THE FUNCTIONAL CONSIDERATION OF RENIN ANGIOTENSIN SYSTEM IN THE HUMAN JEJUNUM

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LITHUANIAN UNIVERSITY OF HEALTH SCIENCES MEDICAL ACADEMY

Mantas Malinauskas

THE FUNCTIONAL CONSIDERATION

OF RENIN ANGIOTENSIN SYSTEM

IN THE HUMAN JEJUNUM

Doctoral Dissertation Biomedical Sciences,

Biology (01B)

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The dissertation has been prepared during the period of 2011–2015 at the following departments: Institute of Physiology and Pharmacology of Medical Academy of Lithuanian University of Health Sciences and the Department of Gastrosurgical Research and Education, Institute of Clinical Sciences, Sahlgrenska Academy, University of Gothenburg, Gothenburg, Sweden.

Scientific Supervisor:

Prof. Dr. Edgaras Stankevicius (Lithuanian University of Health Sciences, Biomedical Sciences, Biology – 01B)

Consultant:

Prof. Dr. Almantas Maleckas (Lithuanian University of Health Sciences, Biomedical Sciences, Medicine – 06B)

Dissertation is defended at the Biology Research Council of the Medical Academy of Lithuanian University of Health Sciences.

Chairperson

Prof. Dr. Antanas Gulbinas (Lithuanian University of Health Sciences, Biomedical Sciences, Biology – 01B)

Members:

Prof. Dr. Vytenis Arvydas Skeberdis (Lithuanian University of Health Sciences, Biomedical Sciences, Biology – 01B)

Prof. Dr. Habil. Albinas Naudziunas (Lithuanian University of Health Sciences, Biomedical Sciences, Medicine – 06B)

Prof. Dr. Gintaras Valincius (Vilnius University, Biomedical Sciences, Biology – 01B)

Assoc. Prof. Dr. Vladimir V. Matchkov (Aarhus University, Biomedical Sciences, Biology – 01B)

The dissertation will be defended at the open session of the Biology Research Council of Lithuanian University of Health Sciences on the 17th June, 2016 at noon in the prof. Vl. Laso auditorium of Lithuanian University of Health Sciences.

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LIETUVOS SVEIKATOS MOKSLŲ UNIVERSITETAS MEDICINOS AKADEMIJA

Mantas Malinauskas

RENINO ANGIOTENZINO SISTEMOS

FUNKCIJOS TYRIMAS

ŽMOGAUS

TUŠČIOJOJE ŽARNOJE

Daktaro disertacija Biomedicinos mokslai, biologija (01B) Kaunas, 2016 3

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Disertacija rengta 2011–2015 metais Lietuvos sveikatos mokslų universiteto Medicinos akademijos Fiziologijos ir farmakologijos institute ir Geteborgo universiteto Sahlgrenskos akademijos Gastrochirurgijos mokslų ir ugdymo fakulteto Klinikinių tyrimų institute, Švedija.

Mokslinis vadovas:

Prof. dr. Edgaras Stankevičius (Lietuvos sveikatos mokslų universiteto Medicinos akademija, biomedicinos mokslai, biologija – 01B)

Konsultantas:

Prof. dr. Almantas Maleckas (Lietuvos sveikatos mokslų universiteto Medicinos akademija, biomedicinos mokslai, medicina – 06B)

Disertacija ginama Lietuvos sveikatos mokslų universiteto Medicinos akademijos biologijos mokslo krypties taryboje:

Pirmininkas:

prof. dr. Antanas Gulbinas (Lietuvos sveikatos mokslų universitetas, biomedicinos mokslai, biologija – 01B)

Nariai:

prof. dr. Vytenis Arvydas Skeberdis (Lietuvos sveikatos mokslų universitetas, biomedicinos mokslai, biologija – 01B);

prof. habil. dr. Albinas Naudžiūnas (Lietuvos sveikatos mokslų universitetas, biomedicinos mokslai, medicina – 06B);

prof. dr. Gintaras Valinčius (Vilniaus universitetas, biomedicinos mokslai, biologija – 01B);

doc. dr. Vladimir V. Matchkov (Aarhus universitetas, biomedicinos mokslai, biolo– gija – 01B).

Disertacija bus ginama viešame biologijos mokslo krypties tarybos posėdyje 2016 m. birželio 17 d. 12 val. Lietuvos sveikatos mokslų universiteto prof. Vl. Lašo vardo auditorijoje.

Adresas: A. Mickevičiaus g. 9, LT-44307 Kaunas, Lietuva.

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LIST OF ABBRIAVIATIONS

ACE – angiotensin-converting enzyme ACE2 – angiotensin-converting enzyme 2 AGT – angiotensingen

Ang – angiotensin Ang(1-7) – angiotensin (1-7) AngII – angiotensin II AngIII – angiotensin III AngIV – angiotensin IV AP-A – aminopeptidase-A AP-B – aminopeptidase-B AP-M – aminopeptidase-M AP-N – aminopeptidase-N

AT1R – angiotensin II, type 1 receptor AT2R – angiotensin II, type 2 receptor AT4R – angiotensin IV receptor BMI – body mass index Cep – epithelial capacitor

EDTA – ethylenediaminetetraacetic acid EIA – enzyme immunoassay

ELISA – the enzyme-linked immunosorbent assay GAPDH – glyceraldehyde-3-phosphate dehydrogenase GI – gastrointestinal

GIP – glucose-dependent insulinotropic peptide GLP-1 – glucagon-like-peptide

GLUT2 – glucose transporter 2

GPCRs – G-protein-coupled receptors HMIT – H+/myo-inositol transporter Iep – epithelial ion current

IHC – immunohistochemistry

IRAP – insulin-regulated aminopeptidase IRS – insulin receptor substrate

Isc – short-circuit current

MAPK – mitogen-activated protein kinase MasR – Mas receptor

NADPH – nicotinamide adenine dinucleotidephosphate NEP – neutral endopeptidase

PBS – phosphate buffered saline PCR – the polymerase chain reaction

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PD – potential difference PPR – prorenin receptor

Ra – apical membrane resistance

RAS – renin angiotensin system Rb – basolateral membrane resistance

Rep – epithelial resistance

Rics – intercellular space resistance

ROS – reactive oxygen species

Rsubep – subepithelial resistance

Rte – transepithelial resistance

Rtj – tight junction resistance

SEM – standard error of the mean

SGLT1 – sodium-dependent glucose transporter STRs – sweet taste receptors

TER – transepithelial electrical resistance TTX – tetrotodoxin

UPM – Ussing pulse method

Va – the voltage of the apical membrane

Vb – the voltage of the basolateral membrane

Vm – mucosal membrane potential

VSMC – vascular smooth muscle cell Vte – transepithelial voltage

WB – western blot

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INTRODUCTION

The classical renin-angiotensin system (RAS) certainly known as endo-crine system is one of the body’s most powerful regulators of body fluid balance and having a role in the cardiovascular system [73, 131]. The classical bioactive peptide of RAS – Angiotensin II (AngII) has been imply-cated in many physiological functions by the activation of AngII type 1 receptor (AT1R) and AngII type 2 receptor (AT2R) [36]. The main enzymes responsible for the biodegradation of RAS peptides are angiotensin-converting enzyme (ACE), ACE2, aminopeptidases A (AP-A), B (AP-B), M (AP-M; also termed aminopeptidase-N) and neutral endopeptidase (NEP) (Figure 2.3.3.1) [8]. AP-A acts on AngII to produce AngIII, which exerts its effects via the AT1R and AT2R [40]. AP-B or M converts AngIII into the hexapeptide AngIV, which exerts its effects via binding to the insulin regulated aminopeptidase receptor (IRAP) and displays a modest affinity for AngII receptors [21]. NEP, conversely, converts AngII into Ang(1-7), which may exert effects distinct (and sometimes even opposite) from those of AngII by activating a specific non-AT1, non-AT2 receptor – the Mas receptor (MasR) [142]. These degradations pathways displays the formation of the so-called alternative RAS axes: AP/AngIV/IRAP or ACE2/Ang(1-7)/MasR (Figure 2.3.3.1).

Recent research has recognized RAS to be locally expressed with paracrine/autocrine functions [90]. Local RAS refers to tissue-based mecha-nisms of Ang peptide formation that operate separately from circulating RAS. Although many different concepts of local RAS have been described, a key feature is the local synthesis of RAS components including angioten-sinogen and enzymes such as renin that cleave angiotenangioten-sinogen to produce Ang peptides independently of the circulating RAS. ACE, (AT1R) and (AT2R) are invariably locally synthesized, but these are also components of the circulating RAS [21]. There are many evidence suggesting the possibi-lity of local RAS that may operate independently of circulating RAS and play pathogenic or protective role [92]. These functional local RAS have been found in such diverse organ systems as the pancreas, heart, kidney, vasculature and adipose tissue as well as the nervous, reproductive systems [132]. Recently, has been shown that RAS expressed also in the digestive tract [144]. There are several reports showing influences by RAS and its key mediator (AngII) on intestinal epithelial fluid and electrolyte transport and data are accumulating, suggesting involvement in GI mucosal inflammation and carcinogenesis [47]. In addition, recent researches reported, that RAS components such as AngII, AT1R and AT2R are located in a variety of cells

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in human gastric mucosa [65]. Moreover, there was investigated, that local AngIV is involved in esophageal aberrations associated with gastro-eophageal reflux disease [16]. There are also evidences about AngII expression in human small intestine and its role on jejunal wall contraction [173, 46].

Small intestinal glucose absorption plays a pivotal role in glycemic control. These principal glucose transporters (SGLT1, GLUT2) of the enterocyte have been extensively studied over the last decades and an intricate regulation of their activity and expression has been established [87, [182]. However, most of the studies have been conducted on rodent models or in cell cultures, and several paradigmatic features have not yet been confirmed in human [162]. Wong et al. showed in rats that the small intesti– nal epithelial brush border membrane has the capacity for local formation of AngII. The authors also demonstrated that activation of the AT1R inhibited SGLT1-mediated glucose absorption [176]. Furthermore, this mechanism was shown to be downregulated in experimental (streptozotocin-induced) type-1 diabetes in association with a relocation of GLUT2 to the brush border of the enterocytes resulting in an un-controlled glucose uptake [177].

There are also evidence that the an alternative axis of the RAS AP/AngIV/IRAP is expressed and exerts its effects in the mucosa [17] and the muscular tissue [37] of digestive tract. Some data has demonstrated, that IRAP is co-expressed with the glucose transporter GLUT4 in adipocytes and myocytes, thus enhanced insulin regulated glucose uptake [80].

Previous studies has been devoted to the action of RAS in rat’s small intestine. The present thesis focuses on the expression and function of RAS in human jejunum. The scientific background of the thesis and the results from new research are summarised and discussed below.

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1. THE AIM AND GOALS OF THE THESIS,

SCIENTIFIC NOVELTY OF THE WORK

The aim of the thesis

Investigate the expression and the potential functions of the renin angiotensin system (RAS) components in the jejunal mucosa of healthy individuals and jejunal muscular layer of the obese patients in vitro.

The goals of the study

1. To assess the expression of AngII receptors (AT1R, AT2R), AngIV receptor (IRAP), aminopeptidases-A, B, M, and glucose transport-ters (SGLT1, GLUT2) in the jejunal mucosa of healthy subjects. 2. To investigate the effects of AngII via both receptor subtype and

AngIV on glucose induced SGLT1 mediated transport in the jeju-nal mucosa of healthy individuals.

3. To assess the expression of aminopeptidases-A, B, M as well as AngIV receptor (IRAP) in the jejunal muscular tissue of obese patients.

4. To investigate the contractile function of AngIV in both longitu-dinal and circular jejunal muscular layers of obese patients.

The scientific novelty of the work

Unfortunately, there is a lack of data suggesting the presence and actions of the RAS in the small intestine, particularly in the human jejunum, furthermore no data suggesting the impact of the RAS in glucose regulation in the human jejunal mucosa.

The thesis demonstrates that the RAS locally expressed in the human jejunum, and in turn have a role on glucose absorption. The experiments was confirmed the presence of the “classical” RAS in the human jejunum mucosa. Moreover, the present data showed that the AngII type 1 and type 2 receptors (AT1R and AT2R) simultaneously exert the opposite effect on SGLT1-mediated glucose transport in human jejunum epithelium. The investigations also revealed the presence of an alternative RAS, the enzyma-tic capability responsible to AngIV and IRAP (AngIV receptor) formation, thus there was a key to following investigations, which was suggested an AngIV-mediated stimulation of glucose absorption. The jejunal mucosa biopsies were collected from healthy volunteers and was analysed using Western blot, ELISA and immunohistochemistry. The square wave current pulse analysis was investigated for its applicability in Ussing chambers for

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assessing mucosal epithelial resistance (Rep), which in turn enables

calcula-tion of the epithelial ion current (Iep). The square wave current pulse method

was first time tested only in our investigations assessing the glucose transport in the human jejunal mucosa.

In order, to expand the map of the RAS in the human jejunum, the investigations was carried on with the expression of an alternative RAS and its bioactive peptide AngIV contractility actions in human jejunum musculature. Previously, was only one study in rats, suggested the AngIV contractility effect in smooth muscles [37].

Summarizing, the novelty of thesis is that the present investigation first time provides the data in the healthy human jejunal mucosa, where was successfully investigated the locally expressed RAS and its effects on glucose uptake, indicating an AT1R mediated inhibition and AT2R enhan-cement of SGLT1-mediated glucose uptake. Moreover, the investigation in the human jejunal musculature first time demonstrate the local expression of AP/AngIV/IRAP axis of an alternative RAS, and pharmacological analyses

in vitro strongly indicate that the AngIV-induce contraction of both

longitu-dinal and circular muscles.

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2. LITERATURE REVIEW

2.1. The human jejunum 2.1.1. Gross anatomy and physiology

The jejunum is the midsection of the small intestine, connecting the duodenum to the ileum. It is about 2.5 m long, which has a median external diameter of 4 cm and an internal diameter 2.5 cm [159]. The suspensory muscle of duodenum is a thin muscle connecting the junction between the duodenum, jejunum, and duodenojejunal flexure to connective tissue sur-rounding the superior mesenteric artery and coeliac artery. It is also known as the ligament of Treitz, which marks the formal division between the first and second parts of the small intestine, the duodenum and the jejunum [109].

No anatomical feature separates the jejunum from the ileum, and the structure of the jejunum and ileum is basically similar. With distance from the duodenum, there is a gradual decrease in diameter, in the thickness of the wall, and in the number of folds. In the upper half of the jejunum the circular folds are large and numerous; however from this point, down to the middle of the ileum, they decrease significantly in size. The function of the folds slow the passage of the food along the intestine and thereby increase the surface for absorption. The folds arecovered with small projections called villi. Each unit of villi called villus, in turn is covered with microvilli. Villi in duodenum is mainly leaf-like, whereas in the jejunum and ileum mainly finger-like. The microvillous surface of the jejunum is known as the brush border is specialized for the absorption of small nutrient particle, which have been previously digested by enzymes in the duodenum. It is interesting to note that the villi in the jejunum are longer than those in the duodenum and the ileum ensure the greater absorption of the nutrients such as proteins, carbohydrates, amino acid, sugar, fatty acid particles, vitamins, minerals, electrolytes, and water [73]. Morphometric data obtained by light and electron microscopy of biopsies demonstrate that villi and microvilli together amplify the small intestinal surface area by 60–120 times. It follows that the total mucosal surface area of the small intestine (duodenum, jejunum and ilium) averages 30m2. Interestingly, the total area of the skin is about 1.8 m2[71].

Motility of the jejunum is organized to optimize the process of absorp-tion. Contraction of the jejunum are effected by the activities of two layers of smooth muscle cells: an outer longitudinal layer, and an inner circular layer. The jejunum is richly innervated by elements of the autonomic

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vous system. Preganglionic parasympathetic nerve fibers of the vagus nerve synapse with the cells of the terminal ganglia in the myenteric plexus or Auerbach plexus. The postganglionic nerves stimulate muscle contraction and gland secretion. Postganglionic sympathetic nerve fiber arising largely from the prevertebral ganglia mostly innervate their target effector cells directly [167].

2.1.2. Tissue structure

The jejunum as well as whole small intestine is subdivided into various tissues layers. From inside out, there are four characteristic layers (Fig. 2.1.2.1). The most inner layer is mucosa that consist of an epithelium, the lamina propria, and the muscularis mucosae. There are exocrine and endocrine cells in the epithelium, which secrete mucus and digestive enzymes into the lumen, and proportionately release gut hormones into the blood.

Fig. 2.1.2.1. The structure of jejunum tissue

The figure shows the composition of the four essential human jejunum layers from inside out, i.e. mucosa, submucosa, muscularis externa and serosa.

These enteroendocrine cells are situated among the epithelial surface, thereby constitute the gut endocrine system. The lamina propria contains of small blood vessels, nerve fibers, and lymphatic cell/tissues. The muscularis

mucosae is a thin muscle layer responsible for controlling mucosal blood

flow and gut secretion.

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The following layer submucosa made of connective tissue with major blood and lymphatic vessels, passing along with a network of nerve cells, called the submucosal nerve plexus [93].

The third layer muscularis externa is a thick muscle and its contraction contributes to major gut motility (segmentation and peristaltysis). The muscularis consists of inner circular smooth muscle and outer longitudinal smooth muscle. Previously mentioned myenteric nerve plexus located between the circular and longitudinal smooth muscle layers. The myenteric nerve plexus of Auerbach and the submucosal nerve plexus of Meissner constitute the enteric nervous system [167].

The last layer serosa also called pariental layer is the most exterior layer mainly consists of connective tissue and it connects to the abdominal wall, thus supporting the GI tract in the abdominal cavity [93].

2.1.3. Epithelial tissue structure

A simple columnar epithelium are columnar epithelial cells that is mono-layered and lines most organs of digestive tract as well as small intestine. At least five types of cells are found in intestinal mucosal epithelium. These type of cells are exist in the intestinal glands and on the surface of the villi. They comprise:

• Enterocytes, whose primary function is absorption [31].

• Goblet cells, and their main secretory product, mucus lubricates and protects the inner surface of the human intestine [15].

• Paneth cells, these cells synthesize and secrete substantial quanti-ties of antimicrobial peptides and proteins [33].

• Enteroendocrine cells – located within the intestinal crypts and villi, and comprising ~1% of the epithelial cell population [112], these cells regulate numerous processes in the body, including controlling glucose levels, food intake, and stomach emptying [186].

• M cell –found in the follicle-associated epithelium of the Peyer's patch, and the main function of Mcells seems to be the rapid uptake and delivery of particular antigens and microorganisms to the immune cells of the lymphoid follicle to induce an effective immune response [86].

The process of the differentiation of enteroendocrine cells begins as proliferating pluripotent stem cells at the base of the intestinal crypt [60]. As daughter cells of the pluripotent stem cells migrate from the base of the crypt towards the surface epithelial cuff, they commit to one of the four cellular lineages [64]. Enteroendocrine cell-committed differentiating cells

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migrate slowly in comparison with differentiating cells of the goblet and enterocyte lineages. Paneth cell-committed differentiating cells migrate towards the base of the crypts as they mature (Fig. 2.1.3.1).

Fig. 2.1.3.1. Development of intestinal enteroendocrine cells

The process of differentiation of enteroendocrine cells begins as proliferating pluripotent stem cells at the base of the intestinal crypt and progresses as daughter cells migrate, in an upward linear fashion, towards the epithelial cuff at the luminal surface Modified from: Gunawardene et al., Int. J. Exp. Pathol. 2011.

2.1.4. Transcellular and paracellular transport

There are two routes for transport of molecules and ions across the epithelium of the gut (Fig. 2.1.4.1):

• Transcellular transport, the movement of ions and small molecules through a cell, is accomplished by the differential distribution of membrane transporters/carriers on opposite sides of a cell.

• Paracellular transport, the movement between adjacent cells, is accomplished by regulation of tight junction permeability, which is regulated and varies among epithelia in tightness and ion selecti-vity. This regulated back diffusion can significantly modify the molecular composition of transcellular transport [129, 5].

Water transport by the intestine is closely coupled with solute move-ment and is passive. Theoretically, water flow could occur by the transcel-lular and paraceltranscel-lular routes, but the prevailing evidence indicates that water transport by the intestine occurs through the paracellular pathway [93].

In order for such transepithelial transport to occur, the epithelial cell must be polarized, with different sets of transport proteins localized in the

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basolateral and apical surfaces. There are three types of transport proteins that are found in the plasma membrane of intestinal cells: pumps, channels and carriers [71].

Fig. 2.1.4.1. Transepithelial transport pathways

The transcellular route is both active and passive and based on the activity of transmem-brane pumps, channels, and carriers expressed in a polarized fashion. Paracellular transport is only passive and driven by the gradients secondary to transcellular transport mechanisms. The major barrier in the paracellular route is the tight junction. Modified from: Anderson, Physiology. 2001.

Pumps such as sodium pump (Na+, K+-ATPase) and proton pump (H+/K+-ATPase) are energy-driven and capable of transporting ions against electrochemical gradients. In the intestinal epithelium, for instance, the sodium pump is essential for establishing and maintaining electrochemical gradients (low intracellular Na+ and electronegative membrane potential) that are required for other types of passive and facilitated transport process-ses [95, 100].

Channel proteins form pores through the membrane, allowing the free passage of any molecule of the appropriate size. Ion channels allow the passage of inorganic ions such as Na+, K+, Ca2+, and Cl- across the plasma membrane [31]. For instance, electrogenic Cl- secretion in the gut causes a potential difference (serosa is positive relatively to lumen) across the mucosa that promotes passive transport of Na+ resulting in net sodium chloride secretion [59, 84].

In contrast, to channel proteins, carrier proteins or transporters selecti-vely bind and transport specific small molecules, such as glucose rather than forming open channels, carrier proteins act like enzymes to facilitate the passage of specific molecules across membranes [31]. Several types of transporters exist in the intestinal epithelium. Uniporters that is involved in

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facilitated diffusion, transport one molecule at a time down a concentration gradient. This type of transporter, for example, moves glucose across the intestinal epithelium [160]. In contrast, antiport and symport processes are associated with secondary active transport, meaning that one of the two substances are transported in the direction of its concentration gradient, utilizing the energy derived from the transport of such substance (mostly Na+, K+ or H+ ions) down its concentration gradient. Antiporters or exchangers such as the Na+/H+ and Cl-/HCO3-, exchange one molecule for

another [138, 19]. Symporters, such as sodium-glucose transport proteins simultaneously carry glucose and Na+ across the membrane. However, transport by these carriers occurs only if all solutes are present, andglucose transport will not occur if an inwardly directed Na+ gradient is absent [182]. The distinction exist between these two transport ways. Transcellular transport often involves energy expenditure whereas paracellular transport is unmediated and passive down a concentration gradient. Paracellular trans-port also has the benefit that absorption rate is matched to load because it has no transporters that can be saturated [6]. Intestinal absorption of nutrients is considered to be dominated by trancellular transport, for instance, glucose is absorbed through the glucose transporters situated in the apical and basolateral membranes of the cells [180]. Paracellular absorption therefore plays only a minor role in glucose absorption, whereas glucose is absorbed transcellularly, the paracellular pathways opens and becomes more permeable to small nutrients. The basolateral deposition of Na+ and nutrients establishes an osmotic gradient that provides the force for water flow. Small nutrients can then be carried across the paracellular pathways with the water by solvent drag. This paracellular transport is facilitated by opening of tight junctions following transcellular Na+-nutrient cotransport. Thus, total glucose absorption continuous to increase well after transcellular absorption [28].

Tight junctions control paracellular permeability. Claudins and occludin are two major transmembrane proteins in tight junctions, which directly determine the paracellular permeability to different ions or large molecules. Intracellular signalling pathways including Rho/Rho-associated protein kinase, protein kinase Cs, and mitogen-activated protein kinase, modulate the tight junction proteins to affect paracellular permeability in response diverse stimuli [74]. Cytokines, growth factors and hormones in organism can regulate the paracellular permeability via signalling pathway [136].

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2.1.5. Bioelectric properties of epithelia

Transepithelial electrical resistance (TER) measurements are widely

used as real-time, non-destructive, and “label-free” measurements of epithelial barrier function (Fig. 2.1.5.1).

Transepithelial electrical resistance probably the most sensitive measure of mucosal barrier function and this measurement reflects the degree to which ions traverse tissue. Ions passively traverse epithelium both transcel-lularly and paraceltranscel-lularly. In the small intestine, apical epithelial membranes have a resistance to passive flow of ions 1.5–3 log units greater than that of epithelial monolayer as a whole. Thus measurements of TER mostly reflect the paracellular resistance, which has been calculated to account for 75– 94% of the total passive ion flow across small intestinal epithelium [18]. An example of such a probe is mannitol, which has been used in permeability studies either as mucosal to serosal fluxes in tissues placed in Ussing chambers [104, 18].

Epithelial tissues are able to transport ions and as a result, generate a transepithelial voltage Vte. This voltage also usually termed as a transmural/

transepithelial electrical potential difference PD. An electrical potential difference can be measured across the wall of the animal and human small intestine, the polarity being serosa positive to mucosa [94]. The net movement of negative or positive charges from the apical to the basolateral side generates a voltage that is equal to the difference between the voltage of the apical membrane Va and of basolateral membrane Vb, respectively.

Measurements of Vte are performed in current-clamp and often referred to as

open-circuit recordings. They are useful in studying absorptive tissues, Va is

mostly effected by the activity of Na+ channels, whereas in secretory epithelia, Va is determined by a Cl- conductance [95].

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Fig. 2.1.5.1. Measurements of transepithelial electrical resistance

Diagram of the intestinal epithelium representing components that contribute to the electrical measurement of TER. Note the critical components of this monolayer that make up the components of TER, including the epithelium itself and the paracellular space. Rtj,

tight junction resistance; Rics, intercellular space resistance; Ra, apical membrane resistance;

Rb, basolateral membrane resistance. Modified from: Blikslager et al., Physiol. Rev. 2007. Addition of D-glucose to the mucosal fluid resulted in a significant depolarization of the mucosal membrane potential Vm in rat duodenum,

jeju-num, and ileum accompanied an increase in the transepithelial potential difference PDt. In case of Phlorizin addition (inhibitor of sodium/glucose

cotransporter) to the mucosal fluid inhibits the sugar-dependent changes in PDt and Vm. According to the analysis with an equivalent circuit model for

the epithelium, it was concluded that an actively transported solute induced not only a depolarization of the mucosal (brush border) membrane but also a hyperpolarization of the serosal (basolateral) membrane of an epithelial cell. The hyperpolarization of the serosal membrane in the presence of an actively transported solute was clarified to a mechanism of serosal electro-genic sodium pump stimulated by the increase in the extrusion rate of Na+ co-transported into the cell with sugar [123].

The third essential electrogenic measurements of epithelia is the short-circuit current Isc and determined as the charge flow per time when the

tissue short-circuited (i.e. Vte is clamped to 0 mV). To measure Isc, a current

that is adjusted by a feedback amplifier to keep Vte at 0 mV is injected

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across the epithelium. In conditions when Rte and Vte are known is possible

to calculate Isc There are simple equation for Isc calculation 𝐼𝑠𝑐=_𝑅𝑉𝑡𝑒 𝑡𝑒 [95].

2.2. Intestinal absorption of monosaccharides 2.2.1. Hexoses/glucose transporters

Hexoses are major metabolic fuel for numerous cell types, and the original studies of their movement across cell membranes and epithelia led to the development of the whole field of membrane transport proteins [105]. There are two types of hexose transporter in the mammalian cells: Na+/glucose cotransporter, which are associated with the secondary active transport of glucose, and sodium-independent facilitative hexose transporters [145].

There are three families of glucose transporters in the human genome, SLC2, SLC5 and SLC50. The SLC2 family of facilitated transporters (GLUTs), the SLC5 family of “active” glucose transporters, and the SLC50 family of uniporters, SWEETs. The founding member of the SLC5 family is the intestinal glucose cotransporter, SGLT1, which is encoded by the SLC5A1 gene and is responsible for the “active” absorption of glucose from the intestine, glucose-lactose-malabsorption, and is the rationale of Oral Rehydration Therapy for Diarrhea. The second member of this family, SGLT2, is encoded by the SLC5A2 gene, is primarily responsible for glu-cose reabsorption in the kidney, and is a new target for drugs to treat diabetes [179]. Sweets and their prokaryotic homologues are monosaccha-ride and disacchamonosaccha-ride transporters that are present from bacteria to plants and humans. Sweets play crucial roles in intercellular transport and cellular secretion. The human genome contains a single SWEET homologue, which functions as a glucose transporter. Their bacterial homologues, which are called Semisweet are among the smallest known transporters [188].

SGLT family consists of 12 proteins, which include sugar cotranspor-ters of anions, vitamins and short-chain fatty acids. Some of them also have a function of glucose sensors as well. GLUT family consist of 14 proteins grouped in 3 subclasses based on similarities in their architecture. They differ from one to another in affinity to glucose, tissue distribution and type of signals that cause their translocation to the cell membrane what results in different levels of sugar transport into the tissues [103].

As was mentioned above GLUT proteins are encoded by the SLC2 genes and are members of the major facilitator superfamily (MFS). The 14 GLUT proteins are comprised of ~500 amino acid residues and can be categorized into 3 classes based on sequence similarity: Class 1 (GLUTs1-4,

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14) Class 2 (GLUTs 5, 7, 9, and 11); Class 3 (GLUTs 6, 8, 10, 12, 13 (H+/myo-inositol transporter)) (Fig. 2.2.1.1).

Fig. 2.2.1.1. The glucose transporter family members.

Unrooted phylogenetic tree showing the relationship between the 14 human GLUT protein family members. Modified from: Augustin et al., IUBMB Life. 2010.

GLUT1, encoded by the SLC2A1 gene, was one of the first membrane transporters to be purified and cloned and is likely one of the most exten-sively studied of all membrane transport systems. GLUT1 is highly expres-sed in the endothelial cells of erythrocytes and the blood-brain barrier where it is mainly involved in glucose transport into the brain [135].

GLUT2 was first characterized by cDNA cloning the Slc2a2/SLCA2 gene from rat and human liver and is the major glucose transporter of heap-tocytes. It is also expressed by the intestinal absorptive cells. In the intes-tine, GLUT2 is the major basolateral glucose transporter isoform. [114].

GLUT3 was the third glucose transporter to be cloned and was origin-nally designed as the neuronal glucose transporter. More, recently GLUT3 has been studied in other cell types with quite specific requirements for glucose, including sperm, preimplantation embryos, circulating white blood cells, and an array of carcinoma cell lines [151].

GLUT4 is expressed most prominently in adipocytes, skeletal muscle, and cardiomyocytes. GLUT4 is regulated by its intracellular localization. In the absence of insulin, GLUT4 is efficiently retained intracellularly within storage compartments in muscle and fat cells. Upon insulin stimulation (and contraction in muscle), GLUT4 translocates from these compartments to the cell surface where it transports glucose from the extracellular milieu into the cell. Its implication in insulin-regulated glucose uptake makes GLUT4 not

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only a key player in normal glucose homeostasis but also an important element in insulin resistance and type 2 diabetes [62].

GLUT5 was identified as the SLC2A5 gene and was initially cloned from human small intestine. The small intestine regulates fructose absorp-tion from dietary sources and, therefore, the availability of fructose to other tissues. After apical transport mediated by GLUT5, fructose is transported across the basolateral membrane by GLUT2. GLUT5 is expressed in skele-tal muscle and fat tissue, has consistently been found in the spermatozoa, and also GLUT5 mRNA is abundant in the cytosol, and in the apical plasma membrane of proximal tubule cells [43].

GLUT6 (formerly designated GLUT9) was cloned from human leuco-cyte cDNA by (RACE)-PCR and identified as the SLC2A6. GLUT6 mRNA is predominantly identified in the brain, spleen and peripheral leucocytes, however is not expressed in insulin-sensitive tissues [96].

Recent data suggest that a transporter GLUT7 exist and primarily ex-pressed in the small intestine and colon (Fig. 2.2.1.2). The protein is expres-sed in the apical membrane of the small intestine and colon and it has a high affinity for glucose and fructose. The richness of GLUT7 in the small intes-tine does change in parallel with the dietary carbohydrate, however the distribution of this protein along the small intestine does not entirely match with the availability of glucose and fructose [30].

GLUT8 was the first isoform of the extended SLC2A family to be identified by database mining and was cloned from the human, rat and mou-se testis cDNA samples. Recently, demonstrated that GLUT8 is a plasma membrane-localized hepatic transporter and that GLUT8 is rate limiting for hepatic fructose uptake [35].

GLUT9 mRNA is detected almost exclusively in the kidney and liver and at low levels in the small intestine, placenta, lung, and leucocytes. In the human kidney, GLUT9 is present in proximal convoluted tubules, whereas in rodents it is most probably in distal connecting tubules. Functional studies confirmed that GLUT9 is a urate transporter, whether GLUT9 plays role as a glucose/fructose transporter that potentially links fructose uptake by the liver [133].

GLUT10 exhibits a very wide tissue distribution and is expressed in pancreas, placenta, heart, lung, liver brain, fat, muscle, and kidney. Recent study suggest, that GLUT10 is mainly localized to the mitochondria of aortic smooth muscle cells and insulin-stimulated adipocytes [91].

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Fig. 2.2.1.2. Hexose transporter proteins expressed

in the intestinal enterocyte

There are approximately 4 hexose transporters expressed in the apical membrane of intestinal epithelial cells (enterocytes). The Na+-coupled high affinity glucose/galactose transporter SGLT1 and the facilitated transporter GLUT5 are both expressed constitutively, although the level of SGLT1 can be modulated up or down. The low affinity glucose/fruc-tose facilitative transporter GLUT2 is transiently expressed in the apical membrane when luminal hexose concentration are high. There appears to be only one facilitative hexoses transporter, GLUT2, in the basolateral membrane. The high affinity glucose/fructose facilitative transporter GLUT7 is also expressed in the apical membrane, though possible regulation has not been determined. Modified from: Cheeseman et al., Physiology. 2008.

Human GLUT11 (encoded by the solute carrier 2A11 gene, SLC2A11) is a novel sugar transporter, and it was exclusively detected in heart and skeletal muscle. The closest relative of GLUT11 is the fructose transporter GLUT5 (sharing 41.7% amino acid identity with GLUT11) [42].

GLUT12 is a member of class II glucose transporters, which preferen-tially transports D-glucose and 2-deoxy-D-glucose over other hexoses. The presence of targeting motifs similar to GLUT4 and GLUT8, and localization primarily in insulin-sensitive tissues, has led to research to clarify whether GLUT12 may represent a second insulin sensitive GLUT. In human skeletal muscle, GLUT12 translocates to the plasma membrane following euglyce-mic insulin infusion [107].

GLUT13 (H+/myo-inositol transporter, HMIT) belong to the Major Facilitator Superfamily. HMIT is encoded by a member of SLC2 gene family, SLC2A13. SLC2A13 is predominantly expressed in the brain and its expression is induced under hypertonic conditions [147].

GLUT14 (identified as the SLC2A14) was shown to specifically expres-sed in testis. Two alternative spliced forms of GLUT14 were identified [10].

Interestingly, that the expression of hexose transporters in the small intestine differs in parallel with the abundance of their hexose substrates in the diet, so that both facilitative (GLUTs) and concentrative

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dependent glucose transporters (SGLTs) increase when the carbohydrate intake is held at a particular level for several days [50].

2.2.2. The classical model of sugar absorption

The predominant carbohydrate of the body is glucose. Glucose in the blood is called “blood sugar” and is a major fuel source for most cells of the body. The major site of glucose absorption in the body is the epithelial of the small intestine, whereas reabsorption of glucose is carried out by epi-thelial cells of proximal tubule nephron [67].

Half a century ago it was established that glucose transport across the small intestine occurred by active transport, i.e., the sugar could be absorbed uphill against its concentration gradient both in vivo and in vitro and this uptake was blocked by metabolic poisons. The energy for active glucose transport is provided by the sodium gradient across the cell membrane, the Na+ glucose cotransport hypothesis first proposed in 1960 by Crane [182].

The classical model of glucose absorption (Na+_{-glucose cotransport) by}

epithelial cells of the small intestine that was developed from Crane’s initial proposed model is shown in Fig. 2.2.2.1. The entry step of glucose across the luminal membrane is via the Na+_{-glucose cotransporter SGLT1, and it is}

inhibited by phlorizin [44]. The driving force of this transporter is the Na+

electrochemical gradient that is generated and maintained by the basolateral Na+_/K+_{-ATPase Na}+_{pump can be blocked by quabain. Na}+ _{transported into}

the cell with glucose exits the cell across the basolateral membrane by the Na+ _{pump, which transports three Na}+ _{for exchange of two K}+_{is recycled}

back across the basolateral membrane via K+ channels, aiding in the maintenance of the negative membrane potential of the epithelial cell, which contributes to the electrochemical gradient for Na+_{entry across the apical}

membrane (Fig. 2.2.2.1) [66].

Glucose transported into the cell and accumulated by SGLT1, whereas across the basolateral membrane via glucose transporter 2 (GLUT2) [10].

The transepithelial transport of Na+ contributes to a charge separation across the epithelium, providing a driving force for Cl- _{absorption via the}

paracellular pathway. The absorption of Na+_{, Cl}-_{, and glucose generates a}

slight osmotic gradient across the epithelium, providing a driving force for water absorption via both trancellular and paracellular pathways (Fig. 2.2.2.1) [67].

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Fig. 2.2.2.1. Cellular model of the Na+ -glucose-coupled cotransport in leaky absorptive epithelium (jejunal cell).

The details of this cellular model are given in the text. SGLT1, Na+-glucose cotransporter 1; GLUT2, Glucose transporter 2. Modified from: Hamilton et al., Adv. Physiol. Educ. 2013.

Plasma glucose is maintained at ∼5 mmol/l. When the glucose concent-ration in the lumen is lower than in plasma, SGLT1 transports glucose uphill against its concentration gradient, however the concentration of fructose against a gradient is not necessary; the entry of fructose is therefore ensured by GLUT5, a facilitative transporter.

The classical model of intestinal glucose absorption is elegant in its sheer simplicity and effective in its ability to explain glucose absorption in many conditions, most notably at low concentration of glucose or in expe-riments using in vitro preparations of intestine, in which the lack of vascular clearance causes tissues accumulation [81].

2.2.3. GLUT2 at the apical surface of intestinal epithelium

Recent studies in morbidly obese humans as well as in obese mice suggest that GLUT2 may be present permanently in the apical membrane of enterocytes, thereby contributing to overshooting postprandial glucose levels.

GLUT2 is rapidly inserted into the apical membrane after a meal and thus intestinal glucose absorption by the apical GLUT2 pathway can be 3 to 5-times greater than by SGLT1 at the high concentration of sugar. The

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apical insertion of GLUT2 is triggered by membrane depolarization via SGLT1, resulting in a large influx of Ca2+_{, which induces a global}

cytoske-letal rearrangement, and leading to apical localization and activation of protein kinase C βII [72, 81]. A high sugar intake is a physiological regula-tor of GLUT2 recruitment into enterocyte brush border membrane and increasing sugar uptake threefold [61].

The recruitment of GLUT2 in brush border membrane was also obser-ved in conditions of increased calorie demand and glucagon-like peptide 2 treatment [9].

The outcome of animal experiments suggests that intestinal glucose absorption is strongly regulated in the presence of luminal glucose or sweet tastants via activation of intestinal sweet taste receptors (STRs) which are heterodimers of G protein-coupled receptors T1R2 and T1R3. Upon acti-vation, STRs have the capacity to promote accumulation of both apical and basolateral membrane GLUT2 during fasting has been reported in morbidly obese subjects [121].

Some data suggest that the insulin has an impact on intestinal sugar absorption. Acute insulin treatment before sugar intake prevented the inser-tion of GLUT2 into the brush border membrane and low in the basolateral membrane [163].

More recently was found that hyperglycaemia causes GLUT2 apical insertion to the brush border membrane of proximal tubule cells, and under insulin insensitivity or in lack of insulin, GLUT2 is not internalized, leading to increased reabsorption of glucose [34].

Recently was revealed that GLUT2 also can be inserted into the apical membrane of human enterocytes. In morbidly obese subjects, GLUT2 accu-mulated in apical and endosomal membranes. Apical GLUT2 was triggered by drugs such as metformin to provide a glucose exit pathway in to the lu-men contributing to the drug hypoglycaemic effect [1].

Unfortunately, GLUT2 trafficking in cell types other than enterocyte and kidney cell is not fully understood.

2.3. Renin angiotensin system (RAS) 2.3.1. The origin of the RAS

The renin-angiotensin system (RAS) has been known for more than a century as a cascade that regulates body fluid balance and blood pressure.

The history of the discovery of the RAS began in 1898 with studies made by Tigerstedt and Bergman, who reported that the arterial blood

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pressure could be regulated after the discovery of a soluble protein extracted from the kidney that increased blood pressure in rabbits, called “renin”[12].

In 1934, Harry Goldblatt induced experimental hypertension in dogs by clamping a renal artery. Using the same technique as Goldblatt, Braun-Menendez et al in 1940 isolated a vasoconstrictor substance responsible for the reno-vascular hypertension from renal venous blood of the hypertensive kidney dog, calling it “hypertensin”. Later study made by Page et al independently describe a vasoconstrictor substance by injecting renin into cats, called “angiotonin” and also described angiotensinogen, first referred to as a “renin activator“. And finally, the name “angiotensin” for the vaso-constrictor substance “hypertension” by Braun-Menendez and “angiotensin” by Page emerged in 1958 after they both agreed on this hybrid name, since these 2 substances proved to be the same potent vasoactive octapeptide [49].

2.3.2. The classical RAS

Angiotensin II (AngII) is an octapeptide produced from the substrate angiotensinogen through sequential enzymatic cleavages by renin and angiotensin converting enzyme (ACE). Renin cleaves angiotensinogen, forming angiotensin I (AngI) that in turn is converted to AngII by ACE. The angiotensinogen substrate is produced in the liver, while renin is produced by kidney and AngII in the vascular tissue [13].

Angiotensinogen (AGT) is a glycoprotein that is the unique substrate of the RAS. Many components of the renin angiotensin system such as renin, ACE, and AngII have been comprehensively studied in animal models.Conversely, many features of AGT has received less attention. This may be partially due to the traditional view of AGT as a passive substrate and lack of pharmacological inhibitors that directly reach the protein. Human AGT is a heterogenous plasma glycoprotein, mainly synthesized in hepatocytes [185].Recent published studies investigating the association between polymorphisms in the AGT gene showed association between polymerphisms and lung cancer [174], as well as relationship between AGT gene M235T polymorphism and hypertrophic cardiomyopathy [189].

In the normal, unstressed adult mammalian kidney, renin is found in a few cells of afferent arteriole at the entrance of glomerulus. Renin is an aspartyl protease synthesized as prorenin, a proenzyme that contains an additional 43-amino acid N-terminal fragment. The physiological matura-tion of prorenin into active renin takes place exclusively in the juxtaglome-rular cells of the kidney. Several proteins like mannose-6-phosphate recep-tor has been shown are able to bind renin and prorenin in several human tissues [120]. Prorenin receptor (PPR), also known as ATP 6-associated

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protein 2 (atp6ap2) [119]. At the beginning, the roles of PRR were thought to enhance the tissue (RAS) by binding PRR to its ligand, while also indu-cing intracellular signal transductions such as mitogen-activated protein kinase (MAPK) pathways independent of the RAS. However, recent findings suggest that PRR has a role in the kidney physiology and diabetic conditions [126].

Cathepsin D is a widely distributed aspartyl protease that enzymatically breaks down a variety of different protein substrates within the lysosomal compartment of most cells [63]. In certain circumstances, cathepsin D takes the same role as the renin, it can also generate AngI from angiotensinogen. This is likely because cathepsin D and renin share overall structural homo-logy and topohomo-logy, and their respective active sites are similar [118].

Angiotensin I is basically formed by the action of renin on angiotensi-nogen. Renin cleaves the peptide bond between the leucine and valine residues on angiotensinogen, creating the ten amino acid peptide AngI [13]. However, AngI appears to have no biological activity and exists solely as a precursor to angiotensin II.

ACEcatalyzes the conversion of AngI to AngII – a potent vasoconstric-tor by cleaving the C-terminal dipeptide His-Leu from AngI. ACE also metabolizes bradykinin – a potent vasodilator by removal of Phe-Arg from the C-terminus [109, 181]. The gene encoding human ACE has been locali-zed to chromosome locus 17q23 and comprises 26 exons. There are two forms of ACE: somatic and testicular. Somatic ACE has two active-site domains, designated the N- and C-domains, while testicular ACE contains only the C-domain active site [149, 82].

The therapeutic use of ACE inhibitors has been an integral part of the standard of care for hypertension and related diseases for decades. However, it was noticed, that long-term treatments with ACE inhibitors is often associated with so called “angiotensin escape” characterized by the return of plasma AngII to pre-treatment levels. It is often assumed that this rebound generation of AngII occurs through the action of the serine proteases such as cathepsin G and chymase (also known as chymostatin-sensitive AngII-generating enzyme). These two serine proteases cleaves AngI to AngII at a substantially greater rate than does ACE [99].

The classical RAS produces the AngII octapeptide through cleavage of inactive AngI by ACE, chymase or cathepsin-G. AngII peptide binds to G-protein-coupled receptors (GPCRs), namely AT1R or AT2R [139]. All classic physiological effects of AngII, such as vasoconstriction, aldosterone and vasopressin release, sodium and water retention, and sympathetic facilitation, are mediated by the AT1R. AngII, via its AT1 receptor, is also involved in cell proliferation [32], left ventricular hypertrophy [158],

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nephrosclerosis [69], endothelial dysfunction and processes leading to athero thrombosis [150]. AT2R has a role in cardiovascular, brain and renal function as well in the modulation of various biological processes involved in development, cell differentiation [191], tissue repair and apoptosis [78].

The classical RAS is depicted in the Fig. 2.4.2.1, where the figure sum-marizes the degradation of AGT a peptide with the sequence of 14 amino acids (1-14) to AngI (1-10) and AngII (1-8). The enzymatic pathways are mediated by renin, cathepsin D, ACE, cathepsin G and chymase. Some effects are mediated by Prorenin receptor, whereas the major effects of AngII are mediated by AT1, AT2 receptors (Fig. 2.3.2.1).

Fig. 2.3.2.1. The pathway of classical RAS peptides degradation.

The details of this figure are given in the text. ACE, angiotensin converting enzyme; AT1R, angiotensin type 1 receptor; AT2R, angiotensin type 2 receptor.

2.3.3. Alternative (novel) RAS

Angiotensin (1-12) [Ang(1-12)], a new member of the renin- angio-tensin system, is recognized as a renin independent precursor. However, the processing of Ang(1-12) in the circulation in vivo is not fully established. Although, some studies suggested, that Ang(1-12) is present in plasma and peripheral tissues including aorta, heart, and kidneys [117]. Early studies of Ang(1-12) metabolism showed that renin did not generate AngII from Ang(1-12), whereas a novel contribution of chymase as an Ang(1-12) degra-ding enzyme was found in neonatal cardiac myocytes of the rats [111].

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A homologue of ACE (a dipeptidase), known as ACE2 (a monopep-tidase), was described over a decade ago. In vitro experiments indicate that AngI may be converted in humans by ACE2 into Ang(1-9). In human heart tissue, the main products of AngI degradation are both Ang(1-9) and AngII [108]. Moreover, Ang(1-9) was detected in human plasma and a higher than AngII concentration of this peptide was found in the kidney [110]. It was also observed, that Ang(1-9) exerts an AngII-like prothrombotic effect due to the conversion to AngII in the circulatory system of rats, and that plate-lets are involved in this process [85].

ACE2 has distinct enzyme activity and substrate affinity, and cleaves a single amino acid from AngII to produce Ang(1-7). Ang(1-7) exerts its effects primarily via the Mas receptor. Together, these components (ACE2/ Ang(1-7)/Mas) comprise the alternative RAS pathway distinguishing them from the (ACE/AngII/AT1R) pathway which has become known as the classical RAS [55]. Ang(1-7) formation partly responsible of the neprilysin (NEP). NEP is a zinc-containing metallopeptidase catalysing the conversion of AngI to Ang(1-7), a potent vasodilator, thus counteracting the deleterious effects of AngII [153].

The next essential alternative RAS axis consist of Angiotensin III (AngIII), which is produced from the metabolism of AngII by aminopepti-dase-A (AP-A), cleaving theAsp1-Arg2 bound [54], than Angiotensin IV (AngIV) is generated by the enzyme aminopeptidase-B B) or- M (AP-M; also termed aminopeptidase-N), due to eliminate the amino acid arginine from the N-terminus of AngIII sequence [169]. AngIV binds to the AngIV receptor (AT4R), which has been identified as insulin-regulated aminopepti-dase (IRAP) [3]. A number of researches suggested that the AngIV/IRAP strongly influences memory disturbance and protects against brain ischemia [122].

The scheme of whole RAS system including the classical RAS as a precursor of the alternative RAS pathwaysis presented in the Figure 2.3.3.1. The details of the classical RAS pathwaywas mentioned in a previous section. As shown in Figure 2.3.3.1 there are five extra bioactive peptides: Angiotensin(1-12) produced directly from angiotensinogen by a non-renin enzyme; Angiotensin(1-9) produced from AngI by ACE2; Angiotensin(1-7) can be produced by the action of tissue endopeptidases especially neprilysin on AngI or from AngII by the action of ACE2, Ang(1-7) activate Mas receptor to produce counteracting effects mediated by AngII; AngIII and AngIV are also produced by cleavage of AngII respectively degrading by the AP-A, AP-M or AP-B and eventually AngIV binds to it’s receptors (IRAP) (Fig. 2.3.3.1).

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Fig. 2.3.3.1. The RAS cascade.

The details of this figure are given in the text. AP-A, aminopeptidase-A; AP-B, aminopep-tidase-B, AP-M, aminopeptidase-M; IRAP, insulin-regulated aminopeptidase.

2.3.4. RAS are locally expressed

Local RAS refers to tissue-based mechanisms of Ang peptide formation that operate separately from circulating RAS. Although many different concepts of local RAS have been described, a key feature is the local synthesis of RAS components including angiotensinogen and enzymes such as renin that cleave angiotensinogen to produce Ang peptides independently of the circulating RAS. ACE,(AT1R) and (AT2R) are invariably locally synthesized, but these are also components of the circulating RAS [23].

There are many evidence suggesting the possibility of local RAS that may operate independently of circulating RASand play pathogenic or pro-tective role [92]. These functional local RAS have been found in such diver-se organ systems as the pancreas, heart, kidney, vasculature and adipodiver-se tissue as well as the nervous, reproductive and digestive systems [132]. Local RAS operate in an autocrine, paracrine or intracrine manner and exhibit multiple physiological effects at the cellular level. In addition to hemodynamic actions, the local RAS has multiple and novel functions including the regulation of cell growth, differentiation, proliferation and

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apoptosis, reactive oxygen species (ROS) generation, tissue inflammation and fibrosis, and hormonal secretion [152].

Pancreatic RAS plays different roles in the pancreatic physiology and pathophysiology regulation. Overexpression of the local RAS components including angiotensinogen, renin and ACE suggest a potential role of the pancreatic RAS in acute pancreatitis [135, 182, 169]. An inhibition of AT1R by valsartan can activate the local RAS to protect against experimental acute pancreatitis through inhibition of microcirculation disturbances and inflammation [129].

The RAS has been established as an important modulator of cardiac remodelling mediating hypertrophy, fibrosis, as well as inflammation [11, 125, 72]. Some evidenceindicated that the intracellular and extracellular renin angiotensin aldosterone system is involved in the regulation of cell volume in the normal and particularly in the failing heart [38]. There are findings suggesting, that ACE and ACE2 expression seems to be a deter-minant factor on the regulation of cell function and cell volume and that the beneficial effect of ACE inhibitors or AT1R blockers during myocardial ischemia and heart failure be related to the prevention of cell swelling induced by extracellular renin and angiotensin II [39].

The complex and extensive actions of AngII on renal function are mediated by the widespread distribution of AngII receptors throughout the kidney in various nephron segments as well as in the vasculature and interstitium [143]. In addition to the classic RAS pathways, prorenin recep-tors and chymase are also involved in local AngII formation in the kidney [84]. The current studyestablished the enhanced excretion of ACE2 in both type 1 (streptozotocin-induced) and type 2 (db/db strain) diabetic models and reveals the potential role of the enzyme as a urinary biomarker of diabetic injury and/or activation of the intrarenal RAS [187].

Almost all components of RAS in vasculature were determined in adults [132]. Whereas, the dynamical expression patterns of RAS during development of vasculature in embryonic human have been investigated and showed, that the AT1R is predominantly expressed in vascular smooth muscle cell (VSMC) of umbilical cord vessels, whereas the AT2R seems to preferentially localize in VSMC of systemic blood vessels [14].

Components of the RAS are expressed in a number of areas in the brain involved in cardiovascular control [130]. However, the brain RAS is an important component in the development of dementia, Alzheimer's and Parkinson's diseases [184]. The brain has been shown to express the RAS components regulating neuronal development including learning and memo-ry processes [183].

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The accelerated adipose tissue growth and fat cell hypertrophy during the onset of obesity precedes adipocyte dysfunction. All components of the RAS are expressed in and have independent regulation of adipose tissue. This local adipose RAS exerts important auto/paracrine functions in modu-lating lipogenesis, lipolysis, adipogenesis as well as systemic and adipose tissue inflammation [52]. Upregulation of adipose RAS in obesity condi-tions, leads to development of hypertension, insulin resistance and other metabolic derangements [76].

Recently, has been shown that RAS can be expressed also in the digestive tract [144]. There are several reports showing influences by RAS and its key mediator AngII on intestinal epithelial fluid and electrolyte transport and data are accumulating, suggesting involvement in GI mucosal inflammation and carcinogenesis [47]. Also recent researches reported, that RAS components such as AngII, AT1R and AT2R are located in a variety of cells in human gastric mucosa and may have influence on gastric mucosa epithelial properties [65]. Moreover, there was investigated, that local AngIV is involved in esophageal aberrations associated with gastroesopha-geal reflux disease [17]. Furthermore, it has also been observedthat alocal RAS regulate gastrointestinal tract smooth muscles activities. Findings suggesting existence of a local RAS in human oesophageal muscular layer and its role in the physiological motor control for patients with achalasia [26]. There are evidences about AngII expression in human small intestinal and its role on jejunal wall contraction [173, 174].

Besides, there are new evidences suggesting, that RAS can be locally expressed in periodontal tissue and have an preventive role on alveolar bone loss in periodontal disease conditions [141].

2.3.5. RAS-mediated effects on glucose transport

Activation of the RAS causes insulin resistance directly. In muscle, AngII inhibits phosphorylation of insulin receptor substrate (IRS)-1, preven-ting increases in phosphatidylinositol 3 (PI3)-kinase and subsequent translo-cation of glucose transporter (GLUT4) to the cell membrane [124]. The mechanism through which AngII decrease insulin-dependent GLUT4 trans-location involves of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase and the formation of reactive oxygen species [175, 41]. Conversely, ACE inhibitors and AT1R blockers increase GLUT4 translocation to the membrane and improve skeletal muscle glucose uptake in rats [89]. AT1R blockers can also increase GLUT4 translocation and improve insulin sensitivity in cultured adipocytes and rodent model [53, 126]. Furthermore,

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activation of ACE2/Ang(1-7)/MasR axis results in increase basal as well as insulin-stimulated glucose uptake by murine adipocytes in vitro [98].

IRAP co-localizes with GLUT4 in neurons in regions of the brain associated with cognition, raising the question as to whether there is an analogous system in neurons as found in adipocytes and muscle cells. However, IRAP inhibitors could facilitate memory by potentiating glucose uptake into neurons [2]. In brain regions that are important in memory processing, such as in the pyramidal neurons of the hippocampus, IRAP is found in the same vesicles as GLUT4. And also was demonstrated, that in utilized hippocampal slices, IRAP inhibitors increased the amount of gluco-se taken up into active neurons via GLUT4 [4, 48].

Recently, was reported, that low dose of Ramipril effectively reduced the renal cortical GLUT2 content to a non-diabetic level [154].

The effects of intestinal RAS on glucose transport was investigated in several studies in rat models and including our group experiments in human tissue, where was shown, that the RAS bioactive peptides AngII [176, 177, 26] and AngIV [104] can alter the glucose uptake across brush border membrane.

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3. METHODOLOGICAL CONSIDERATIONS

3.1. Ethics

All studies were performed according to the principles of the Declaration of Helsinki and was approved by the Regional Ethical Review Board in Gothenburg, Sweden: Papers I, II (Diary no:810-11), Paper III, (Diary no: 261-13). All study participants were informed verbally and in writing and signed a consent form. The investigations was performed at the Department of Gastrosurgical Research and Education, at the Sahlgrenska University hospital in Gothenburg, Sweden.

3.2. Subjects and tissue 3.2.1. Subjects

In Paper I and Paper II was used the mucosa of jejunum. Endoscopic jejunal biopsies were taken from healthy volunteers: Paper I (n=28); Paper II (n=16). Inclusion and exclusion criteria were specified for participation in the study. Inclusion criteria: voluntary participation, self-reported general healthy state, age between 18 and 65 years, body mass index between 18 and 25 kg/m2. Exclusion criteria were: overweight or obesity (BMI > 25 kg/m2), history of drug abuse, use of prescription medication within 14 days (with the exception of contraceptives), pregnancy or breast feeding, or potentially childbearing women not using adequate birth control (e. g. intrauterine device, barrier method, oral contraceptive, abstinence), in the investigator’s judgement, clinically significant abnormal-lities at the screening examination or in the laboratory test results.

The muscle tissue of jejunum was used in Paper III. Jejunal tissue of full wall thickness was taken from patients (n=23) undergoing gastric by-pass surgery for morbid obesity. Two of these 23 patients were taking medi-cine for hypertension.

3.2.2. Tissue handling

Tissue collection, handling and preparation were performed in a stan-dardized manner, e. g. refrigeration, transportation to the laboratory, buffer preparations and tissue stripping. On collection, the tissue specimens were immediately placed in fixation medium, Krebs solution or liquid nitrogen depending on the subsequent analysis method.

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3.2.3. Jejunal mucosa

Jejunal biopsies were taken from healthy volunteers in the fasting state. After concious sedation with midazolam and alfentanil, an enteroscope (i.e. a thin-calibered pediatric colonoscope) was introduced into the gastroduo-denum and proximal jejunum. Eight to twelve (Paper I) and eight to ten (Paper II) biopsies were harvested in the jejunum approximately 50 cm distal to the ligament of Treitz. Four to six jejunum biopsies were either snap frozen or chemically fixated for later WB, Immunohistochemistry, EIA analyses. The remaining biopsies were prepared for functional assessments in mini-Ussing chambers.

3.2.4. Jejunal muscle tissue

In Paper III was used jejunal muscle tissue. A full-wall specimen was resected from the jejunum between the gastro-entero and the entero-entero anastomosis as the loop was divided to create a Roux-en-Y construction as already reported by Spak [155]. During contractile experiments in vitro muscular layer ofjejunum tissue were separated from the mucosa/submu-cosa and snap frozen or chemically fixated for later Western blot analyses (WB), Immunohistochemistry or enzyme immunoassay (EIA).

3.3. Western blot

The quantitative amounts of the specific proteins were estimated by the antibody-based method Western blot (WB; immunoblotting), which was used in all papers. Tissue specimens were first solubilized by sonication with ultrasound by sonication with ultrasound energy in PE buffer (10 mM potassium phosphate buffer, pH 6.8 and 1mM EDTA) containing proteinase blockers (3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate). After centrifugation (10,000 × g, 10 min at 4°C) cell debris was removed and the supernatant was analysed for protein content by the method of Bradford [22]. The samples were diluted in SDS (sodium dodecyl sulfate buffer), and heated at 70°C for 10 min before loaded onto a NuPage 10% Bis-Tris gel, and the proteins were separated according to size by gel electrophoresis in an electrical field. After the electrophoresis, the proteins on the gel were transferred (blotted) to a polyvinyldifluoride transfer membrane, Hybond, 0.45 µm, RPN303F to make the proteins detectible to the antibodies. The membrane was then incubated with the specific primary antibody and with a secondary antibody to enhance the signal (see Tables 3.3.1, 3.3.2). The secondary antibody was conjugated with alkaline pho-sphatase that, upon reaction with the reagent CDP-Star, that in turn

THE FUNCTIONAL CONSIDERATION OF RENIN ANGIOTENSIN SYSTEM IN THE HUMAN JEJUNUM

Mantas Malinauskas