the Mastersizer2000 laser diffraction particle size analyzer (Malvern Instruments, Herrenberg, Germany). The measurements are presented as volume distribution (mean of three measurements). The 43 morphology of the PS-b-PEO polymersomes was further analyzed by cryo-scanning electron microscopy (SEM). In a high pressure freezer, the polymersome dispersion in citric acid solution 250 mM at pH 2.0 was vitrified at -160 °C and mechanically broken. The resulting surface was coated and imaged by SEM (Zeiss Gemini 1530, Zeiss, Oberkochen, Germany) according to the literature [288].

Polymer quantification

The PS-b-PEO polymer concentration in the polymersome dispersion was determined after dilution in N,N-dimethylformamide (final polymersome concentration 5% (v/v) for 1 mL and 2.5% (v/v) for 16 mL set-up) by spectrophotometry at 271 nm (Tecan Infinite 200 Pro, Maennedorf, Switzerland). Alternatively, the PS-b-PEO and PBD-b-PEO polymer concentration in the polymersome dispersion was determined after lyophilization and resuspension in N,N-dimethylformamide with gel-permeation chromatography with the same organic solvent (0.5 mL/min) as mobile phase equipped with a at 35°C two ViscoGEL columns (GMHHR-M, poly(styrene-co-divinylbenzene)) and coupled to a refractive index detector (Viscotec GPCmax VE-2001 with Viscotek TDA 302 Triple Detector Array, Viscotek, Malvern Panalytical Ltd, Malvern, UK). For the molecular weight and dispersity (Đ) determination, the polymers were dissolved in THF at 2 mg/mL and analyzed by refractive index measurement, comparing to poly(methylmethacrylate) standards (2500- 89300, PSS polymer, Mainz, Germany).

Ammonia uptake in side-by-side cells

The transmembrane pH-gradient was generated in side-by-side diffusion cells (PermeGear Inc., Hellertown, PA) maintained at 37°C by dilution in phosphate buffer (final concentration 50 mM) at pH 6.8 and 300 mOsmol/kg in the absence and presence of the bile salts sodium cholate (SC), sodium deoxycholate (SDC; bile salts for microbiology, Sigma Aldrich Chemie, 1:1 mixture of SC and SDC), and sodium taurocholate (STC, abcr, Zug, Switzerland) at 25/25/0 mM (liposome, PBD-b-PEO, and hypoosmolal PS-b-PEO experiments), 12.5/12.5/0 (digestive enzyme experiment), or 30/30/30 mM (other experiments). In the hypoosmolal conditions, the osmolality was set to 160 mOsmol/kg by modifying the sodium chloride concentration and the phosphate concentration to 10 mM. In the hyperosmolal conditions, the osmolality was set to 620 mOsmol/kg by modifying the sodium chloride concentration. In the digestive enzyme experiment, trypsin from porcine pancreas 1 mg/mL (approximately 10 000 IU/mL, Sigma Aldrich Chemie), α-chymotrypsin from bovine pancreas (Type II) 1 mg/mL (approximately 40 IU/mL, Sigma Aldrich Chemie), lipase from porcine pancreas (Type II) 3 mg/mL (approximately 300 IU/mL, Sigma Aldrich Chemie) were added to the bile salt-containing phosphate buffer.

The dual-chamber system was separated by a track etch polycarbonate membrane (pore size = 100 nm, Sterlitech, Kent, WA), physically isolating the vesicles on one side. The lipid or polymer and the ammonia concentrations within the diffusion cells were 1.75 mg/mL and 1.5 mM (unless stated otherwise, Sigma Aldrich Chemie), respectively. At the allotted time, aliquots were taken from the vesicle-free compartment. The ammonia concentration of the samples was determined by the Berthelot reaction. The ammonia capture capacity was quantified using equation 1 with the total mass of sequestrant being the total mass of lipid or polymer:

Ammonia capture capacity = Total ammonia (µmol) – free ammonia (µmol) (1) Total mass of sequestrant (g)

Ammonia uptake with microparticles

To evaluate their ammonia uptake, zeolite (Sigma Aldrich Chemie), PMAA (Lewatit CNP-105, Sigma Aldrich Chemie), mesostructured silica (Sigma Aldrich Chemie), or activated charcoal (Norit, Cabot, Alpharetta, GA) microparticles were incubated at 30 mg/mL in phosphate buffer 50 mM (pH 6.8, 37°C) with an ammonia concentration of 1.5 mM under orbital shaking. At the allotted time points, the dispersions were filtered with a 0.2-µm syringe filter and the ammonia concentration of the samples was determined by the Berthelot reaction. The ammonia capture capacity was quantified using equation 1 with the total mass of sequestrant being the total mass of microparticles.

The cation competition experiments with zeolite microparticles (20 mg/mL) were conducted similarly with a modified buffer (HEPES 100 mM at pH 7.0 supplemented with 50 to 2000 mM sodium, potassium, or calcium) and an incubation time of 30 min at 37°C.

Dietary hydrogel experiments

PS-b-PEO(2000) polymersomes with PS fragments between 3300 and 4500 (Table 3.2) were used for the dietary hydrogel experiments. PS-b-PEO polymersomes were incubated in an isotonic phosphate buffer 10 mM at pH 6.8 with different Metamucil® (Metamucil® Regular, Procter & Gamble Switzerland, Petit-Lancy, Switzerland) concentrations (5-50%, m/m) for 24 h at 37°C. Subsequently, the hydrogel was diluted 1:18.5 (m/v) in isotonic phosphate buffer 50 mM at pH 6.8 (final PS-b-PEO polymer concentration: 1 mg/mL) containing 1.5 mM ammonia. After 3 h incubation at 37°C under orbital shaking, the dispersions were centrifuged for 5 min at 4500 x g. After filtration with a 0.2-µm syringe filter, the ammonia concentration of the solution was determined with the L-glutamate dehydrogenase-based enzymatic ammonia kit (AM1015, Randox Laboratories, Crumlin, UK).

The preparation of the fluorescent PS-b-PEO polymersomes was analogous to the non-fluorescent ones except for the modified polymer amount (30 mg) and inner phase: HPTS 10 mM in isotonic citric acid 250 mM at pH 2.0 for HTPS-containing transmembrane pH gradient polymersomes and HPTS 10 mM and p-xylene-bis(N-pyridinium bromide) (DPX) 30 mM in isotonic phosphate buffer 50 mM at pH 6.8 for HTPS/DPX-containing polymersomes. To account for incomplete extraction from the hydrogel matrix, the fluorescence intensity of DPX/HPTS-containing polymersomes were corrected with the extraction efficiency of HPTS-only polymersomes (Figure 3.1). To remove the free dye, the polymersomes were purified using a MidiTrap G-25 column (GE Healthcare, Glattbrugg, Switzerland) with an isotonic phosphate buffer at pH 6.8 [289].

Figure 3.1. Normalized fluorescence recovery after incubation of HPTS-containing polymersomes without transmembrane pH gradient in Metamucil©-based hydrogels at pH 6.8. Fluorescence emission intensity normalized to the one at 0% Metamucil® in relation to Metamucil® concentration. Dye concentration: 30 µM. Inner phase: isotonic phosphate buffer 50 mM at pH 6.8. All results as means ± SD (n = 3).

The dye concentration was determined upon dilution to 5% (v/v) in dimethylformamide and the measurement of the fluorescence emission intensity at 510 nm excited at 413 nm (Tecan Infinite 200 Pro). The final dye concentration for the HTPS-containing transmembrane pH gradient polymersomes and the HTPS/DPX-containing polymersomes 50 µM and 30 µM, respectively. The fluorescence emission at 510 nm excited at the isosbestic wavelength 413 nm was measured (Tecan Infinite 200 Pro). For the fluorescent transmembrane pH gradient polymersomes, additionally the pH-dependent excitation at 455 nm at the same emission wavelength was measured as well (Tecan Infinite 200 Pro).

Lyophilization

PS(2500)-b-PEO(2000) polymersomes frozen in liquid nitrogen and lyophilized (Alpha 2-4 LSC, Christ, Osterode am Harz, Germany) over 24 h. The volume of sublimated water was subsequently readded and the dispersion mixed and sonicated in an ultrasound bath for 10 min. The ammonia uptake was subsequently analyzed in bile salt-containing medium in side-by-side diffusion cells (see above).

In vivo experiments

The animal experiments adhered to national guidelines and were approved by the local ethics committee of the Centre de Recherche du CHUM (CIPA, Montreal, QC, Canada). Secondary biliary cirrhosis was induced in 38 male Sprague‐Dawley rats (ca. 220 g) (Charles River Laboratories, St. Constant, QC, Canada) by BDL.

Firstly, the effect of orally applied polymersomes on blood ammonia levels was evaluated in the absence of laxatives. 31 days after BDL surgery, two groups of cirrhotic rats (n = 4 per group) received

46 polymersome dispersion in citric acid solution (twice daily 0.9 g polymer/kg and 0.48 g citric acid/kg) or citric acid solution alone (twice daily 0.48 g citric acid/kg) by gavage for five days. A group of healthy rats (n = 2) served as control. Blood ammonia levels were measured in fresh blood samples from the saphenous vein 31 and 36 (at sacrifice) days after BDL surgery and in healthy control rats using the PocketChem BA (mode F1). The weight of the rats was recorded daily during the treatment.

Secondly, the laxative effects of poly(ethylene glycol) (3350) (PEG, 1 g/kg, n = 3, Pegalax®, Aralez Pharmaceuticals, Mississauga, ON, Canada), sodium picosulfate (25 mg/kg, n = 19, i.e., same animals used in second part of study, Dulcolax® Picosulfate, Sanofi Avenis, Vernier, Switzerland), and the negative control (water, n = 6) were determined in BDL rats. The laxatives were diluted in water and gavaged once daily at 8.30 a.m. (5 mL/kg) in BDL rats at three weeks after surgery. Fresh stool samples were collected after 8 and 24 h and dried at 70°C overnight. The weight loss of the stool samples was quantified gravimetrically.

Finally, the effects of the polymersomes (1 g/kg per day, diblock copolymers from Table 3.2) was assessed in BDL rats supplemented with sodium picosulfate (25 mg/kg per day). Three weeks after surgery, blood was sampled from the saphenous vein and centrifuged at 1500 x g at 4°C for 7 min after addition of heparin (Sandoz Canada, Boucherville, QC, final concentration approx. 20 IU/mL). The ammonia levels in the heparinized plasma were immediately measured with the PocketChem BA PA- 4140 (20 µL sample, mode F6; PocketChem BA PA-4140, Arkray, Kyoto, Japan). Two groups with similar ammonia levels were created (n = 18-19 per group) which received sodium picosulfate (once daily 25 mg/kg) and polymersome dispersion in citric acid solution (twice daily 0.5 g polymer/kg) or citric acid solution alone (twice daily 0.24 g citric acid/kg, i.e., corresponding to citric acid dose received by the polymersome group) by gavage twice daily (10.30 a.m. and 5 p.m.) for one week (total gavage volume: 15 mL/kg per day). A group of age-matched healthy rats (n = 6) served as control and received citric acid solution (twice daily 0.24 g/kg) by gavage for one week (15 mL/kg per day). At day seven, the second dose was administered at 1 p.m. and the rats were sacrificed at 3.30 p.m. Arterial blood was collected by cardiac puncture and centrifuged at 1500 x g at 4°C for 7 min after addition of heparin (final concentration 10 IU/mL). The ammonia levels in the heparinized plasma were immediately measured with the PocketChem BA (20 µL sample, mode F6). The three deaths in the polymersome group and the death in the citric acid group occurred right after the gavage procedure. The weights of the rats were measured daily in the fourth week after surgery.

Ammonia assays

With the Berthelot reaction, the ammonia-containing sample was added to equivolumetric amounts of alkaline sodium hypochlorite and phenol nitroprusside solutions (both from Sigma Aldrich Chemie). After 25 min of incubation at room temperature, the absorbance was measured at 636 nm (Tecan Infinite 200 Pro).

With the enzymatic ammonia assay (AM1015, Randox), the ammonia concentration was measured based on the manufacturer’s instructions with a modified sample, reagent buffer, and enzyme solution volume (20/200/2 µL or 30/300/3 µL) in a 96-well plate [36,106]. In brief, the ammonia-containing solution, blank, or calibrator was added to the reagent buffer (trometamol 150 mM at pH 8.6) in the well 47 and incubated for 5 min at room temperature. After measuring the baseline absorbance at 340 nm (Tecan Infinite 200 Pro), the enzyme was added and the absorbance at the same wavelength was measured after 5 min incubation at room temperature.

With the PocketChem BA, 20 µL of plasma or whole blood sample was applied to a strip (Ammonia Test Kit II, Arkray) and measured after 3 min using mode F6 (plasma) or F1 (whole blood).

Statistical analysis

The statistical calculations were carried out by SigmaPlot (version 13.0). For the comparison of two groups, an unpaired t-test was used. For the comparison of three or more groups, a one-way ANOVA (Holm-Sidak test) assuming a normal distribution of the data was used. In ammonia uptake kinetics with side-by-side diffusion cells, the time point at 24 h was compared between groups with this test. A p- value of <0.05 was deemed statistically significant.

3.3. Results and Discussion

Screening of microparticle and vesicular formulations

To identify a formulation for ammonia detoxification in the intestine, several strategies based on microparticles and transmembrane pH-gradient vesicles were tested in buffers and simulated intestinal fluids (Figures 3.1, 3.2). A series of organic and inorganic microparticles, which bind ammonia with ionic and hydrophobic interactions as well as hydrogen bonds, was evaluated. Of the four tested microparticle systems, aluminosilicate-based zeolites exhibited the highest uptake capacity with 40.0 ± 1.7 µmol ammonia/g material after 4 h at pH 6.8 and 37°C (Figure 3.2A). This value was slightly higher than the mesoporous carbon adsorbent microparticles AST-120 (approx. 25 µmol ammonia/g as calculated from reference [264]), an ammonia-sequestering agent which was reported to decrease plasmatic ammonia levels in BDL rats upon oral administration [264]. Co-incubating zeolites with mono- and divalent cations (potassium, sodium, calcium) led to a concentration-dependent displacement of the ionized form of ammonia (Figure 3.2B). Therefore, we assume that ionic binders such as zeolites bind physiologically relevant cations in vivo, resulting in a risk of reduced ammonia capture and, potentially, local and systemic electrolyte imbalances [290–292].

Figure 3.2. Ammonia sequestration by microparticles at near-neutral pH and 37°C. Ammonia capture capacity of different microparticles (activated charcoal, PMAA, mesoporous silica, zeolite) at 30 mg/mL 48

(A). Buffer composition: phosphate buffer at pH 6.8; ammonia concentration: 1.5 mM; n = 3. Normalized ammonia capture capacity of zeolite microparticles at 20 mg/mL in the presence of increasing cation (potassium, sodium, calcium) concentrations (B). Buffer composition: HEPES buffer 100 mM at pH 7.0; ammonia concentration: 1.0 mM; n = 3-13. All results as means ± SD. *p < 0.05, **p < 0.01, and ***p < 0.001.

In the past, we developed a highly efficient and relatively selective transmembrane pH-gradient liposome formulation for systemic ammonia removal via peritoneal dialysis [36,105,106]. With regard to their high ammonia uptake capacity and selectivity [106], we subsequently decided to investigate the utility of transmembrane pH-gradient vesicles in intestinal ammonia capture. Conceptually, the ammonia generated by urease-producing bacteria in the gut diffuses through the amphiphilic membrane bilayer into the acidic core of the vesicle (Figure 3.3). Ammonia is subsequently protonated to ammonium whose diffusion out of the vesicle is hindered because of its positive charge. Thus, ammonia molecules are stably sequestered and excreted with the feces.

Figure 3.3. Schematic representation of ammonia generation by urease-producing intestinal bacteria and ammonia sequestration by transmembrane pH-gradient vesicles.

However, the GI tract is a harsh environment for vesicular structures due to the high bile salt concentrations, extreme osmolarity and pH values, and high enzymatic activity (e.g., phospholipases) [101]. With regard to these challenges, vesicles composed of lipid or diblock copolymers with low or high phase transition temperatures were tested in simple buffers and bile salt-containing media. As liposomes made of phospholipids with phase transition temperatures (Tm) below 37°C are easily destabilized, two phospholipids DPPC (Tm = 41°C) or DSPC (Tm = 55°C) with relatively high Tm were chosen. Low or high glass transition temperature (Tg) non-biodegradable diblock copolymers of PEO and PBD (Tg = -92°C [293]) or PS (Tg = 97°C [294]) were selected for the polymersomes. All formulations showed an appreciable ammonia uptake in isotonic phosphate buffer at pH 6.8 (Figure 3.4). In the presence of physiologically relevant bile salt concentrations [295], however, liposomes were destabilized (Figure 3.4A), and only the polymersomes made of the high Tg diblock copolymer PS-b- PEO preserved their ammonia capture capacity (Figure 3.4B). After 4 h incubation in the bile salt-

49 containing medium at pH 6.8 and 37°C, the capture capacity of PS-b-PEO polymersomes was 416 ± 36

µmol NH3/g polymer which exceeded the capture capacity of AST-120 microparticles (approx. 25 µmol

NH3/g [264]) by more than 15-fold. As reported in the literature, the destabilization of liposomes by bile salts is mediated by the insertion of the amphiphiles into the bilayer and the partitioning of phospholipids into the bile salt micelles [85–90]. PBD-b-PEO polymersomes were reported to be destabilized by surfactants with a similar mechanism [99]. In contrast, the high glass transition temperature of PS yields a much tougher membrane. To the best of our knowledge, this study is the first to provide evidence on the high stability of PS-b-PEO polymersomes with short PS fragments. One study reported that PS-b- PEO polymersomes composed of very long PS fragments (>25’000) were resistant to surfactants [45]. However, the vesicle preparation methods of polymersomes with such large hydrophobic fragments were associated with low yields and water-miscible organic solvents [1,45].

Figure 3.4. Ammonia sequestration by transmembrane pH-gradient vesicles. Ammonia capture capacity of cholesterol-containing DPPC and DSPC liposomes at 1.75 mg/mL in an isotonic buffer at pH 6.8 with or without bile salts (A). Ammonia capture capacity of polymersomes made of PBD(1500)-b-PEO(2000) or PS(5000)-b-PEO(2000) at 1.75 mg/mL in an isotonic buffer at pH 6.8 with or without bile salts (B). Bile salt composition: cholate and deoxycholate, 25 mM each, for PBD(1500)-b-PEO(2000) polymersomes and liposomes; cholate, deoxycholate, and taurocholate, 30 mM each, for PS(5000)-b- PEO(2000); ammonia concentration 1.5 mM; temperature: 37°C. All results as means ± SD (n = 3). *p < 0.05, **p < 0.01, and ***p < 0.001.

Optimization and characterization of PS-b-PEO polymersomes

To identify the optimal preparation method for PS-b-PEO polymersomes, film rehydration, nanoprecipitation and emulsification processes were investigated. Only emulsification with a water- insoluble solvent under sonication led to a highly concentrated polymersome formulation (>30 mg/mL) with high ammonia uptake in bile salt-containing media (Figure 3.5A). While both dichloromethane and chloroform are suitable solvents for this method (Figure 3.5A), dichloromethane was selected due to its lower toxicity and higher vapor pressure (i.e., easier removability) [296]. With this method, highly concentrated PS-b-PEO polymersome dispersions with polymer concentrations of up to 107.3 ± 3.5 mg/mL (n = 6) and a batch volume of 16 mL could be prepared. PS-b-PEO polymersome solutions with high ammonia uptake could not be produced by the same process using a miscible solvent (Figure 3.5A). Indeed, nanoprecipitation-based methods reported for PS-b-PEO polymersomes would not be 50 adapted for a large scale preparation method due to the low speed of the procedure and the low polymer concentrations (typically <2.5 mg/mL) [6,24,25]. Furthermore, potential difficulties in removing the water- miscible organic solvent from the dispersion could result in toxic solvent residues. Impaired membrane stability may further result from these residues due to their plasticizing properties (i.e., enhancing membrane fluidity and permeability) [25]. The film rehydration method did not yield polymersomes with an appreciable ammonia uptake in bile salt-containing media (Figure 3.5A). Concentrated dispersions of well-structured polymersomes were not formed because the PS fragment is presumably too hydrophobic for efficient hydration and rapid partition from the solid into the water phase.

To determine the optimal citric acid concentration in the polymersome core, the ammonia uptake capacity of polymersome formulations containing different citric acid concentrations at pH 2.0 was determined. This pH is generally regarded as safe for ingestion as it is similar to the pH values of widely consumed beverages (e.g., fruit juices, soft drinks) [297]. All investigated citric acid concentrations (100 – 500 mM) led to the significantly higher ammonia uptake than the negative control (PS-b-PEO polymersomes without transmembrane pH-gradient) after 24 h of incubation in bile salt-containing buffer at pH 6.8 (Figure 3.5B). An isotonic citric acid solution of 250 mM was selected for further analyses, as it was the best compromise between ammonia capture capacity, citric acid exposure, and osmolality. This concentration showed a significantly higher capture capacity than the lowest (100 mM, p=0.016) and highest citric acid concentrations (500 mM, p<0.001). The lower uptake capacity for the 100 mM citric acid concentration may be associated with its low buffering capacity which leads to a rapid increase in core pH upon ammonia influx and a subsequent loss of the transmembrane pH gradient. Probably due to the same reason, the capture capacity of PS-b-PEO polymersomes with a citric acid concentration of 250 mM at a core pH of 3.0 was lower than with pH 2.0 (Figure 3.5C). Concerning the 500 mM citric acid concentration, its hyperosmolarity may impede the influx of solutes such as ammonia into the core due to the unfavorable osmolality gradient across the membrane. Furthermore, the ammonia capture capacity of PS-b-PEO polymersomes was similar at different physiologically relevant ammonia concentrations (Figure 3.5D) [298,299].

Figure 3.5. Preparation and characterization of PS-b-PEO polymersome formulations. Ammonia capture capacity of PS-b-PEO polymersomes prepared with film rehydration, sonication-enhanced nanoprecipitation in THF, and sonication-enhanced emulsification in chloroform and dichloromethane in a bile salt-containing isotonic medium at pH 6.8 (A). Ammonia capture capacity of PS-b-PEO polymersomes with different citric acid concentrations in the core (100 to 500 mM) at pH 2.0 and control PS-b-PEO polymersomes without a transmembrane pH gradient (core pH 6.8) in an isotonic bile salt- containing buffer at pH 6.8 (B). Ammonia uptake capacity of PS-b-PEO polymersomes with a citric acid solution 250 mM at pH 2.0 or 3.0 and in an isotonic bile salt-containing buffer at pH 6.8 (C). Ammonia uptake capacity of PS-b-PEO polymersomes incubated with increasing concentrations of ammonia (1.5 to 6 mM) (D). Bile salt composition: cholate, deoxycholate, and taurocholate, 30 mM each; polymer concentration: 1.75 mg/mL; ammonia concentration 1.5 mM. All results as means ± SD (n = 3-8). *p < 0.05, **p < 0.01, and ***p < 0.001.

Cryo-SEM analysis confirmed the vesicular morphology of the polymersomes (Figure 3.6A). Laser diffraction measurements revealed a volume distribution in the low micron range with a D50 at 5.1 µm (Figure 3.6B). Submicron-sized vesicles are suitable for local delivery in the intestine due to their poor diffusion across the intestinal mucus layer and the intestinal membrane [300,301].

Figure 3.6. Morphology and size distribution of PS-b-PEO polymersomes. Cryo-SEM image of PS-b- PEO polymersomes in isotonic citric acid solution 250 mM at pH 2.0 (A). Volume distribution of PS-b- PEO polymersomes at different hydrodynamic radii as determined laser diffraction (B). The mean D10, D50, and D90 values were 2.4 µm, 5.1 µm, and 11.2 µm, respectively.

Evaluation of PS-b-PEO library in simulated intestinal fluids

Having identified PS-b-PEO polymersomes containing an isotonic citric acid 250 mM solution at pH 2.0 as the lead formulation, a library of bile salt-resistant PS-b-PEO(2000) polymersomes was created to select the optimal PS fragment length. PS-b-PEO(2000) polymersomes with PS fragments between 2500 and 6000 showed a preserved ammonia uptake over 24 h in a solution containing 90 mM bile salts at pH 6.8 (Figure 3.7A). These bile salt concentrations exceed extreme physiological ones two-fold [295]. PS-b-PEO(2000) polymersomes with PS fragments lower than 2500 showed a very low ammonia uptake capacities, potentially due to the formation of instable polymersomes or micelles. PS(2000)-b- PEO(2000) formed instable polymersomes as ammonia sequestration was observed in phosphate buffer but not in bile salt-containing media (Figure 3.7B). Therefore, a certain length of the PS fragment seems to be needed for resistance against bile salts at a concentration of 90 mM. While PS(6000)-b- PEO(2000) polymersomes took up ammonia in the presence of bile salts, the capture capacity was comparably low, potentially due to more aggregation (i.e., formation of fewer vesicles) of the long, highly hydrophobic PS fragment during the preparation process. Polymersomes composed of PS fragments of 2500 and 5000 were further exposed to bile salt-containing media of physiologically relevant extreme hypo- and hyperosmolality (160 and 620 mOsmol/kg [91–93]). In these media, the polymersomes exhibited a similar capture capacity as under isotonic conditions (Figure 3.7C). PS(5000)-b-PEO(2000) polymersomes further took up ammonia in the presence of a mixture of the enzymes trypsin, chymotrypsin, and lipase and bile salts (Figure 3.7D). As proteins undergo ammoniagenic degradation reactions (aminolysis) in solution [302], the incubation time was limited to 4 h. A library of PS-b- PEO(2000) polymersomes is therefore provided with high resistance to key vesicle-destabilizing features of the GI tract (i.e., extreme bile salt concentrations and osmolality, digestive enzymes). For the subsequent in vitro and in vivo experiments, a PS fragment of approx. 3500 was selected (Table 3.2) due to its comparably low hydrophobicity (i.e., high polymersome yields) and its distance to the molecular weight range around 2000 where instable polymersomes are formed.

Figure 3.7. Ammonia uptake of PS-b-PEO polymersomes in simulated intestinal fluids. Ammonia capture capacity of PS-b-PEO(2000) polymersomes with PS fragments ranging from 1000 to 6000 in an isotonic bile salt-containing phosphate buffer at pH 6.8 (A). Capture capacity of PS(2000)-b- PEO(2000) polymersomes in isotonic phosphate buffer at pH 6.8 with and without bile salts (B). Capture capacity of PS(5000)-b-PEO(2000) and PS(2500)-b-PEO(2000) polymersomes incubated in hypo- (160 mOsmol/kg), norm- (300 mOsmol/kg), and hyperosmolal (620 mOsmol/kg) bile salt-containing phosphate buffer at pH 6.8 (C). Capture capacity of PS(5000)-b-PEO(2000) corrected for ammoniagenic protein degradation upon incubation in a trypsin- (1.0 mg/mL), chymotrypsin- (1.0 mg/mL), lipase- (3.0 mg/mL), and bile salt-containing buffer at pH 6.8 (D). Bile salt composition in enzyme-free isotonic and hyperosmolal solutions: cholate, deoxycholate, and taurocholate, 30 mM each; in hyposomolal solution: phosphate buffer 10 mM, cholate and deoxycholate, 25 mM each; in enzyme-containing solution: cholate and deoxycholate, 12.5 mM each; polymer concentration: 1.75 mg/mL; ammonia concentration 1.5 mM; temperature 37°C. All results as means ± SD (n = 3). *p < 0.05, **p < 0.01, and ***p < 0.001.

Stability in dietary fiber-based hydrogels

To simulate the environment of the distal GI tract, where the feces thicken due to water resorption, the polymersomes were incubated in dietary fiber-based (Metamucil®, psyllium husk) hydrogels at pH 6.8. The Metamucil® concentrations employed in this assay (5-50% (m/m)) strongly exceeded typical concentrations of hydrogel-forming agents (approx. 2% (m/m)) in liposome-containing hydrogels [303]. After 24 h of incubation, the polymersomes were exposed to an ammonia-containing solution and showed a Metamucil® concentration-dependent decrease in capture capacity (Figure 3.8A). To investigate the destabilization mechanism in detail, the pH-dependent fluorescent dye HPTS was

54 encapsulated in the polymersome core containing citric acid solution 250 mM at pH 2.0. With increasing Metamucil® content, the fluorescence emission ratio rose up to values observed for control polymersomes without a transmembrane pH gradient (Figure 3.8B). The rise in fluorescence emission ratio, which indicates a pH increase in the environment around the fluorescent dye [304], could either stem from a loss of the pH gradient across the polymersome membrane or the release of the dye into the outer phase at pH 6.8. To investigate the leakage of the dye from the polymersome, the dye/quencher pair HTPS/DPX was subsequently incorporated in the core of PS-b-PEO polymersomes without transmembrane pH gradient. As DPX is a collisional quencher of HPTS, an increase in HPTS fluorescence emission indicates a spatial separation of the dye/quencher pair (i.e., loss of encapsulated dye and/or quencher due to impaired membrane integrity) [289]. A Metamucil® concentration-dependent increase in fluorescence intensity was observed which reached up to 30% of a corresponding polymersome-free DPX/HPTS solution (positive control) (Figure 3.8C). Therefore, we observed a disruption of the polymersome membrane leading to an at least partial loss of the transmembrane pH- gradient and the encapsulated cargo in a dietary fiber-based hydrogel. The polysaccharide-based hydrogel-forming dietary fibers probably bound a large fraction of the free water molecules, resulting in a strong osmolality increase and a subsequent destabilization of the membrane. This hypothesis of a loss of membrane integrity upon dehydration is supported by a lyophilization experiment in which a lyophilized polymersome dispersion showed a very low ammonia uptake capacity after resuspension (Figure 3.8D). The rather high water permeability of PS could allow an outflow of water in extremely hyperosmolal environments which may impair the polymersomes’ membrane integrity and result in the loss of the transmembrane pH gradient [24,305]. In addition to the increase in osmolality, the ordered structure of the hydrogel might cause mechanical stress on the polymersome membrane. As the flexural modulus generally decreases with vesicle size [306], the limited mechanical resistance may be partially explained by the large size of the polymersomes investigated here. Unfortunately, decreasing the size of PS-b-PEO polymersomes has only been achieved using considerable amounts of water-miscible organic solvents as plasticizers [25]. We have tried several methods to decrease the polymersome size without adding large amount of water-miscible solvents (extrusion, changes in sonication parameters and solvent composition, high-pressure homogenization) without success. Microfluidic methods could be used to yield defined, submicron-sized polymersomes but the scale-up to large volumes and highly concentrated polymersome dispersions would be challenging [307,308]. With regard to stability and safety considerations associated with water-miscible organic solvents (see above), this approach seems inappropriate for the present study. In the light of these results, we decided to use a laxative agent, which increases the water content in the stools, as an add-on to the PS-b-PEO polymersome treatment in the subsequent in vivo experiments.

Figure 3.8. Stability of PS-b-PEO polymersomes in dietary fiber (Metamucil®)-based hydrogels and after lyophilization. Ammonia capture capacity of transmembrane pH-gradient PS-b-PEO polymersomes after 24 h incubation in dietary fiber-based hydrogels at Metamucil® concentrations of 0 to 50% (m/m) (A). Fluorescence emission ratio (FR) of transmembrane pH-gradient HPTS-containing PS-b-PEO polymersomes after 24 h incubation in dietary fiber-based hydrogels (B). Normalized fluorescence intensity (%FImax) of DPX/HPTS-containing polymersomes after 24 h incubation in dietary fiber-based hydrogels (C). Capture capacity of lyophilized and resuspended PS-b-PEO polymersomes in a bile salt- containing buffer (D). Polymer concentration: 1 mg/mL (A), 1.75 mg/mL (D); dye concentration: 50 µM (B) and 30 µM (C); buffer composition: isotonic phosphate buffer at pH 6.8 (A-C) containing cholate, deoxycholate, and taurocholate, 30 mM each (D); temperature 37°C. All results as means ± SD (n = 3). *p < 0.05, **p < 0.01, and ***p < 0.001.

In vivo evaluation

To assess the effect of orally administered PS-b-PEO polymersomes on systemic ammonia levels, a rat model of secondary biliary cirrhosis induced by BDL was selected. BDL rats show typical features of liver cirrhosis (hyperammonemia, pathologic alterations in bilirubin, liver enzymes, albumin, hepatomegaly) and HE (neurologic symptoms, brain edema), and have been successfully treated with gut-targeting ammonia-lowering strategies [264,309,310].

Firstly, PS-b-PEO polymersomes were orally applied in BDL rats at four weeks after surgery. Citric acid- containing polymersome dispersion (1.8 g/kg per day) or citric acid solution alone (0.96 g/kg per day) were gavaged twice daily for five days. At sacrifice, neither differences in blood ammonia levels nor in average weight gain per day were observed between the two treated groups (Figure 3.9). The PS-b- PEO polymersomes were likely destabilized in the distal intestine, where the chyme is dehydrated, similarly as in the dietary fiber hydrogels.

Figure 3.9. In vivo evaluation of PS-b-PEO polymersomes without laxatives. Study plan: 31 days after BDL surgery, two groups of cirrhotic rats (n = 4 per group) received polymersome dispersion in citric acid solution (twice daily 0.9 g polymer/kg) or citric acid solution alone (twice daily 0.48 g citric acid/kg) by gavage for five days. A group of healthy rats (n = 2) served as control. Blood ammonia levels in BDL rats at 31 days and at sacrifice at 36 days (both from saphenous vein) after BDL surgery and in healthy control rats (non-BDL control group n = 2, polymersome group n = 4, citric acid group n = 4) (A). Average weight gain per day in treated BDL rats (both n = 4) (B). All results as means ± SD. No significant differences were observed with respect to weight gain or blood ammonia levels between the two treated groups.

To mitigate the limited stability of PS-b-PEO polymersomes upon dehydration observed both in vitro and possibly occurring in vivo, the intestinal environment was hydrated by orally administered laxatives (Figure 3.10A). The use of laxative agents in chronic HE patients is common, as current practice guidelines recommend lactulose as first-line therapy in these patients [126]. The stool-hydrating properties of two orally administered laxatives, PEG (1 g/kg body weight) and sodium picosulfate (25 mg/kg, both administered once daily in the morning), were assessed in BDL rats. In contrast to PEG, sodium picosulfate significantly increased the stool water content compared with the water control over a period of 24 h (Figure 3.10B). While oral administration of sodium picosulfate increased the fecal water content, it led to a significantly decreased weight gain per day compared with BDL rats before laxative administration (Figure 3.10C). To investigate the efficacy of an oral PS-b-PEO polymersome application, BDL rats at 21 days after surgery were treated twice-daily (1 g/kg per day) for seven days. The plasmatic ammonia levels were significantly increased in BDL rats at day 28 compared with healthy control rats but the plasma levels of the polymersome and citric acid control group were not significantly different (Figure 3.10D). We hypothesize that the increase in stool hydration was not sufficient to create a non- destabilizing intestinal environment for the polymersomes as they are destabilized at an 80% (m/m) water content in dietary fiber hydrogels. 57

Figure 3.10. In vivo evaluation of PS-b-PEO polymersomes. Study plan: Three weeks after BDL surgery, two groups of cirrhotic rats (n = 16-17 per group) received sodium picosulfate (once daily 25 mg/kg) and polymersome dispersion in citric acid solution (twice daily 0.5 g polymer/kg) or citric acid solution alone (twice daily 0.24 g citric acid/kg) by gavage for one week. A group of age-matched healthy rats (n = 6) served as control and received citric acid solution (twice daily 0.24 g/kg) by gavage for one week (A). Stool water content in BDL rats gavaged with water (5 mL/kg), PEG(3350) (PEG, 1 g/kg), or picosulfate (25 mg/kg) (n = 3-19 stool samples) (B). Average weight gain per day in age-matched healthy control rats and BDL rats (non-BDL control group n = 6, BDL weeks 1-3 n = 37, BDL week 4: polymersome group n = 16, citric acid group n = 17) (C). Plasma ammonia levels in BDL rats at 21 days (saphenous vein) and at sacrifice at 28 days (cardiac puncture) after BDL surgery and in age-matched healthy controls (non-BDL control group n = 6, polymersome group n = 16, citric acid group n = 17) (D). All results as means ± SD. *p < 0.05, **p < 0.01, and ***p < 0.001.

The polymersome dose should have led to an appreciable decrease in ammonia based on their capture capacity. Assuming no destabilization in the GI tract, the seven-day polymersome treatment could cumulatively remove 2.8 mmol ammonia/kg body weight. In contrast, the cumulative removal of 0.6 mmol ammonia/kg by four LSPD sessions in the same animal model significantly decreased systemic ammonia levels [36]. Albeit potentially less selective to ammonia and with a lower ammonia uptake capacity, a three-day treatment with orally applied AST-120 microparticles at 4 g/kg with a cumulative removal of 0.3 mmol ammonia/kg led to a significant decrease in plasma ammonia in BDL rats [264]. 58

The present study shows that vesicles, which are stable under extreme in vitro conditions, may still be destabilized in the GI tract, especially as the chyme is dehydrated. Therefore, more in-depth studies are warranted to investigate the stability of vesicular carriers in the GI tract, especially when they are used to deliver sensitive drugs such as insulin [101].

3.4. Conclusion

The aim of this chapter was to develop an ammonia-sequestering formulation for ammonia detoxification via the oral route. Polymersomes made of PS-b-PEO were found to resist high concentrations of bile salts, extreme osmolality conditions, and digestive enzymes. In dietary fiber-based hydrogels, however, these polymersomes showed a fiber concentration-dependent reduction in ammonia uptake. As the stability of the transmembrane pH-gradient was limited in dietary fiber-based hydrogels, the animals in the subsequent in vivo study were supplemented with a laxative agent to hydrate the intestine. The efficacy of the polymersomes in terms of reducing plasmatic ammonia concentrations was evaluated in BDL rats, an established model of liver cirrhosis-associated hyperammonemia. Despite the significant increase in stool water content by the laxative agent, a reduction in plasmatic ammonia levels was not observed in the polymersome group compared with the buffer control group.

Chapter 4

4. Development of a PS-b-PEO polymersome-based ammonia assay

4.1. Introduction

The endogenous metabolite ammonia plays an important role in pH homeostasis and in amino acid and protein metabolism [282]. At pathologically elevated blood concentrations (hyperammonemia), however, it also accumulates in the brain and leads to neurotoxic effects (astrocyte swelling, impaired astrocyte buffering, neuronal dysfunction) [117,311]. The main metabolic pathway for ammonia detoxification is the UC in the liver [283]. The second ammonia-removing metabolic pathway relies on GS, which converts glutamate and ammonia to the transient ammonia sink glutamine, in the liver, muscles and the brain [283]. In patients suffering from liver disease (e.g., liver cirrhosis, acute liver failure) and inborn UCD, the UC insufficiently detoxifies ammonia and the GS pathway is rapidly saturated [282,283]. The resulting hyperammonemia is associated with low grade to severe neuropsychiatric symptoms (cognitive impairments, ataxia, lethargy, hepatic coma with a risk of death) [126]. In liver disease patients, this complication is referred to as hepatic encephalopathy (HE) and is highly prevalent among liver cirrhosis patients, affecting up to 80% of patients at the time of diagnosis [126]. The plasma ammonia levels both in UCD and HE were correlated with symptom severity and clinical outcome such that ammonia levels are routinely measured for diagnosis, disease staging, and prognosis [127,192,284].

In UCD patients, ammonia-lowering treatment options include hemodialysis in acute hyperammonemic crises and for maintenance therapy phenylbutyrate, which is metabolically coupled to glutamine and thus leads to the removal of this transient ammonia sink [282]. In HE patients, current practice guidelines recommend treatments which target the gut ammonia production by urease-producing bacteria, the main source of systemic ammonia [126]. While the first-line therapy lactulose leads to the expulsion of ammonia and modulates the microbiota due to its prebiotic effects, the second-line drug rifaximin decreases the bacterial ammonia production with its antibiotic action [282]. To evaluate the response to these treatments, the ammonia levels are generally determined in hyperammonemic patients [126,127].

The diagnostic gold standard for the measurement of ammonia in plasma, the preferred matrix for ammonia quantification, is an L-glutamate dehydrogenase (GLDH)-based enzymatic ammonia assay [312]. GLDH catalyzes the conversion of alpha-ketoglutarate and ammonia to L-glutamate [312]. As this conversion necessitates the oxidation of a nicotinamide adenine dinucleotide phosphate (NADPH) molecule to NADP+, the reaction can be followed by the decrease in absorbance at 340 nm upon the formation of the less absorbing NADP+ [312]. Several interferences such as hemolysis, icterus, and hyperlipidemia, which are commonly found in liver disease patients, were described for this test [312]. As this assay relies on an enzymatic reaction, it is prone to timing errors and thus not ideally suited for the processing of large amounts of samples (high-throughput).

The PocketChem BA (Arkray) is a strip-based, handheld device for ammonia quantification in whole blood and its derivatives [313]. Upon application on the strip, the blood is alkalized and ammonium is deprotonated. The thus formed ammonia is hydrophobic enough to diffuse across an ammonia-selective

60 poly(propylene) membrane [313]. Drawbacks of the PocketChem BA include its low throughput (3 min per measurement which can only be performed one at a time), its high price (approx. 15 USD per strip), and its constant underestimation of ammonia levels compared with the GLDH ammonia assay [313]. Furthermore, the device is currently not FDA-cleared for clinical use.

Two investigative ammonia assays were recently reported in the literature. To quantify ammonia in whole blood, a bicompartimental system with a selectively permeable membrane was developed [314]. The two compartments are separated by a membrane made of the cation-selective sulfonated tetrafluoroethylene polymer derivative Nafion® [314]. Upon addition of ammonia to the first compartment, which contains an acidic buffer, ammonia is ionized to ammonium and diffuses across the membrane [314]. In the second compartment, a solution of phenol and hypochlorite leads to the formation of the strongly absorbing indophenol in the presence of ammonia (Berthelot reaction) [314]. The ammonia- selective membrane is needed due to the low selectivity of hypochlorite towards ammonia in the presence of primary amines. The ammonia concentration is subsequently determined spectrophotometrically. Drawbacks of this approach include the long incubation time of the sample due to the slow diffusion of ammonium and potential selectivity issues in the presence of Nafion®-permeable small molecules with primary amines [314].

The second investigative ammonia assay is based on an electrochemical sensor device [315]. Similarly to the PocketChem, the sample is alkalized such that the thus formed gaseous ammonia can permeate across a polytetrafluoroethylene-based ammonia-selective membrane [315]. Ammonia then deprotonates polyaniline nanoparticles embedded in a silver electrode which leads to a quantifiable change in impedance (i.e., resistance to an electrical current upon application of a voltage) [315]. Negative points of this device include the long incubation time at elevated pH and the potentially low selectivity in the presence of other small, poly(tetrafluoroethylene)-permeable amines [315].

In this chapter, an ammonia assay based on transmembrane pH gradient PS-b-PEO polymersomes is proposed. As laid out in chapter 3, these polymersomes efficiently capture ammonia in their acidic core. Conceptually, the increase in core pH is a measure of the ammonia concentration outside the polymersome. Therefore, the incorporation of a pH-sensitive dye in the core could enable the quantification of the pH increase in the core and thus the extravesicular ammonia concentration (Figure 4.1).

Figure 4.1. Schematic depiction of the ammonia sensing mechanism of transmembrane pH gradient polymersomes.

4.2. Materials and Methods

HPTS fluorescence excitation spectra

The fluorescence excitation spectra of HPTS at 67 µM in isotonic citric acid solution 5.5 mM (pH 2.0- 6.0) or isotonic phosphate buffer 50 mM (pH 6.5-10.0) was determined at a fixed emission wavelength of 510 nm (Tecan Infinite 200 Pro) at room temperature.

PS-b-PEO polymer synthesis

PS-b-PEO(2000) synthesis was carried out by atom transfer radical polymerization (ATRP) [6]. PEO(2000) monomethyl ether (Sigma-Aldrich Chemie, Buchs, Switzerland) was converted to an ATRP macroinitiator by reacting it with 2-bromopropionyl bromide (Sigma-Aldrich Chemie) in dry tetrahydrofuran (THF, Acros Organics) and further used to polymerize styrene in bulk. Briefly, the ATRP macroinitiator (2.0 mmol) was loaded in a flame dried Schlenk flask, along with copper bromide (CuBr, 3.0 mmol Alfa Aesar) and 4,4’-dinoyl-2,2’-dipyridyl (2.64 mmol, TCI) as the catalyst and ligand, respectively. The Schlenk flask was evacuated and refilled with argon through several cycles to remove oxygen. In a separate flask, 11.5 mL of styrene (100 mmol, Sigma-Aldrich Chemie) was deoxygenated by bubbling argon through it for 0.5 h, and then loaded in the Schlenk flask. The mixture was then heated at 115 °C during 16 h and the brown product solution was dissolved in THF, filtered through a basic alumina column and precipitated twice in hexane. The precipitate was filtered and dried under vacuum. The feeding molar ratio of [monomer]/[initiator] was 50. The PS/PEO composition was determined in deuterated acetone by 1H nuclear magnetic resonance spectroscopy (Bruker AV-400, Billerica, MA) at room temperature. The polymer synthesis was carried out by Dr. Yinyin Bao (Laboratory of Drug Formulation and Delivery, ETH Zurich).

The PS-b-PEO(2000) polymers used in this chapter have PS fragments between 2800 and 4500, and are characterized in Tables 3.1 and 3.2.

Polymersome preparation

PS-b-PEO polymersomes were produced using an oil-in-water (o/w) emulsification method. 60 mg of polymer were dissolved in 100 µL dichloromethane or chloroform. The polymer-containing dichloromethane solution was added dropwise to mixture of 0.9 mL citric acid solution 5.5 mM at pH 5.7 (final pH 5.5, final citric acid concentration 5 mM, unless stated otherwise) at 300 mOsmol/kg and 0.1 mL HPTS 100 mM under sonication in an ice bath at amplitude 10 (3.1 mm sonotrode, Fisher Scientific™ Model 705 Sonic Dismembrator™, 700 W, 50/60 Hz, Fisher Scientific) for 2 min. The organic solvent was evaporated using a rotary evaporator for at least 5 minutes at 40°C at 70 kPa.

Purification and quantification

The fluorescent polymersomes were purified using MidiTrap G-25 columns (GE Healthcare Europe) to remove the free dye and exchange the external phase with sodium chloride-containing phosphate buffer 50 mM at pH 7.4 at 300 mOsmol/kg [289].

To determine the dye concentration in the polymersome dispersion, the polymersomes were diluted in N,N-dimethylformamide (final polymersome dispersion volume fraction 5%, v/v). The samples were subsequently centrifuged at 5000 x g for 2 min. The fluorescence emission of the supernatant was determined at 510 nm excited at 413 nm (Tecan Infinite 200 Pro) and compared to a HPTS standard curve in the same phosphate buffer / N,N-dimethylformamide mixture, and the HPTS concentration in the polymersome dispersion was calculated. The purified polymersomes were stored protected from light at 4°C and used in less than five days after purification except for the stability experiment (5 months of storage at 4°C).

Ammonia sensing

The samples, standards, or blank were co-incubated with polymersomes in isotonic phosphate buffer 50 mM at pH 7.4 at RT for 10 min (unless stated otherwise). The volume fraction of the sample, standard, or blank was 17% and 43% (both v/v) per well with a final HPTS concentration of 16 and 40 µM, respectively. The fluorescence emission at 510 nm was measured at an isosbestic (413 nm) and at a pH-dependent excitation wavelength (455 nm) to determine the fluorescence emission ratio (i.e., fluorescence emission at pH-dependent excitation wavelength normalized to the emission at the isosbestic excitation wavelength). The fluorescence emission ratio of the ammonia standards was corrected by subtracting the one of the blank (corrected fluorescence emission ratio). The accuracy was defined as the percentage of the mean of the measurements normalized to the nominal ammonia concentration. The precision was defined as coefficient of variation (i.e., standard deviation normalized to mean). The lower limit of quantification (LOQ) was determined based on equation 4.1 which relates to the current guidelines for bioanalytical method validation of the European Medicine’s Agency (EMA, EMEA/CHMP/EWP/192217/2009, legal effective date 2012/02/01)

휎 퐿푂푄 = (4.1) 푚 where σ stands for the standard deviation of the intercept and m the slope of the standard curve.

Selectivity

The ammonia concentration of a 100 µM ammonia solution (17% (v/v)) was determined in the presence of a mixture of drugs and metabolites (dopamine (TCI), levofloxacin (Sigma Aldrich Chemie), propranolol (Sigma Aldrich Chemie), glycine (Acros), alanine (Roth, Arlesheim, Switzerland), each at 500 µM), or di- (Sigma Aldrich Chemie, 100 or 200 µM) or trimethylamine (TCI, 100 µM) (17% (v/v)). The ammonia concentration was quantified as described above by comparison to an ammonia standard curve in isotonic phosphate buffer 50 mM at pH 7.4.

Healthy and BDL rat plasma samples

The animal experiments adhered to national guidelines and were approved by the local ethics committee of the Centre de Recherche du CHUM (CIPA, Montreal, QC, Canada). To induce secondary biliary cirrhosis, male Sprague‐Dawley rats (ca. 220 g) (Charles River Laboratories) underwent BDL. Weight- matched healthy male Sprague‐Dawley rats were used as control animals (n = 6). Approximately 5 mL of arterial blood were collected by cardiac puncture and centrifuged at 1500 x g at 4°C for 7 min after addition of heparin (Sandoz, final concentration 10 IU/mL). The samples were frozen and stored at - 80°C. After thawing on ice, the ammonia concentration was analyzed with the polymersome assay (compared with a standard curve in phosphate buffer at pH 7.4) as described above using a different plate reader (Tecan Spark®, Tecan), the enzymatic GLDH-based ammonia assay (AM1015, Randox Laboratories), and the PocketChem BA PA-4140 (Arkray).

In the GLDH enzymatic ammonia kit, the samples were measured based on the manufacturer’s instructions with a modified sample (20 µL), reagent buffer (200 µL), and the enzyme solution volume (2 µL) in a 96-well plate [36,106]. Based on an ammonium chloride standard curve (25-200 µM), the lowest concentration with acceptable accuracy (between 80-120%) and precision (CV below 20%) was determined (50 µM). The measurements below this value were set as 25 µM in accordance with the literature [36]. To measure the ammonia levels with the PocketChem BA, 20 µL of sample was applied to the strip and measured in mode F6 according to the manufacturer’s instructions.

To assess potential hemolysis, the absorbance at 340 nm of the healthy rat plasma diluted in trometamol 150 mM buffer at pH 8.6 (final plasma volume fraction 9%, v/v) was measured using a plate reader (Tecan Spark®), and the absorbance of a water control (i.e., water diluted in the same buffer) was subtracted.

Statistical analysis

The statistical calculations were carried out by SigmaPlot (version 13.0) and Microsoft Excel 2016 (determination of R2). Three or more groups were compared using one-way ANOVA (Holm-Sidak test) assuming a normal distribution of the data. For the comparison of two groups, a t-test was used. For the comparison of two independent samples (i.e., polymersome baseline fluorescence ratio, Figure 4.4B), an unpaired t-test was employed. For the comparison of two dependent samples (i.e., time dependence of fluorescence ratio, Figure 4.3B), a paired t-test was used. A p-value of <0.05 was deemed statistically significant.

4.3. Results and Discussion pH-dependent fluorescence of HPTS

The ammonia sensing mechanism of transmembrane pH gradient polymersomes depends on a pH- sensitive dye, which senses the pH increase upon ammonia influx into the vesicular core. The fluorescent dye HTPS was selected due to its pH-responsive excitation spectrum in the acidic range [304]. Its high hydrophilicity and three negative charges hinder the diffusion of HPTS through the polymersome membrane, trapping the dye in the polymersome core. In accordance with the literature [304], the wavelengths of 413 nm and 455 nm were selected as isosbestic (i.e., pH-independent) and pH-dependent excitation wavelengths, respectively, and the fluorescence was measured at an emission wavelength of 510 nm (Figure 4.2A). The fluorescence emission intensity ratio of the pH-sensitive and the isosbestic excitation wavelength showed a sigmoidal pH dependence with an inflexion point at pH 7.56 (Figure 4.2B). The normalization to the fluorescence emission intensity of the isosbestic excitation wavelength, which correlates with the dye concentration independently of pH, allowed for a correction for slight differences in dye concentration.

Figure 4.2. Fluorescence profile of HPTS at different pH values. Fluorescence emission intensity at 510 nm excited from 350 to 490 nm (A). Ratio of fluorescence emission intensity at 510 nm of an excitation wavelength at 455 nm (pH-dependent) to the one at 413 nm (isosbestic) at different pH values (B). Buffer solutions: citrate buffer 5.5 mM (pH 2.0-5.7), phosphate buffer 50 mM (pH 6.3 – 7.4), or tris 5 mM (pH 8.5, 11.1), all at 300 mOsmol/kg. Dye concentration 100 µM.

Ammonia sensing with HPTS-containing PS-b-PEO polymersomes

HPTS-containing PS-b-PEO polymersomes showed a linear ammonia sensing profile in the physiologically relevant range of 12.5 to 800 µM (coefficient of determination R2 = 0.9932, Figure 4.3A). In order to meet the criteria of the EMA on precision and accuracy (CV <15% (<20% at LOQ) and accuracy between 85-115%) in the range of 25 to 800 µM, the linear regression curve was split at 200 µM (Table 4.1).

Table 4.1. Accuracy and precision of standard curve in Figure 4.3A calculated based on a linear regression curve with the range 12.5-200 µM or 50-800 µM (n = 3-10).

12.5–200 µM 50–800 µM

Ammonia conc. (µM) Accuracy (%) Precision (%) Accuracy (%) Precision (%)

12.5 58.7 57.5 25 101.1 19.1 50 107.8 9.1 25.9 50.9 100 103.4 8.6 12.9 91.8 200 97.5 6.4 7.8 107.4 400 5.2 107.0 600 4.4 100.6 800 4.4 97.1

As the plasma ammonia cut-off is at 50 and 100 µM in adults and newborns, respectively, a high accuracy and precision at low concentrations are essential in the diagnosis of hyperammonemia [126,127,282]. In hyperammonemic crises, plasma ammonia levels often exceed 300 µM, and peak concentrations are correlated with disease severity and clinical outcome in acutely hyperammonemic HE and UCD patients [284,316,317]. A broad linear range is therefore highly advantageous for disease staging, outcome prediction, and response assessment to ammonia-lowering treatments in severe hyperammonemia. By splitting the linear regression curve, high accuracy and precision were achieved for the polymersome assay both below (non-hyperammonemic to moderately hyperammonemic) and above 200 µM (highly hyperammonemic) with acceptable precision and accuracy between 25 and 800 µM. The LOQ was at 39 and 27 µM for the 17% (v/v) and 43% (v/v) volume fraction set-up, respectively. The linear range and coefficient of determination of the polymersome assay were similar to the reported parameters for the diagnostic gold standard (GLDH-based enzymatic assay) apart from a slightly higher LOQ (Table 4.2). Furthermore, the polymersome assay exhibited a considerably broader linear range than the ones reported for the other tests.

Table 4.2. Comparison of the performance parameters of the polymersome assay with reported parameters for other ammonia assays.

Assay Linear range (µM) Coefficient of determination LOQ (µM) R2 Polymersome assay 25-800 0.9932a 27b 39c

GLDH-based enzymatic 7-940 0.9806 7 ammonia assay (Randox) [312] PocketChem BA [313] 7–286 0.977 7 Polyaniline nanoparticles 25–200 0.9868 36 [315] Bicompartimental well with 25–500 0.9757 25 ammonia-selective membrane [314] a12.5–800 µM (17% (v/v); n = 5); b12.5–200 µM (43% (v/v); n = 3); c12.5–200 µM (17% (v/v); n = 5)

To assess the impact of potential timing errors on the read-out, the change in response was evaluated from 2.5 to 15 min after adding the polymersomes to the ammonia-containing buffer. There was no statistically significant change in fluorescence emission ratio between 10 and 15 min (Figure 4.3B). Therefore, timing errors are presumably less relevant in this time frame than with the GLDH-based ammonia assay which relies on an enzymatic reaction. A low time-dependency would allow for measuring large amounts of samples and controls (standards, calibrators, blanks) simultaneously. Furthermore, the time needed to reach the equilibrium in the polymersome assay corresponds to the incubation time in the GLDH-based ammonia assay. A 10-min incubation time at room temperature should therefore not be associated with artefacts (e.g., ammoniagenic degradation reactions [302]). In contrast, an investigative ammonia assays based on an ammonia-selective membrane and the Berthelot reaction necessitated an incubation time of at least 30 min [314].

Figure 4.3. Ammonia sensing with HPTS-containing PS-b-PEO polymersomes. Blank-corrected fluorescence emission ratio at different ammonia concentrations (A). Fluorescence emission ratio normalized to 2.5 min at different ammonia concentrations over 15 min (B). Inner phase: isotonic citrate buffer 5 mM (A) or 1 mM (B) at pH 5.5; outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS 67 concentration: 16 µM. Corrected fluorescence emission ratio (FR): FR of blank subtracted from FR at given ammonia concentration. Normalized FR: FR at given time point normalized to FR at 2.5 min. All results as mean ± SD (n = 3). No statistically significant differences were observed after 10 min for a given ammonia concentration (B).

As the stability of the transmembrane pH gradient is crucial for shelf-life, the ammonia sensing capacity of PS-b-PEO polymersomes stored in an outer phase at pH 7.4 and 4°C was assessed after five months. The polymersomes were still capable of sensing ammonia in a physiologically relevant range in a linear fashion (R2 = 0.9932, Figure 4.4A). However, the baseline fluorescence emission ratio was significantly higher after five months, indicating a slight reduction in transmembrane pH gradient over time. Alternatively to a storage at physiological pH, the polymersomes could instead be stored in acidic buffer to prevent proton exchange between the inner and outer media. Provided that the buffering capacity of the outer phase is low, the transmembrane pH gradient could be created in situ upon addition of the analyte (e.g., plasma, ammonia-containing buffer).

Figure 4.4. Stability of HTPS-containing PS-b-PEO polymersomes stored at pH 7.4 and 4°C for five months. Blank-corrected fluorescence emission ratio at different ammonia concentrations (A). Fluorescence emission ratio of blank at baseline and after five months of storage at pH 7.4 and 4°C (B). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 16 µM. Corrected fluorescence emission ratio (FR): FR of blank subtracted from FR at given ammonia concentration. All results as mean ± SD (n = 3). *p < 0.05.

Selectivity

To assess their selectivity towards ammonia, the polymersomes were co-incubated with ammonia and physiologically relevant endogenous metabolites and drugs. The selected weakly basic molecules exhibited a protonable nitrogen moiety, a logP between -3.2 and 3.0, and a molecular weight below 400 g/mol (Table 4.2). These properties render the molecules amenable to an uptake by transmembrane pH-gradient vesicles [318]. Indeed, several of the investigated molecules were reported to be taken up by transmembrane pH-gradient liposomes [106,318–322].

Table 4.2. Relevant physicochemical properties of the investigated molecules (information retrieved from PubChem, National Center for Biotechnology Information).

Molecule Molecular weight pKa of corresponding protonated nitrogen logP Zwitterion at (g/mol) physiological pH Ammonia 17 9.25 -0.7 Dimethylamine 45 10.73 -0.2 Trimethylamine 59 9.80 0.3 Alanine 89 2.34 (carboxylic acid), 9.60 (amine) -3.0 x Glycine 75 2.34 (carboxylic acid), 9.60 (amine) -3.2 x Dopamine 153 8.81 (phenol), 10.90 (amine), 13.68 (phenol) -1.0 Propranolol 259 9.42 (amine) 3.0 Levofloxacin 361 6.24 (carboxylic acid), 8.74 (piperazinyl) -0.4 x

The di- and tri-methylated analogs of ammonia (DMA, and TMA, respectively) were chosen because of their structural similarity to ammonia and their physiological relevance [323,324]. Even at concentrations exceeding physiological serum concentrations >25-fold [323,324], DMA and TMA did not significantly influence the measured ammonia concentration (Figure 4.5). Furthermore, there was no statistically significant overestimation at pathologically elevated serum DMA levels which can be observed in hemodialysis patients (up to 200 µM [325]). Moreover, a mixture of drugs and metabolites (levofloxacin, propranolol, glycine, alanine, dopamine, each in five-fold molar excess over ammonia) did not interfere with the ammonia assay. The concentrations of the drugs and metabolites ranged from physiologically relevant (glycine, alanine) to highly elevated levels (levofloxacin, propranolol, dopamine) [326–330]. In contrast to transmembrane pH gradient liposomes [106,318–321], PS-b-PEO polymersomes selectively sequestered ammonia in the presence of weakly basic molecules. In addition, the destabilization of liposomes reported for propranolol was not observed with the investigated polymersomes [106]. We hypothesize that the highly hydrophobic, rigid membrane of PS-b-PEO polymersomes hindered the diffusion of bulkier and more hydrophilic substrates than ammonia.

Figure 4.5. Selectivity of PS-b-PEO polymersomes. HTPS-containing transmembrane pH gradient polymersomes co-incubated with ammonia and DMA (100-200 µM), TMA (100 µM), or a mixture (Mix) of drugs and metabolites (levofloxacin, propranolol, glycine, alanine, dopamine, each at 500 µM). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 16 µM. All results as mean ± SD (n = 3). No statistically significant differences were observed between the groups.

Ammonia sensing in plasma of healthy rats

To evaluate the performance of the polymersome ammonia assay in plasma, the ammonia concentration of plasma samples from healthy rats was determined by the polymersome assay, the GLDH-based enzymatic assay (diagnostic gold standard), and the PocketChem BA. In four of the six investigated rats, the polymersome assay yielded similar to slightly increased ammonia levels compared with the comparator tests (Figure 4.6A). In rats E and F, on the other hand, the ammonia levels obtained by the enzymatic kit were more than two-times higher than the polymersome assay and PocketChem BA. Generally, ammonia samples from healthy rats do not exceed 80 µM [36,264]. As observed visually and by the increased baseline absorbance at 340 nm (i.e., absorbance from released hemoglobin [331], Figure 4.6B), the samples E and F were slightly hemolyzed. Hemolysis is a reported interference for the enzymatic ammonia assay which may have resulted in an overestimation of the ammonia levels [312]. In contrast, the polymersome assay and the PocketChem BA did not show strongly increased ammonia levels, which highlights their usefulness in slightly hemolyzed samples.

Figure 4.6. Ammonia quantification in healthy rat plasma. Arterial plasma samples (cardiac puncture) from six healthy male rats measured with the polymersome assay (white), the GLDH-based ammonia assay by Randox (gray), and the PocketChem BA (black) (A) (n = 3-10). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 40 µM. Plasma volume fraction in well 43% (v/v). Corrected absorbance at 340 nm in the same six rats (B) (n = 3-10). All results as mean ± SD (n = 3). *p < 0.05, **p < 0.01, and ***p < 0.001.

Ammonia sensing in plasma of BDL rats

The performance of the polymersome assay was further evaluated in the plasma of BDL rats. This animal model exhibits several hallmarks of liver cirrhosis (hyperammonemia, icterus, muscle wasting) [36,310]. The plasma samples from these animals exhibit typical pathological changes in clinical 70 laboratory parameters of liver cirrhosis (hyperbilirubinemia, hyperlipidemia) which are known interferences of ammonia measurements with the GLDH-based ammonia assay [36,312]. With the same set-up as for the measurements in the healthy rats, the polymersome assay showed significantly higher ammonia levels than the enzymatic assay (overestimation of 38.3 ± 14.3%) in seven of the nine tested animals (Figure 4.7A). This overestimation is potentially related to the hyperbilirubinemia (i.e., the accumulation of bilirubin in the blood, icterus), a typical feature of BDL plasma. Bilirubin exhibits a broad fluorescence emission peak around 525 nm when excited at 455 nm [332]. Therefore, exciting HPTS at the pH-dependent excitation wavelength also excites bilirubin, which presumably leads to an increased fluorescence intensity at the employed emission wavelength 510 nm and thus an overestimation of the ammonia concentration. Furthermore, bilirubin slightly interferes with the isosbestic excitation wavelength (413 nm) with regard to its absorbance spectrum at pH 7.4 (peak at 440 nm, Figure 4.7B). In consequence, a slightly lower emission intensity was observed at the isosbestic excitation wavelength, further contributing to the overestimation. The polymersome and the GLDH-based assays yielded significantly higher ammonia levels than the PocketChem BA in eight and nine of the nine tested animals, respectively. This finding was in accordance to the reported constant proportional underestimation of ammonia concentrations by the PocketChem BA compared with the GLDH assay [313].

Figure 4.7. Ammonia quantification in BDL rat plasma (high volume fraction set-up). Arterial plasma samples (cardiac puncture) from nine male BDL rats measured with the polymersome assay (white), the GLDH-based ammonia assay by Randox (gray), and the PocketChem BA (black) (A) (n = 3-4). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 40 µM. Plasma volume fraction in well 43% (v/v). All results as mean ± SD. *p < 0.05, **p < 0.01, and ***p < 0.001. Absorbance spectrum of bilirubin at pH 7.4 (B).

To decrease the interference of bilirubin, the volume fraction of plasma per well was decreased from 43% to 17% (v/v). With this low volume fraction set-up, the polymersome assay yielded higher ammonia levels than the enzymatic kit in only one of the five BDL plasma samples (Figure 4.8A). In accordance with the literature [313], the significantly lower ammonia levels determined by the PocketChem BA persisted compared with both the polymersome and the GLDH assays.

Standard curves in phosphate buffer yielded a less steep slope and slightly higher LOQ with the low compared with the high volume fraction set-up (Table 4.2, Figure 4.8B), probably due to the lower final ammonia concentration per well. Nonetheless, the LOQ remained relatively low such that a lower plasma volume fraction could be beneficial in hyperbilirubinemic samples without impairing the performance at low ammonia concentrations.

Figure 4.8. Ammonia quantification in BDL rat plasma (low volume fraction set-up). Arterial plasma samples (cardiac puncture) from five male BDL rats measured with the polymersome assay (white), the GLDH-based ammonia assay by Randox (gray), and the PocketChem BA (black) (A). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 16 µM. Plasma volume fraction in well 17% (v/v). *p < 0.05, **p < 0.01, and ***p < 0.001. Blank-corrected fluorescence emission ratio at different ammonia concentrations in the low (17%, v/v) and high (43%, v/v) volume fraction set-up (B). Inner phase: isotonic citrate buffer 5 mM at pH 5.5, outer phase: isotonic phosphate buffer 50 mM at pH 7.4. HPTS concentration: 16 µM (17%, v/v) and 40 µM (43%, v/v). All results as mean ± SD (n = 3-5).

3.4 Conclusion

The aim of this chapter was to develop a PS-b-PEO polymersome-based ammonia assay. Encapsulating the pH-sensitive fluorescent dye HPTS in the polymersome core allowed the quantification of ammonia in solution with high selectivity and sensitivity and low time-dependence after 10 min. Moreover, the broad linear range promises to reliably assess both low and high ammonia quantifications. The polymersome assay performed similarly as the comparator assays in plasma of healthy and BDL rats.

Chapter 5

5. General discussion and outlook

As laid out in chapter 1, the naturally occurring metabolite ammonia is not sufficiently detoxified and accumulates in the blood of a high number of liver disease patients [282]. The resulting hyperammonemia is associated with potentially lethal neuropsychiatric symptoms [282]. The existing treatments often provide insufficient therapeutic benefit, and can be associated with adverse reactions [126]. Especially in HE, the main ammonia-lowering treatments targeting the gut, lactulose and rifaximin, inadequately control hyperammonemia in a large part of the patient population [126]. Furthermore, lactulose led to compliance-impairing adverse reactions (e.g., diarrhea) and the prolonged intake of the antibiotic rifaximin may entail the risk of antibiotic resistance [282].

In chapter 3, we aimed at developing a novel ammonia-sequestering oral treatment for HE based on transmembrane pH-gradient polymersomes. A small library of PS-b-PEO(2000) polymersomes with high ammonia capture in simulated GI fluids (e.g., high bile salt concentrations, hypo- and hyperosmolar conditions, digestive enzymes) was identified. The polymersomes were further incubated in dietary fiber- based hydrogels to simulate the colonic environment where the feces thicken upon water resorption. In this matrix, the polymersomes lost their ammonia-sequestering capacity in a fiber concentration- dependent manner. Further experiments revealed that this result was likely related to a loss of the transmembrane pH gradient. In the subsequent in vivo evaluation of the polymersomes, the BDL rats were supplemented with a laxative to increase the water content of the stools. Nevertheless, the polymersome treatment did not lead to an appreciable decrease in plasma ammonia compared with the control treatment. Seeing that a significant decrease in capture capacity was observed at the water content corresponding to stools under laxative treatment (80%, m/m), the hydration of the stools was probably not strong enough to stabilize the vesicles. In the future, a more in-depth investigation of this destabilization process is warranted. We hypothesize that highly hyperosmolar conditions are created in the hydrogel matrix due to the binding of free water to the polysaccharide-based hydrogel matrix. Unfortunately, the exact conditions in the hydrogel matrix remain poorly understood, mainly due to the difficulty of measuring the osmolality of highly concentrated hydrogels with conventional osmometers. Potentially, extrapolations based on the osmolality of low fiber concentrations could be envisaged.

An important piece of information on the susceptibility of PS-b-PEO polymersomes to osmotic pressure gradients is the water permeability of the membrane. As adaptive behavior to osmotic stress (i.e., shrinking and swelling) may transiently destabilize the vesicular membrane and lead to a loss of the pH gradient [36], a low water permeability would be favorable to resist extreme osmolality conditions. Even though direct evidence on the water permeability of the investigated PS-b-PEO polymersomes is lacking, a slight loss of transmembrane pH gradient was observed after storage of fluorescent PS-b- PEO polymersomes at 4°C for five months. While this result may also be explained by the loss of HPTS, a leakage of the much smaller and less charged hydronium ion is more likely. Therefore, the membrane of PS-b-PEO polymersomes is slightly permeable to small hydrophilic molecules such as hydronium ions under isotonic conditions, suggesting a certain water permeability at baseline as reported for PS [305]. To assess the permeability to water, changes in vesicle size or morphology in media with different 73 osmolalities could be assessed with microscopy techniques [24]. PS-b-PEO polymersomes were recently reported to show low water permeability in media with low amounts of organic solvent [24]. However, the diblock copolymer PS(14000)-b-PEO(2000) employed in this study had a considerably longer PS fragment than the ones investigated in chapter 3, and the polymersome preparation process (nanoprecipitation) differed such that the membrane packing and permeability might differ.

Moreover, mechanical stress in the hydrogel matrix could contribute to the destabilization of the membrane. The large size of the vesicles make them particularly prone to mechanical stress, and the highly rigid membrane may not allow enough flexibility for membrane rearrangement upon mechanical stress [306]. As we were not able to reduce the size of PS-b-PEO polymersomes, the resistance of the membrane to mechanical stress could be investigated for instance by micropipette aspiration or atomic force microscopy experiments [306,8,333]. As a comparator, polymersomes made of low Tg diblock copolymers such as PBD-b-PEO could be employed.

Furthermore, the ammonia capture capacity could be evaluated in more biorelevant media, such as caecum fluid and stools. Rat stools could be lyophilized and the lost water fraction could be added in form of polymersome-containing buffer such that the most distal part of the colon could be simulated. By adding different amounts of water, the conditions in the laxative-supplemented rats could be approximated. Investigating the stability over 4-12 h will give further insights of stability over time, which is of particular interest with regard to the accelerated transit time upon laxative administration [334]. High and variable ammonia concentrations in stools may complicate such experiments, though.

More fundamental studies on the presented PS-b-PEO polymersome library would also be of interest. Ammonia uptake experiments in a bile salt-containing medium showed, for instance, that PS-b-PEO copolymers with PS fragments <2000 did not take up ammonia to an appreciable degree. Therefore, stable polymersomes were not formed by sonication-enhanced emulsification. Other macromolecular assemblies (e.g., micelles, nanoparticles, polymersomes) could have formed. Micelles composed of PS(100000)-b-PEO(10000) were reported in water/(N,N)-dimethylformamide mixtures with low water content (4.5wt%) [335]. The macromolecular assembly could be characterized with scanning or transmission electron microscopy. Moreover, hydrophilic and hydrophobic fluorescent dyes could be incorporated and analyzed with fluorescence microscopy in order to investigate the potential formation of vesicular structures with a fluorescent core and membrane, respectively, especially in the absence of high bile salt concentrations.

In the light of the negative in vivo results, several modifications to the polymersome treatment could be applied. Longer PS fragments could be used in order to stabilize the polymersomes. As they are more hydrophobic and may thus be associated with a higher risk of aggregation during the preparation process, longer PEO fragments could be used to keep the PS/PEO ratio constant. Alternatively, different polymersome-forming polymers may be investigated. With regard to the lower water permeability of aliphatic polymers compared with PS [305], polymersomes made of aliphatic hydrophobic fragments (e.g., PEE, see chapter 2) could be tested under extreme osmolality conditions and in dietary fiber hydrogels. However, the high membrane fluidity of these polymersomes (Tg of PEE = -45 °C [336]) may make them susceptible to destabilization by bile salts.

Other approaches to selectively capture ammonia in the gut may also be investigated. Zeolite microparticles, for instance, bind both ammonia and a variety of cations such as ammonium [337,338]. Therefore, they could be coated with a hydrophobic ammonia- (e.g., poly(propylene)) or hydrophilic ammonium-selective coatings (e.g., Nafion®). For the latter approach, achieving selectivity over potassium with its identical charge and similar ionic radius may be challenging, though.

As laid out in chapter 4, the quantification of plasma ammonia is of paramount importance in disease staging, prognosis, and the response assessment to ammonia-lowering treatments in hyperammonemia. Current ammonia assays have significant drawbacks such as interferences and low upper levels of quantification. We therefore aimed at developing a novel ammonia assay using the transmembrane pH gradient PS-b-PEO polymersomes investigated in chapter 3. The polymersome- based ammonia assay allowed the quantification of ammonia in a pathophysiologically relevant range in buffers, and performed similarly to the comparators in plasma from healthy and BDL (i.e., cirrhotic and hyperammonemic) rats.

The further development of the PS-b-PEO polymersome ammonia assay could be divided in two phases: an explorative and a validation phase. In the explorative phase, each component of the assay should be optimized. The intravesicular dye HPTS, for instance, could be replaced with a dye less prone to interference by plasma constituents or contaminants (e.g., hemoglobin, bilirubin). A dye in the near- infrared region would be most suitable, as naturally occurring metabolites generally do not interfere in this range of the electromagnetic spectrum. A near-infrared dye based polymersome assay would further enable ammonia sensing in other matrices, notably whole blood, which could enable point-of-care ammonia testing. A promising near-infrared dye with a pH-dependent fluorescence emission in the near- infrared spectrum with an inflexion point at pH 3.7 was identified in preliminary experiments (Figure 5.1A). Upon encapsulation in PS-b-PEO polymersomes, a linear response to ammonia in a physiologically relevant range was observed (Figure 5.1B). However, the low fluorescence emission intensity and the missing isosbestic wavelength led to high variability and a comparably low coefficient of determination (R2 = 0.9353). Therefore, higher concentrations of dye would need to be encapsulated to improve the signal-to-noise ratio. Alternatively, another dye with higher quantum yield and a pH- independent excitation or emission wavelength could be selected. Another possibility to enable the quantification of the pH-dependent dye would be its ratiometric coupling of a pH-independent dye [339].

Figure 5.1. pH dependence of IRDye 680RD (LI-COR Biosciences, Bad Homburg, Germany) and ammonia sensing with PS-b-PEO polymersomes encapsulating IRDye 680RD. Fluorescence emission intensity at 696 nm excited at 666 nm at different pH values (A). Isotonic citrate solution 5.5 mM. Dye concentration 10 µM. Fluorescence emission intensity at 696 nm excited at 666 nm of IRDye 680RD- containing polymersomes at different ammonia concentrations (B). Polymer concentration 0.72 mg/mL. Inner phase of polymersomes: IRDye 680RD 40 µM in citric acid solution 20 mM at pH of 3.0. Results as mean ± SD.

Furthermore, the PS fragment length may be varied to optimize stability and uptake kinetics. Longer PS fragments promise to enhance stability and thus increase shelf life because of their higher membrane hydrophobicity and therefore presumably lower proton permeability. However, the ammonia uptake kinetics may be decelerated because of the longer diffusion distance. In addition, the higher hydrophobicity of the PS fragment may lead to more aggregation in the preparation method. As the diffusion of more hydrophilic or bulkier substrates is presumably hindered, a more hydrophobic membrane could further increase the selectivity towards ammonia. Based on the selectivity experiments in chapter 4, an improvement of this parameter is probably not needed, though.

Once the best polymer candidate is identified, the polymer synthesis, polymersome preparation, and polymersome purification processes will be scaled up to an industrial scale. As sonication-based protocols can only be scaled up with difficulty, another method to emulsify the polymer-containing organic phase (e.g., using a homogenizer) and the dye-containing water phase needs to be established [340]. Column-based purification methods are also difficult to scale up such that other methods such as cross-flow filtration and centrifugation should be investigated [341]. Crucially, these methods need to remove large amounts of free dye and potentially exchange the outer phase while preserving the polymersome integrity.

A thorough optimization of the volume fractions of the assay components (i.e., polymersome dispersion, ammonia-containing solution, and buffer) is of high importance. Indeed, the volume fraction of the standards influenced the sensitivity of the assay as it determines the final ammonia concentration in the well. For plasma samples in BDL rats, the composition had an influence on the measured ammonia levels, probably due to an interference by bilirubin. Therefore, the ideal volume fractions of the ammonia- containing standards, plasma samples, and the best polymersome concentration need to be thoroughly investigated in biorelevant media, notably rat and human plasma samples. For instance, potentially interfering substances such as bilirubin and hemoglobin could be added at pathophysiologically relevant concentrations to determine the polymersome concentration at which the interfering substance is tolerated.

Once the assay is optimized, a validation based on the current guidelines for bioanalytical method validation by the EMA and FDA should be carried out in biorelevant media. The defining parameters of the assay (e.g., lower limits of detection and quantification, linear range, robustness, etc.) should be determined and stability experiments at different temperatures and outer phases should be conducted to establish the shelf-life of the formulation.

In conclusion, an oral formulation of PS-b-PEO polymersomes was developed for ammonia sequestration in the intestine. While resistant to simulated GI fluids in vitro, the PS-b-PEO polymersomes did not lower plasmatic ammonia levels in hyperammonemic rats. Presumably, the vesicles were destabilized in the thickening feces in the distal GI tract. However, fluorescent PS-b-PEO polymersomes were capable of quantifying ammonia with high sensitivity and selectivity in biorelevant media. In plasma of healthy and BDL rats, the polymersome assay performed similarly as the comparators.

6. List of abbreviations

BDL: bile duct ligation; BCAA: branched-chain amino acid; CoA: coenzyme A; DPX: p-xylene-bis(N- pyridinium bromide); ELAD: extracorporeal liver assist device; FDA: Food and Drug Administration; FR: fluorescence emission ratio; GI: gastrointestinal; GS: glutamine synthase; HA: hyperammonemia; HE: hepatic encephalopathy; HPTS: 8-hydroxypyrene-1,3,6-trisulfonate; IP: intraperitoneal; IV: intravenous; MARS: molecular adsorbent recirculation system; MELD: model end stage liver disease; NADPH/NADP+: nicotinamide adenine dinucleotide phosphate; OTC: ornithine transcarbamylase; PS: poly(styrene); PEG(3350): poly(ethylene glycol) 3350; PEO: poly(ethylene oxide); PBD: poly(butadiene); SC: sodium cholate; SDC: sodium deoxycholate; SEM: scanning electron microscopy;

STC: sodium taurocholate; THF: tetrahydrofuran; Tg: glass transition temperature; Tm: phase transition temperatures; UC: urea cycle; UCD: UC disorders.

7. Curriculum Vitae

Personal information

Simon Matoori

Schweighofstrasse 335

8055 Zurich

Switzerland

OrcID: 0000-0002-1559-0950

Born in Zurich on November 30th, 1988

Swiss citizen

Marital status: unmarried, no children

EDUCATION

2014 – 2018 Ph.D. studies in Drug Formulation and Delivery under the supervision of my Ph.D. advisor Prof. Dr. Jean-Christophe Leroux, Institute of Pharmaceutical Sciences (IPW), Department of Chemistry and Applied Biosciences (DCHAB), ETH Zurich; supported by the doctoral scholarship of the Swiss Chemical Industry

• 04/2017-05/2017, 11/2017-12/2017, 04/2018-05/2018, 09/2018-10/2018 In vivo studies for Ph.D. thesis in the Laboratory of Prof. Dr. Christopher Rose, CRCHUM, Université de Montréal (Canada)

2013 – 2014 Master of Science degree in Medicinal and Industrial Pharmaceutical Sciences, IPW, DCHAB, ETH Zurich

2011 – 2013 Master of Science degree in Pharmaceutical Sciences and Federal Diploma for Pharmacists, IPW, DCHAB, ETH Zurich; supported by the ETH Excellence Scholarship & Opportunity Program; graduated with distinction

2008 – 2012 Bachelor of Science degree in Pharmaceutical Sciences, IPW, DCHAB, ETH Zurich; graduated with distinction

2001 – 2007 Gymnasium and Matura Certificate, Kantonsschule Wiedikon (Zurich)

1995 – 2001 Primary School, Schulhaus Friesenberg (Zurich)

EMPLOYMENT HISTORY

09/2014 – 12/2018 Ph.D. studies in Drug Formulation and Delivery (Ph.D. advisor Prof. Dr. Jean-Christophe Leroux), IPW, DCHAB, ETH Zurich; supported by the doctoral scholarship of the Swiss Chemical Industry (100%)

07/2014 – 08/2014 Internship at the pharma consulting company Dr. J. Fröhlich Beratungen GmbH (100%)

09/2013 – 02/2014 Research project SIgA-CD71 Complex Formation - Signaling, Transcription, Translation in the research group of Dr. Nadine Cerf- Bensussan, Laboratory of Intestinal Immunology, INSERM UMR1163 (previously U989), Institut Imagine, Hôpital Necker and Université Paris Descartes - Sorbonne Paris Cité, Paris, France

11/2012 – 08/2013 Assisting Pharmacist (Praktische Assistenzzeit) at Apotheke im Zollikerberg (Zollikerberg, Zurich) under the supervision of Daniela Koller, as part of the MSc Pharmaceutical Sciences

02/2012 – 08/2012 Master’s thesis Polymer-Conjugated Prolyl Endopeptidases – Stability, Mucoadhesion, Labeling under the supervision of Dr. Gregor Fuhrmann and Prof. Dr. Jean-Christophe Leroux, IPW, DCHAB, ETH Zurich. Awarded with Amedis Award for Best Master’s Thesis in Pharmaceutical Sciences at ETH Zurich and University of Basel

11/2011 – 12/2011 Research project Michaelis-Menten Kinetics of Polymer-Conjugated Prolyl Endopeptidases under the supervision of Dr. Gregor Fuhrmann and Prof. Dr. Jean-Christophe Leroux, IPW, DCHAB, ETH Zurich

INSTITUTIONAL RESPONSIBILITIES

01/2016 – 09/2016 Member of the committee for the renewal of the MSc programs (Studiengangsrevision) in Pharmaceutical Sciences and Medicinal Industrial Pharmaceutical Sciences MIPS, chaired by Prof. Dr. Cornelia Halin-Winter and PD Dr. Vivianne Otto

SUPERVISION OF STUDENTS 80

11/2016 – 07/2017 Supervision of MSc Pharmaceutical Sciences student Julia Müller (research project and master’s thesis)

11/2015 – 07/2016 Co-supervision of MSc Pharmaceutical Sciences student Olha Voznyuk with Prof. Dr. Georgios Sotiriou (research project and master’s thesis)

TEACHING ACTIVITIES

09/2017 Examinator in Federal Exam for Pharmacists, part Arzneimittelherstellung in kleinen Mengen (Christa Meier and Dr. Vivianne Otto), 4 half-days

09/2015, 09/2016 Assistant in Federal Exam for Pharmacists, part Arzneimittelherstellung in kleinen Mengen (Christa Meier and Dr. Vivianne Otto), 4 half-days

01/2015, 01/2016, Assistant in practical course Arzneimittelherstellung in kleinen

01/2017 Mengen (Dr. Johannes M. Fröhlich), 9 half-days per year

10/2014-11/2014, Assistant in practical course Allgemeine Chemie Praktikum für

10/2015-11/2015, Pharmazeuten und Biologen (Dr. Reinhard Kissner), 15 half-days per

01/2017 year

ACTIVE MEMBERSHIPS IN SCIENTIFIC SOCIETIES

Since 2017 Student member of the European Society of Radiology

PRIZES, AWARDS AND FELLOWSHIPS

2018 SNF Early.Mobility Fellowship

2018 First Prize Poster Award at Swiss Pharma Science Day 2018

2015 Doctoral Scholarship Award of the Swiss Chemical Industry

2012 Amedis Award for Best Master’s Thesis in Pharmaceutical Sciences at ETH Zurich and University of Basel

2011 ETH Excellence Scholarship & Opportunity Program Award (scholarship for MSc in Pharmaceutical Sciences)

PERSONAL SKILLS

Language skills German First language

English Business fluent in speech, writing and reading (CAE)

French Business fluent in speech, writing and reading

Italian Fluent in speech, writing and reading (CELI C1)

7. Scientific contributions

PUBLICATIONS IN PEER-REVIEWED SCIENTIFIC JOURNALS

• Giacalone G, Matoori S, Agostoni V, Forster V, Kabbaj M, Eggenschwiler S, Lussi M, De Gottardi A, Zamboni N, Leroux JC. Liposome-supported peritoneal dialysis in the treatment of severe hyperammonemia: An investigation on potential interactions. J Control Release. 2018; 278:57- 65. o https://www.ncbi.nlm.nih.gov/pubmed/?term=giacalone+matoori

• Moroz E*, Matoori S*, Leroux JC. Oral delivery of macromolecular drugs: Where we are after almost 100 years of attempts. Advanced Drug Delivery Reviews 2016; 101:108-21. o https://www.ncbi.nlm.nih.gov/pubmed/26826437

• Matoori S and Leroux JC. Recent advances in the treatment of hyperammonemia. Advanced Drug Delivery Reviews 2015; 90:55-68. o https://www.ncbi.nlm.nih.gov/pubmed/25895618

• Fuhrmann G, Grotzky A, Lukić R, Matoori S, Luciani P, Yu H, Zhang B, Walde P, Schlüter AD, Gauthier MA, Leroux JC. Sustained gastrointestinal activity of dendronized polymer-enzyme conjugates. Nature Chemistry 2013; 5:582-9. o https://www.ncbi.nlm.nih.gov/pubmed/23787748

• Matoori S, Fuhrmann G, Leroux JC. Celiac Disease: a Challenging Disease for Pharmaceutical Scientists, Pharmaceutical Research 2013; 30(3):619-26. o https://www.ncbi.nlm.nih.gov/pubmed/23229860

*Co-first author

PROFESSIONAL ARTICLES

• Matoori S, Fuhrmann, G, Schulz, JD, Leroux, JC. Gluten binden und spalten – zwei neue adjuvante Therapieansätze für die Zöliakie. Swiss Medical Forum 2012; 12(37):716-717. o https://medicalforum.ch/fr/resource/jf/journal/file/view/article/fms.2012.01252/fms-01252.pdf/

PATENTS AND LICENSES

Matoori S (contribution 50%, see confirmation by ETH Transfer), Wuerthinger O, Leroux JC (title not disclosed), US62557256

Matoori S (contribution 47.5%, see confirmation by ETH Transfer), Schmidt A, Leroux JC (title not disclosed), EP16184371.9

INTERNATIONAL CONFERENCES

• NanoDDS 2018, Sept 21-23 2018, Poster presentation “Development of a novel polymersome-based ammonia assay”, Simon Matoori, Yinyin Bao, Rafael Ochoa-Sanchez, Mélanie Tremblay, Christopher F. Rose, Jean-Christophe Leroux

8. Acknowledgments

First and foremost, I thank Prof. Dr. Jean-Christophe Leroux for being such an outstanding mentor. His scientific brilliance, passion for science, and impressive work ethics were inspirational, and his support was tireless. I am extremely grateful that he gave me the opportunity to work so closely with him. It was a pulsatile hypo- and hyperammonemic journey that I would not have wanted to make with anybody but him.

I am very thankful that Prof. Dr. Peter Walde accepted to be the co-examiner of this thesis. His lectures on vesicles were enlightening and every scientific discussion with him highly fruitful.

I am deeply grateful to Dr. Yinyin Bao for his tireless polymer syntheses. His diligence, scientific knowledge, and positive attitude made him a great collaborator and friend. I also thank Aaron Schmidt for his support and kindness, and for interesting discussions about polymersomes.

Special thanks go to the research group of Prof. Dr. Christopher Rose for supporting our animal studies with great enthusiasm, especially Rafael Ochoa-Sanchez, Mélanie Tremblay, and Mariana Oliveira.

I would also like to thank my master student Julia Müller who worked hard and remained positive despite a challenging project. Furthermore, my thanks go to Olha Wuerthinger for the support.

I thank Eric Fischer for his support with the headspace GC-FID measurements, which mostly took place in the evenings and weekends. I greatly appreciated his can-do spirit.

My thanks also go to Meriam Kabbaj and Vincent Forster who work very hard to translate academic developments into patient benefit.

I thank Susanne Freedrich (EPIC) for her support in animal-related matters.

I would also like to express my thanks to the Scholarship Fund of the Swiss Chemical Industry (SSCI) for awarding me a doctoral scholarship.

I further acknowledge generous funding from the Swiss National Science Foundation (2-77082-16).

My thanks go to Scope M for the cryo-SEM measurements.

I would like to express my thanks to every former and current member of the Leroux group. Every one of you is so special and fantastic that I do not dare to mention anybody by name.

Finally, I thank my family and friends for celebrating the good moments with me and putting the less good ones into perspective. I could not be happier to be surrounded by such wonderful people.

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