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36: Bile Esculin - Biology

36: Bile Esculin - Biology


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Learning Objectives

  • Identify the organism's ability to grow in high bile concentration and to use the sugar esculin

BILE ESCULIN

Bile esculin agar comes in a slant form. It is 2 tests in one---resistance to 40% bile AND the use of the sugar esculin. Bile is a very inhibitory chemical, particularly at that concentration. IF the bacterium grows, the next question is whether it uses the esculin.

If your bacterium cannot grow in the presence of bile, the esculin reaction will have to be obtained a different way, using an esculin sugar disc

MATERIALS NEEDED

  • 1 bile esculin slant per unknown

THE PROCEDURE

  1. Using your unknown, streak the inoculum up the slant.
  2. Incubate the medium at 25º C or 37º C.

INTERPRETATION

  1. Growth on the slant = resistance to 40% bile
  2. Use of esculin sugar = slant turns black coffee (over 1/2 of slant or entire agar) color

QUESTIONS

  1. The presence of growth on bile esculin means __________.
  2. What is esculin?

25 Hydroxycholesterol

Henriikka Kentala , . Vesa M. Olkkonen , in International Review of Cell and Molecular Biology , 2016

1.3 Lipid Ligands of ORPs

OSBP, the archetype of mammalian ORPs, binds 25-hydroxycholesterol (25OHC) within its ORD lipid-binding pocket with a Kd of 10 nM, and other oxysterol species with lower affinity ( Taylor et al., 1984 Dawson et al., 1989 Wang et al., 2002 ). Importantly, OSBP also binds cholesterol with a Kd of 173 nM ( Wang et al., 2005, 2008 ). The close homolog of OSBP in ORP subfamily I, ORP4L, was similarly shown to bind 25OHC with a Kd of 10 nM ( Wang et al., 2002 ), and experiments with the ORP4S variant suggested that ORP4 also binds cholesterol at a lower affinity ( Wyles et al., 2007 ). In ORP subfamily II, ORP1L binds 25OHC (Kd 97 nM), 24(S)OHC, 22(R)OHC, and 7-ketocholesterol (7KC) at relatively high affinity ( Suchanek et al., 2007 Vihervaara et al., 2011 ), while ORP2 shows a high affinity (Kd 14 nM) for 22(R)OHC ( Hynynen et al., 2009 ) but only low affinity (Kd 3.9 μM) for 25OHC ( Suchanek et al., 2007 ), demonstrating that even closely related family members can display distinct differences in ligand specificity. Similar to OSBP and ORP4L, several other ORPs are suggested to bind cholesterol ( Hynynen et al., 2009 Ngo and Ridgway, 2009 Vihervaara et al., 2011 Nissilä et al., 2012 Liu and Ridgway, 2014 ) In fact, recombinant ORP9L failed to bind oxysterols but did bind cholesterol in vitro ( Ngo and Ridgway, 2009 ), and is able to extract cholestatrienol and PI4P from liposomes ( Liu and Ridgway, 2014 ). Similarly, we could not detect oxysterol binding in vitro by ORP10 ( Nissilä et al., 2012 ), which was later on suggested to bind phosphatidylserine (PS) ( Maeda et al., 2013 ). In a study employing photocrosslinkable radiolabeled cholesterol and 25OHC, we observed a positive crosslinking signal for one or both of these sterols for a total of 10 human ORPs ( Suchanek et al., 2007 ). It remained, however, somewhat unclear whether all the observed signals reflected insertion of the labeled sterol derivatives within the ORD ligand cavity. Some of them may have arisen through crosslinking to the surface of the membrane-associated proteins.

Structural studies on the yeast Osh proteins have markedly advanced our knowledge of ORP liganding by lipids. The report by Im et al. (2005) revealed the high-resolution structure of the “short” yeast ORP Osh4p (also known as Kes1p) with five different sterols: ergosterol, cholesterol, and 7KC, 20- and 25OHC. Of importance, de Saint-Jean et al. (2011) determined the structure of Osh4p with PI4P inserted within the ORD ligand cavity, and showed that a bound sterol is readily exchanged for PI4P.

The structure of the Osh3p ORD with bound PI4P ( Tong et al., 2013 ) revealed that the ligand cavity of this protein appears too narrow to accommodate the bulky sterol molecules, suggesting that not all ORPs have the capacity to bind sterols. The authors indicated that the amino acid residues lining the inositol-4-phosphate-binding cleft at the entrance of the ligand cavity are highly conserved among all ORPs, including residues of the “OSBP fingerprint” sequence ( Figure 2 (B)). Together with the similar findings by de Saint-Jean et al. (2011) , this observation brought up the possibility that PI4P binding could be a unifying feature of all ORPs, and that only a subset of the family members could additionally bind sterols. A new aspect in ORP ligand specificity was revealed by Maeda et al. (2013) , who demonstrated that yeast Osh6p and Osh7p specifically bind PS. The authors also crystallized Osh6p and modeled PS into the ORD ligand cavity. The head group and the unsaturated sn-2 fatty acyl chain were oriented toward the entrance of the cavity, while a saturated sn-1 fatty acyl chain was inserted toward the bottom of the pocket. Osh6p extracted very little if any ergosterol from membranes, suggesting that it, similar to Osh3p, may be unable to bind sterols, thus putatively belonging to a new, “glycerophospholipid-binding subgroup” of ORPs. The ORD ligands identified for the yeast and mammalian ORPs are summarized in Table 1 .

Table 1 . Ligands identified for the ORDs of human and S. cerevisiae ORPs by structural analyses or in vitro binding/transport assays.

MammalianLigandsReferences
OSBP25OHC and other oxysterols, cholesterol, PI4P Taylor et al. (1984) Dawson et al. (1989) Ridgway et al. (1992) Wang et al. (2005) Wang et al. (2008) Mesmin et al. (2013)
ORP4/OSBP225OHC, 7KC 20OHC, 22(R)OHC, 22(S)OHC, 7OHC, cholesterol Wang et al. (2002) Wyles et al. (2007)
ORP124(S)OHC, 22(R)OHC, 25OHC, 7KC, cholesterol Suchanek et al. (2007) Yan et al. (2007) Vihervaara et al. (2011)
ORP222(R)OHC, 7KC, 25OHC, cholesterol Suchanek et al. (2007) Hynynen et al. (2009)
ORP3?
ORP6?
ORP7?
ORP5PS, dehydroergosterol, cholesterol? Maeda et al. (2013) Du et al. (2011)
ORP8Cholesterol, 25OHC? Zhou et al. (2011) Yan et al. (2008)
ORP9Cholesterol, PI4P Ngo and Ridgway (2009) Liu and Ridgway (2014)
ORP10PS, cholesterol? Maeda et al. (2013) Nissilä et al. (2012)
ORP11?
S. cerevisiae
Osh1pCholesterol? Schulz et al. (2009)
Osh2pCholesterol Schulz et al. (2009)
Osh3pPI4P, cholesterol? Tong et al. (2013)
Osh4pErgosterol, cholesterol, 7OHC, 20OHC, 25OHC, dehydroergosterol, PI4P, PI(4,5)P2?, PS? Im et al. (2005) Raychaudhuri et al. (2006) de Saint-Jean et al. (2011)
Osh5pCholesterol Schulz et al. (2009)
Osh6pPS Maeda et al. (2013)
Osh7pPS Maeda et al. (2013)

Ileal bile acid transporter inhibition as an anticholestatic therapeutic target in biliary atresia and other cholestatic disorders

Biliary atresia is a rare cholestatic liver disease that presents in infants and rapidly advances to death in the absence of intervention. As a result of blockage or destruction of the biliary tract, bile acids accumulate and drive inflammation, fibrosis, and disease progression. The standard of care, Kasai portoenterostomy (KPE), is typically performed shortly after diagnosis (currently at

2 months of age) and aims to restore bile flow and relieve cholestasis. Nevertheless, most patients continue to experience liver injury from accumulation of bile acids after KPE, since there are no known effective therapeutics that may enhance survival after KPE. Improving cholestasis via interruption of the enterohepatic circulation of bile acids may directly attenuate hepatic bile acid retention and reduce the risk of early organ failure. Directly addressing intrahepatic accretion of bile acids to avoid inherent bile acid toxicities provides an attractive and plausible therapeutic target for biliary atresia. This review explores the novel therapeutic concept of inhibiting the sole ileal bile acid transporter (IBAT), also known as ASBT (apical sodium-bile acid transporter, encoded by SLC10A2), as a means to reduce hepatic bile acid concentration after KPE. By reducing return of bile acids to the cholestatic liver, IBAT inhibitors may potentially lessen or delay liver damage associated with the hepatotoxicity and cholangiopathy of bile acid accumulation. The clinical programs of 2 IBAT inhibitors in development for the treatment of pediatric cholestatic liver diseases, maralixibat and odevixibat, are highlighted.

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Introduction

Bile acids are biologically important cholesterol derivatives produced in the liver and secreted into the bile as glycine- or taurine-conjugated primary bile acids (Chiang, 2009). The main primary bile acids produced are cholic acid (CA) and chenodeoxycholic acid (CDCA) in humans, and CA and muricholic acid (MCA) in mice (Hofmann, 1999 Russell, 2003). The conjugated forms of these primary bile acids are released through the bile duct into the duodenum, where they act as detergents for the emulsification and absorption of dietary lipids (Hofmann, 2009). Beyond their digestive function, bile acids also act as hormone-like regulators of host metabolism (Kuipers et al., 2014) as well as antimicrobials (Begley et al., 2005). Their potency is influenced by several microbial transformations during intestinal transit.

In the small intestine, primary bile acids are deconjugated by bile acid hydrolases which are expressed by a wide variety of intestinal bacterial taxa including species belonging to the Bacteroidetes (Narushima et al., 2006), a dominant group within the gut microbiota. About 95% of the primary bile acids are reabsorbed in the small intestine and recycled via the enterohepatic circulation (Russell, 2003 Ridlon et al., 2005 Hofmann, 2009). The remaining 5% that escape the enterohepatic circulation undergo further microbial transformations in the large intestine (Ridlon et al., 2005, 2016). One of the most important bile acid transformations is 7α-dehydroxylation, generating secondary bile acids, including deoxycholic acid (DCA) and lithocholic acid (LCA). This multistep biochemical pathway, encoded in the bai (𠇋ile acid inducible”) gene operon, is restricted to a narrow phylogenetic group of bacterial species belonging to the Clostridium cluster XIV and has been most extensively studied in the species Clostridium scindens and Clostridium hylemonae (Ridlon et al., 2005, 2016).

Transformation of host-produced conjugated primary bile acids to DCA and LCA is an example of microbial inter-species co-metabolism. 7α-dehydroxylation bacteria depend on other microbial species for deconjugation of the glycine- or taurine-conjugated bile acids, as they can only metabolize unconjugated bile acids. For this reason, bile acid 7α-dehydroxylation by human 7α-dehydroxylating species C. scindens and C. hylemonae cannot occur in mono-colonized animals, in absence of bile acid deconjugating organisms (Narushima et al., 1999).

Secondary bile acids play a key role in resistance to intestinal infection (Begley et al., 2005). The best-studied example is Clostridium difficile infection (CDI), a nosocomial, antibiotic therapy-associated diarrheal infection (Rupnik et al., 2009). Resistance against CDI conferred by a healthy gut microbiota is abolished when perturbed (dysbiosis) by broad-spectrum antibiotic treatment (Britton and Young, 2012). The antibiotic-induced dysbiosis is associated with the abolition of secondary bile acid biosynthesis in the colon (Antunes et al., 2011 Theriot et al., 2016, 2014).

It is well-established that bile acids regulate C. difficile endospore germination and outgrowth as well as vegetative growth (Wilson et al., 1982 Wilson, 1983). Whilst the germination receptors of C. difficile are only partially identified (Francis et al., 2013a Fimlaid et al., 2015), the primary bile acids taurocholic acid (TCA) and cholic acid as well as the secondary bile acid DCA are proven germinants of C. difficile spores (Sorg and Sonenshein, 2008). Moreover, chenodeoxycholic acid (CDCA) as well as alpha and beta stereoisomers of muricholic acid (MCA) are known to inhibit C. difficile germination (Sorg and Sonenshein, 2010 Francis et al., 2013b). Further, DCA as well as LCA, ursodeoxycholic acid (UDCA) and ω-muricholic acid (ω-MCA) are strong inhibitors of spore outgrowth and vegetative cell growth of C. difficile (Francis et al., 2013b Weingarden et al., 2015). Thus, it has been hypothesized that bile acid deconjugating and 7α-dehydroxylating microbial species play a critical role in conferring colonization resistance to C. difficile. Initial evidence supporting this hypothesis was mainly derived from in vitro experiments direct evidence for the importance of microbial bile acid transformation in CDI in vivo is still scarce. Only recently, Buffie and colleagues, by using metagenomic analyses and computational modeling of antibiotic-treated animal and patient samples, were able to infer a strong positive correlation between the presence of C. scindens-related taxa and CDI resistance, and successfully tested causality in vivo by inoculation of antibiotic-treated, CDI-susceptible mice with the C. scindens type strain ATCC35704 (Buffie et al., 2014).

The intestinal microbiology of C. scindens and related species is still understudied. A limitation of previous work has been that C. scindens colonization experiments in vivo required antibiotic-treated animals containing a dysbiotic microbiota with unclear background levels of endogenous bile acid 7-dehydroxylating bacteria. Although this situation mimics the conditions in the CDI patient, it can be an experimental limitation for the study of C. scindens physiology in the healthy gut.

Whilst it is standard to use genetically defined inbred mouse strains for most experimental animal studies, conventional animal models contain still rudimentarily defined, and unstandardized microbial consortia. These consist of hundreds to thousands of different operational taxonomic units and vary considerably between individual animals as well as among research facilities (Rogers et al., 2014 Ericsson et al., 2015 Hoy et al., 2015). Although modern analytical and computational tools facilitate the analysis of complex host–microbiota interactions, it is a challenge to establish the underlying signaling and metabolic networks without the ability to standardize also the microbiota composition. Experimental animal models with a fully standardized microbiota, so-called gnotobiotic animal models, provide a defined and reduced complexity in vivo system for the in-depth elucidation of host–microbiota interactions.

In the present study, we used a recently established defined mouse intestinal microbial consortium, referred to as “oligo-mouse microbiota” (Oligo-MM 12 ), rationally assembled of 12 mouse-intestinal bacterial isolates representative of the major mammalian intestinal bacterial phyla (Brugiroux et al., 2016) as a platform for the in vivo study of C. scindens. The bacterial strains combined in the Oligo-MM 12 consortium were recently described in detail and are openly available from the DSMZ repository (Lagkouvardos et al., 2016). Gnotobiotic Oligo-MM 12 -associated mice (henceforth referred to as “stable defined moderately diverse microbiota, murine 2,” short sDMDMm2 mice) were originally generated by colonization of germ-free mice with a cocktail of cultured bacteria. They have been stably maintained by breeding under gnotobiotic conditions for over 25 generations in three independent facilities without significant compositional deviations, providing mice harboring a physiologically acquired, and developed stable gut microbiota.

We amended the Oligo-MM 12 microbiota by colonization of gnotobiotic sDMDMm2 mice with C. scindens ATCC35704 and assayed the resulting bile acid 7α-dehydroxylation activity and protection against CDI in vivo. For this purpose we first compared the CDI-susceptibility of sDMDMm2 mice with germ-free and antibiotic treatment mouse models of CDI and carried out a comprehensive quantitative analysis of the associated bile acid metabolomes. We then analyzed the effects of the introduction of C. scindens into the preexisting stable sDMDMm2 consortium. We show that C. scindens strain ATCC35704 can be functionally amended to sDMDMm2 to carry out CA and CDCA 7α-dehydroxylation. In correlation with this finding, C. scindens colonization specifically delayed intestinal overgrowth of C. difficile and concomitant intestinal pathology. Only minor direct effects of C. scindens on the composition of the 12 species of the Oligo-MM 12 model consortium were detectable.


The Making of Barrett’s Metaplasia

Most, if not all, esophageal adenocarcinomas arise from Barrett’s esophagus, the condition in which the normal squamous cells lining the distal esophagus are replaced by intestinal-type columnar cells 7 . Barrett’s esophagus develops through the process of metaplasia, the replacement of one adult cell type by another. Metaplasia is thought to arise as a protective response to chronic tissue inflammation 8 , which in the esophagus is thought to be due to GERD. Barrett’s metaplasia can result from either changing fully differentiated esophageal squamous cells directly into intestinal-type columnar cells or from changing the differentiation pattern of esophageal stem cells 8 .


METABOLISM OF LIPIDS - PowerPoint PPT Presentation

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The Systematic Review of Proteins Digestion and New Strategies for Delivery of Small Peptides

4 Faculty of Animal and Veterinary Sciences, Shabestar Branch, Islamic Azad University, Shabestar, Iran.

*Corresponding Author: Tohid Vahdatpour
Faculty of Animal and Veterinary Sciences
Shabestar Branch, Islamic Azad University
Shabestar, Iran.
Tel: +984114777882
E-mail: [email protected]

Received date: March 16, 2016 Accepted date: May 16, 2016 Published date: May 23, 2016

Citation: Vahdatpour S, Mamaghani AP, Goloujeh MS, et al. The Systematic Review of Proteins Digestion and New Strategies for Delivery of Small Peptides. Electronic J Biol, 12:3

Abstract

The digestive system is rich in different proteolytic enzymes, acid, and other secretions, also lined with a layer of mucus, which provides extracellular and intracellular barriers to peptide absorption in all epithelial surfaces of lumen. However, small peptides may be crossing from these barriers and absorbed from different regions of the intestine. These peptides as small, mini or small peptides called bioactive peptides. Bioactive peptides have defined as peptides with quasi-hormone or drug like activity that eventually modulate physiological function through binding interactions to specific receptors on target cells. Bioactive peptides may be classified as antimicrobial, anti-diabetic, antithrombotic, antihypertensive, opioid, immune modulator, mineral binding and anti-oxidative. Most of the final protein digestive products that are absorbed are individual amino acids, with lower values absorption of small peptides. The peptide transporter 1 (PEPT1) is primarily responsible for the absorption of dietary diand tripeptides from the small intestinal. Currently, there is an interest in small peptides in pharmaceutical research and developments are being evaluated in clinical trials. The aim of this review is to summarize the existing knowledge for a better understanding of the challenges about protein digestion, small peptide absorption and oral delivery enhancers with an emphasis on small peptides.

Keywords

Barrier Digestive Peptide PEPT1.

1. Introduction

Almost all proteins and many peptides compound digested in alimentary system. Therefore, pharmacologically active proteins and peptides like hormones cannot be administered orally because of inadequate oral bioavailability, and this may limit the usefulness of these compounds. Lower gastrointestinal bioavailability can be caused by weak aqueous solubility, degradation within the gastrointestinal contents, low membrane permeability or pre-systemic metabolism. Compounds can have poor membrane permeation due to largemolecular weight as is the case with proteins and other macromolecules, or insufficient lipophilicity to partition into biological membranes, as with many hydrophilic low-molecular weight compounds. There are numerous pharmacologically effective compounds currently used that must inject because of inadequate bioavailability by non-injecting routes. Absorption enhancement is the technology aimed at non-injectable delivery of poorly membranepermeable compounds. Consumption of the peptides and proteins show several advantages as compared to conventional drugs. These include high activity, high specificity, low toxicity, and minimal nonspecific and drug-drug interactions [1]. Developments of biotechnology resulted in production of peptides. Oral administration route has advantages, including: patient compliance, ease of administration and reasonably low cost of production. Low oral bioavailability of macromolecular drugs stems mainly from pre-systemic enzymatic degradation and poor penetration across the intestinal membrane. The present review summarizes the physiological barriers to oral delivery of peptides and provides novel pharmaceutical approaches to improve oral bioavailability of bioactive peptides.

2. Digestion of Protein

2.1 Digestion in the stomach

The Pepsin, an important peptic enzyme of the stomach, is most active at a pH of 2.0 to 3.0. The gastric glands secrete a large quantity of hydrochloric acid. This hydrochloric acid is secreted by the parietal (oxyntic) cells in the glands and the pH averages around 2.0 to 3.0, a highly favourable range of acidity for pepsin activity [2]. Pepsin is digesting the protein collagen. Collagen is a major constituent of the intercellular connective tissues therefore, for the digestive enzymes to penetrate meats and digest the other meat proteins, it is first necessary that the collagen fibers be digested. Pepsin only initiates the process of protein digestion, usually providing only 10 to 20 percent of the total protein digestion to convert the protein to proteases, peptones and a few polypeptides. This splitting of proteins occurs as a result of hydrolysis at the peptide linkages between amino acids which are digested to the final stage to form single amino acids and little small peptides. More than 95 percent of the final protein digestive products that are absorbed are individual amino acids, with only 5 percent absorption of di- and tripeptides and very rare absorption of other small peptide molecules. Even these very few absorbed molecules of whole peptides and/or protein can sometimes cause serious allergic or immunologic disturbances [3].

2.2 Digestion in the intestine

Most protein digestion occurs in the initial of small intestine, in the duodenum and jejunum, by proteolytic enzymes from pancreatic secretion. In the small intestine the partial breakdown products of the protein foods attacked by major proteolytic pancreatic enzymes: trypsin, chymotrypsin, carboxypolypeptidase and proelastase. The trypsin and chymotrypsin split protein molecules into small polypeptides carboxy polypeptidase then cleaves individual amino acids from the carboxyl ends of the polypeptides. Proelastase, in turn, is converted into elastase, which then digests elastin fibers that partially hold meats together. The small percentages of the proteins are digested all the way to their constituent amino acids by the pancreatic juices [4]. Therefore, most remain as dipeptides and tripeptides. The last digestive stage of the proteins in the intestinal lumen is achieved by the enterocytes that line the villi of the small intestine, mainly in the duodenum and jejunum. These cells have a brush border that consists of hundreds of microvilli projecting from the surface of each cell. Two types of peptidase enzymes are important, aminopolypeptidase and several dipeptidases that succeed in splitting the remaining larger polypeptides into tripeptides and dipeptides and a few into amino acids. The amino acids with the dipeptides and tripeptides are easily transported through the microvillus membrane to the interior of the enterocytes [5].

3. Absorption of Peptides

Therefore, peptides for absorption must firstly diffuse across the mucus layer before absorption across the epithelia is possible. The aqueous boundary or unstirred water layer can be act as a limiting factor for highly lipophilic peptides. Once a protein crosses the monolayer of intestinal epithelial cells, it can enter either the capillaries of the portal venous system or the lymphatic lacteal [6]. The lipophilic peptides are more likely to be absorbed by the lymphatic system [7]. The lymphatic circulation bypasses the liver and thus the attractive approach to delivery of peptides and proteins. Absorption into the lymphatic lacteals provides very slow systemic delivery over several hours as the lymph moves at a slow rate. Although, absorption into the portal venous system results in rapid delivery within minutes to systemic circulation after an initial hepatic pass.

PEPT1 mediated active absorption is responsible for the high bioavailability of orally active peptides so that the proton-coupled uptake of the more than 8000 different di- and tripeptides is performed by the high-capacity/low-affinity peptide transporter isoform PEPT1 (SLC15A1) [8]. For examples: betalactam antibiotics of the cephalosporin and penicillin classes, ACE-inhibitors, ester prodrugs of enalapril and fosinopril, bestatin, alafosfalin, amino acidconjugated antiviral drugs (valacyclovir), L-DOPA, as well as artificial di- and tripeptides such as Gly- Sar [9]. PEPT1 prefers peptides containing L amino acids over D enantiomers. Several PEPT1 inhibitors such as sulfonylurea antidiabetic drugs, nateglinide, glibenclamide, tolbutamide, chlorpropamide, sartans and ester pro-drugs of ACE inhibitors have been identified. Di-peptides such as Gly-Sar and Val- Ala have also demonstrated inhibitory potential therefore di- and tri-peptides produced by digestion of milk proteins may also inhibit PEPT1-mediated drug absorption, causing reduced exposure of the victim drug (Figure 1).

4. Barriers to Peptide Absorption

Barriers to peptides absorption are extracellular and intracellular barriers. Potent barriers exist to the oral absorption of peptides the development of oral replacements for injectable peptides is a high-priority research for the consumption of peptides [10]. The oral bioavailability of the most peptides is less than one percent. Inhibition of proteolytic enzymes and opening of tight junctions increase para-cellular transport of peptides from intestine [11].

4.1 Extracellular barriers

Peptide absorption from the digestive system is encountering with enzymatic and penetration barriers. Hydrolysis of peptides and proteins in the digestive system can occur in the lumen, in the brush border, intracellular fluid or in the cytosol of the enterocytes [12]. As the protein is ingested, it reaches the stomach, where it is acted on by gastric juice in a very acidic environment. Gastric juice contains a family of aspartic proteinases called pepsins. Pepsins digested the protein into a mixture of polypeptides and move down to the duodenum. As the protein enters the duodenum, the pH rises to about 6.0 to 8.0. This pH change approximately from 2.0 to 8.0 can also cause precipitation of the protein through its iso-electric point and then may not easily re-dissolve [4].

Figure 1:Digestion of proteins and peptides for absorption of amino acids and di- tri- peptides to blood stream.

In the duodenum, the polypeptides are acted by pancreatic proteases. These proteases are classified to endopeptidases such as trypsin, chymotrypsin, and elastase or exopeptidases such as carboxy peptidase A. These enzymes degrade the polypeptides into smaller peptides. These peptides are then further degraded by the proteases in the brush border and the cytosol of the enterocytes. For peptides with four or more residues, more than 90 percent of the proteolytic activity is in the brush border membrane. For tripeptides, the activity is 10 to 60 percent for dipeptides, it is only about 10 percent. Lysosomes and other cell organelles can also act as potential sites of peptide and protein degradation [13,14]. The brush border contains exopeptidases that act at the N-terminal end of the protein (amino peptidases). Amino peptidases in the intestine include leucine amino peptidase and amino peptidases N, A, and B. The amino peptidase activity in the Payer's patches of the jejunum and ileum is only about 20 to 30 percent of that in the neighboring patch-free areas [14]. Knowledge of the distribution of brush border membrane peptidases along the intestine helps to predict the preferential uptake of peptides and proteins from various intestinal regions [13]. Because this distribution may vary for different enzymes, sitespecific oral delivery may be dependent on the amino acid sequence of the peptide. This is because of the substrate specificity of the brush border membrane peptidases. Also, the activities of brush border membrane peptidases may be controlled by the surface pH of mucosal cells rather than the luminal pH [15]. Therefore, it should be realized that several enzymes that act on carbohydrates may also affect the drug if it is a glycoprotein.

4.2 Intracellular barriers

Peptides can move from epithelia by two routes. The transcellular route involves intracellular transfer from the apical to the basolateral surface of an individual epithelial cell. This transport can take place either through specific uptake mechanisms of the cell or through sequential partitioning, events from an aqueous environment to a hydrophobic environment and then back into an aqueous environment. The transcellular route is important for the uptake of lipophilic peptides by sequential partitioning events. The carbohydrates and amino acids are transported by a carrier-mediated process. Although L-amino acids are absorbed by an active transport mechanism, D-amino acids are absorbed by passive diffusion. There are four separate transport systems for amino acids, and a separate system transports dipeptides and tripeptides into the mucosal cells.

These carrier systems can even transport amino acid-type or peptide-type drugs. The active transport mechanisms for the transcellular route include Na+ coupled glucose transport and H+ coupled dipeptides transport. Studies on the kinetics of glycylproline transport in intestinal brush border vesicles have shown that dipeptides transport is saturable process and follows Michaelis-Menten kinetics [16]. Using brush border membrane vesicles (BBMVs) prepared from rabbit duodenum and jejunum and rat upper small intestine, no evidence was found in the oral absorption of TRH by a Na+ or H+ dependent carrier system in the brush border membrane. Of course, it appears that the TRH absorption in vivo may be accounted for by passive absorption of the peptide across a paracellular route and its resistance to luminal hydrolysis.

The paracellular route involves transfer between adjacent cells. The villous cells have tight intracellular junctions, which prevent paracellular transport of solutes. Movement from this route is limited by the junction complex to molecules with a radius less than about 8 Å [17]. If a peptide can pass from the paracellular route, it will not be subject to digestion by the intracellular proteases. Although the paracellular route may be preferable for this reason, structural features of peptides that may encourage their paracellular transport are not well understood. The partition coefficient between n-octanol and water may be one predictor, and drugs with a log P of less than zero (i.e., hydrophilic molecules such as most peptides) are more likely to follow a paracellular pathway. Peptides following a paracellular pathway may also be more affected by penetration enhancers such as zonula occludens toxin (ZOT), a protein from Vibrio cholera that can reversibly open tight junctions between intestinal cells [18]. Paracellular transport is constrained by the physical properties of permeate. Larger peptides (greater than three amino acids) are usually absorbed in small amounts by passive diffusion via the paracellular route. Although larger proteins are not typically absorbed in the digestive tract, protein antigens can be taken up by M cells, which are specialized intestinal epithelial cells that overlie aggregates of lymphoid tissue (Payer's patches). The tight junctions for paracellular diffusion have been reported to be generally impermeable to molecules with radii of more than 11 to 15 Å, which may represent the limit for the hydrodynamic radius for oral delivery of spherical rigid molecules. However, peptides have some conformation flexibility, and even larger molecule scans permeate the tight junctions [19].

5. The Site of Peptide Delivery

The different peptides absorbed from different regions of the intestine by several manners. This is believed to be caused by decreased proteolytic activity in the distal area. Also, the distal region has higher paracellular permeability despite a decreased absorption area [20]. The protease activity in the cytoplasm does not show regional variation, but the same is not true for the brush border or for the luminal fluid. The stomach, with its low pH and enzymatic activity, presents very harsh conditions for a protein drug. A typical approach to prevent dissolution of a dosage form in the stomach is to use enteric coating.

The intestinal segments have progressively fewer and smaller villi in the more distal sections. This leads to a progressively reduced surface area, with the colon having the lowest surface area for a particular length. The colon also has variable pH and the presence of solid fecal matter, which may interfere with drug absorption. However, the colon has relatively low enzymatic activity and is promising in this regard. Using isolated luminal enzymes and studies in intact mucosa, calcitonin was found to degrade much more in the small intestine compared to the colon [21]. The colon has a high population of bacteria largely anaerobic species. This fact has been exploited for an ingenious approach to target peptides and proteins to the colon [22]. In a study polypeptides such as insulin or vasopressin were coated with polymers cross-linked with azo-aromatic groups to protect orally administered polypeptides from digestion in the stomach and small intestine of rats. Once the polypeptide reached the colon, the indigenous micro flora reduced the azo bonds, thus breaking the cross-links and releasing the polypeptide for absorption. The upper half of the large intestine is drained by hepatic portal veins the lower half is drained by lymphatic. If a polypeptide is destroyed in the liver, it may be possible to adjust the thickness or composition of coating so that the drug is released in the lower colon, where it will bypass the hepatic veins. Delivery of insulin and an absorption promoter to the colon has also been attempted using a soft gelatin capsule coated with a poly acrylic polymer (Eudragit) having pH-dependent properties [23]. Delivery of insulin-like growth factor I (IGF-I) to rat and mini pig colonic mucosa under in vitro conditions has been investigated. IGF-I is a 7649- Da protein of 70 amino acids that exerts its biological actions through specific IGF-I receptors. It has been found useful to lower blood glucose levels in insulinresistant diabetic patients in clinical studies. IGF-I was absorbed intact across the rat colonic mucosa as determined by reverse-phase high-performance liquid chromatography (RP-HPLC), sodium dodecyl sulfate&ndashpoly acrylamide gel electrophoresis (SDSPAGE), and Western blotting [24]. A time-based drug release system for colon-specific delivery has been developed. This system exploits the relatively constant small intestinal transit time of dosage forms [11]. Time-based systems can be designed to release their drug after a predetermined lag time, with the lag time independent of normal physiological conditions such as pH, digestive state of the subject and anatomical position at the time of release [19,25].

The apparent permeability of insulin from rat intestine shows a site-dependent variation as measured by the averted rat gut sac technique. The permeability was significantly greater in the jejunum and the ileum than in the duodenum. In these in vitro experiments, insulin was remarkably stable. This suggests that insulin metabolism at the brush border is not significant. However, insulin was metabolized almost completely in intestinal homogenates. Thus, it appears that degradation of insulin under an in vivo situation would be caused by luminal and cytosol enzymes [1]. In situ experiments have shown that the absolute bioavailability of insulin was higher when administered in the more distal region of the rat intestine than that absorbed from a more proximal region of the intestine [26]. Insulin absorption from isobutyl cyanoacrylate Nano capsules administered to diabetic rats was dependent on the site of absorption. The hypoglycemic effect following absorption from various sites was as follows: ileum, stomach, duodenum and jejunum, and colon.

6. Strategies for Delivery of Peptides

The five main methods are most important strategies, including: chemical modification, bioadhesive delivery systems, penetration enhancers, protease inhibitors, carrier systems. The other formulation and manners referred at the end of this review which will be considered in the near future.

6.1 Chemical modification

The carrier molecules used to reversibly destabilize the native peptides. This no covalent interaction between the carrier and partially unfolded protein conformation increases solvent exposure of hydrophobic side chains, thereby increasing lipid solubility and oral absorption of the protein by a passive and transcellular route [27]. The chemical modification approach is more applicable to peptides than to proteins because of the structural complexity of proteins. A peptide can be chemically modified to improve its enzymatic stability or membrane permeation. For example, substitution of D-amino acids for L-amino acids in the primary structure may improve the enzymatic stability of the peptide. An example of chemical modification of a peptide that results in increased enzymatic stability without affecting membrane permeability is the various analogues of the naturally occurring penta-peptide methionine (Met)-enkephalin. The metabolism of these analogues by BBMVs shows large differences in degradation rates, but they all have similar effective permeability across Caco-2 cells [28].

Another clue to chemical modification comes from the fact that a lipophilic peptide, cyclosporine A, is readily absorbed from the gastrointestinal tract. Thus, efforts have been directed toward imparting lipid solubility to peptides by bonding an acyl group of a fatty acid to an amino terminus of the peptide. Using a range from tripeptides to proteins, lipid solubility was achieved for thyrotropin-releasing hormone, tetra-gastrin, insulin, and lysozyme. These new derivatives maintained their biological activity and had increased absorption from the intestine [29]. Another approach to increase lipophilicity can be cyclization, which will remove charged N- and C-terminal groups, reducing overall solvent-accessible surface area of the molecule. Also, more lipophilic synthetic amino acids such as t-butyl glycine, b-naphthyl alanine, and p-phenyl phenylalanine can be used to synthesize peptide analogues provided biological activity is not lost. The use of a conjugate system, which combines structural features of lipids with those of amino acids and peptides, is likely to provide a high degree of membrane-like character for the conjugate, which may allow its passage across membranes [30]. Chemical modification of salmon calcitonin has also been done to make its oral absorption feasible. In a study modified salmon calcitonin by a new method to prepare fatty acid-polypeptide conjugates it can be carried out in aqueous solutions and can regenerate the original active polypeptide in tissues or blood. Using this reversible aqueous lipidization approach, the area under the curve (AUC) of modified calcitonin delivered orally was about 20 times higher than that of unmodified calcitonin [19]. PEGylated proteins may also have a potential for oral delivery. PEGylation of recombinant human granulocyte colony-stimulating factor has been reported to increase its stability and in vivo bioactivity when administered by the intraduodenal route. Its bioavailability by the internal route was 1.8 to 3.5 percent the unmodified protein did not produce any quantifiable response [31].

Chemical modification does not always lead to improved oral absorption. Diacyl derivatives of insulin exhibited higher proteolysis than native insulin in the small intestine of rat under in vitro conditions. This was because insulin association was inhibited by diacylation, making more monomers available for proteolysis [32]. The structural requirements for intestinal absorption of peptide drugs have been reviewed [23]. Barlow and Satoh [9] conducted a series of elementary analyses to define the basic design features for a potent, specific, and absorbable peptide drug. Recognition of the peptide by its target receptor seems to need about 4 to 6 amino acid residues, and the rest of the structure may be &ldquoredundant&rdquo for bioactivity. The resulting structure is still too big for transport by paracellular transport. Also, peptide transporters for molecules larger than the three residues are unlikely to exist, so that active transport is also not feasible. For transport by simple diffusion, the lipophilicity of the peptide needs to be increased. Based on molecular modeling, it was predicted that an active absorbable peptide should have a total surface area of around 350 Å2, of which the polar surface area should be 50 Å2 or less. This could be attempted by methylation the peptide NH groups, eliminating charged termini or cyclization of the molecule so that peptide CO and NH groups are made inaccessible to solve because of intra molecular hydrogen bonding. Computer simulations to design peptides or to predict their oral absorption may be possible. A theoretical analysis to estimate the extent of peptide absorption has been developed on the basis of a mass balance approach. Using this analysis, simulations showed that the intestinal absorption of insulin is approximately 1 percent of the administered dose [33]. Because some peptide transporters are known to exist in the intestinal mucosa, knowledge of its structure will lead to rational design of peptide mimetic having affinity for this receptor. However, passive diffusion will be limited due to substrate specificity. Structural features of peptides to achieve drug delivery are not necessarily the same as those required for bioactivity. Therefore, a collaborative effort by a multidisciplinary team is required for rational design of peptide mimetic with adequate oral absorption [33].

Briefly, among common standard modifications of peptides as follows:

- Unnatural amino acids (6-Aminocaproic acid, Amino butyric acid, Citrulline, Norleucine, etc.)

- Heavy amino acids (labeled with 13C and/or 15N)

- Phosphorylation or sulfurylation (at Ser, Tyr, Thr)

- Conjugation to carrier proteins (BSA, KLH, OVA)

- Branching of peptides (MAPs &ndash multiple antigenic peptides)

6.2 Bioadhesive delivery systems

Bioadhesive delivery systems have been widely investigated to prepare oral peptide consumption [34]. This increases the overall time for peptide absorption as the delivery system will not be dependent on the gastrointestinal transit time for removal. Peptides will not have to diffuse through luminal contents or the mucus layer to reach the mucosal epithelium. Because of intimate contact with the mucosa, a high drug concentration is presented for absorption. Also, site-specific delivery may be possible if bioadhesion can occur at a particular site in the digestive system. Bioadhesive delivery systems may be affected by the mucus turnover time in the digestive system, which varies based on the site. In the digestive system of rats, the colon and cecum were found to be the best location for mucoadhesion of poly carbophil disks. Mucoadhesive intestinal patches have also been investigated for oral delivery of conventional drug molecules [5]. Bioadhesive polymers can be used to improve the oral absorption of peptide drugs. The Bioadhesive polymers, poly carbophil, and chitosan derivatives have been used to enhance the absorption of the peptide drug 9-desglycinamide, 8-argininevasopressin (DGAVP) in the vertically perfused intestinal loop model of the rat [35]. Buccal adhesive systems offer innumerable advantages in terms of accessibility, administration and withdrawal, receptivity, low enzymatic activity, economy and high patient compliance. Adhesions of these drug delivery devices to mucosal membranes lead to an increased drug concentration gradient at the absorption site and therefore improve bioavailability of systemically delivered drugs. Investigations are continuing beyond traditional polymer networks to find other innovative drug transport systems. In the current global scenario, scientists are finding ways to develop buccal adhesive systems through various approaches to improve the bioavailability of drugs used orally by manipulation of the formulation strategies like the inclusion of pH modifiers, enzyme inhibitors, permeation enhancers, etc. The future direction of buccal adhesive drug delivery lies in vaccine formulations and delivery of peptides.

Another important aspect concerns the in vitro and ex vivo techniques which are employed for evaluation of the performance of the materials and dosage forms. Important factors affecting mucoadhesion including:

o Concentration of active polymer

o Chain flexibility of polymer

o Functional Group Contribution

&bull Environmental &ndash Related Factors: 21-25

o Selection of the model substrate surface

6.3 Penetration enhancers

Recently, the peptides are considered as penetration enhancers like skin penetrating peptides (SPPs) and/ or cell penetrating peptides (CPPs) have garnered wide attention in recent years and emerged as a simple and effective non-invasive strategy for macromolecule delivery into the skin or cells, respectively. Generally, penetration enhancers can improve oral absorption by their action on the transcellular or paracellular pathway. For effects on the transcellular pathway, surfactants and fatty acids may alter the membrane lipid organization and may thus increase oral transport. Surfactants can be incorporated into lipid bilayers, thus changing the physical properties of the cell membranes. For effects on the paracellular pathway, chelating agents can disrupt the integrity of occluding junctional complexes by chelating calcium or magnesium around tight junctions [36]. Bile salts such as sodium deoxycholate and sodium cholate can also be used in the formulation to promote the absorption of insulin from the colon mixed micellar systems have also been used to enhance the oral absorption of polypeptides [37]. Such systems are also known to form naturally in the gastrointestinal system to aid the absorption of lipids. The dietary fats are first emulsified by bile salts in the intestine and then acted on by pancreatic lipase to produce mono glycerides and free fatty acids. Lipoidal dispersions of insulin in fatty acids using sodium glycol cholateas an emulsifier and absorption promoter have been investigated. The hypoglycemic effects after oral administration to rabbits were found to be dependent on the fatty acid used. Penetration enhancers may enhance the absorption of drugs preferentially in some specific region of the gastrointestinal tract. Cyclo-dextrins have also been used to enhance the absorption of insulin from the lower jejunal/upper ileal segments of the rat by an in situ closed-loop method.

When penetration enhancers are used to enhance oral absorption, it could be realized that they have limitations that may prevent their general acceptance for usage. Also, the potential lack of specificity of penetration enhancers may have long-term toxicity implications that can only be evaluated in chronic studies. The potential lytic nature of surfactants raises safety concerns because the intestinal epithelium provides a barrier to the entry of toxins, bacteria, and viruses. Similarly, chelators that cause Ca2+ depletion do not act specifically on tight junctions but rather may induce global changes in cells, such as disruption of actin filaments or adherent junctions. Thus, it will be difficult to induce the opening of tight junctions in a rapid, reversible, and reproducible manner [37].

6.4 Protease inhibitors

Protease inhibitors may also promote oral absorption of therapeutic peptides and proteins by reducing their proteolytic breakdown in the gastrointestinal tract. Generally, inhibitory agents may be classified as (1) polypeptide protease inhibitors (e.g., aprotinin) (2) peptides and modified peptides (e.g., bacitracin, chymostatin, and amastatin) (3) amino acids and modified amino acids (e.g., a-aminoboronic acid derivatives), and (4) others (e.g., p-aminobenzamidine and camostat mesilate) [38]. An amino peptidase inhibitor, amastatin, has been reported to reduce the hydrolysis of the penta peptide, leucine (Leu)-enkephalin (YGGFL) at a high pH. At lower pH (below 5.0), the endopeptidase inhibitors, tripeptides YGG and GGF, were found to be effective. Coperfusion of YGGFL with a combination of amino- and endopeptidase inhibitors was most effective to inhibit hydrolysis in the rat jejunum. In the absence of these inhibitors, extensive hydrolysis of YGGFL was observed in the rat jejunum, primarily by brush border enzymes and secondarily by luminal peptidases [39]. In another study, an aminopeptidase inhibitor (puromycin) was able to increase the absorption of metkephamid (MKA), a stable analogue of Met-enkephalin, across the rat intestine. However, in this study, an endo-peptidase inhibitor (thiorphan) was ineffective. This is because the dominant enzyme participating in MKA metabolism during absorption is amino-peptidase [40].

Bile salts, in addition to acting as penetration enhancers, can also act as protease inhibitors to enhance oral absorption. Bile salts have been shown to inhibit brush border membrane and cytosolic proteolytic hydrolysis and would thus be useful to reduce intestinal degradation of peptide drugs [10]. A bacterial protease inhibitor from Brucella abortus called U-Omp19 has been reported as an ideal constituent for an oral vaccine formulation against infectious diseases. When U-Omp19 was coadministered orally with Toxoplasma gondii antigen (Ag), U-Omp19: i) could bypass the harsh environment of the gastrointestinal tract by inhibiting stomach and intestine proteases and consequently increased the half-life of the co-administered Ag at immune inductive sites. Finally, this bacterial protease inhibitor in an oral vaccine formulation conferred mucosal protection and reduced parasite loads after oral challenge with virulent Toxoplasma gondii [41]. In an in situ study with closed small and large intestinal loops in rats, no marked hypoglycemic response was observed when insulin alone was administered. However, a significant hypoglycemic effect was obtained following large intestinal administration of insulin with 20 mM sodium glycocholate, camostatmesilate, and bacitracin [42]. It has been suggested that if a protease inhibitor such as soybean trypsin inhibitor is used to prevent the proteolysis of insulin in the rat intestine, then its absorption is promoted by the endogenous bile acids present in the intestine [43]. In another study a decrease in insulin degradation with the co-administration of protease inhibitors to improve the oral bioavailability of insulin has been reported. A significant decrease of blood glucose levels in both lean and diabetes induced obesity rat models as well as a significant increase in plasma insulin levels 20 min and 135 min post-administration of oral insulin with the peptidase inhibitor have been shown in this study [44]. Very small doses (about 1 mg) of vasopressin in solution produced anti diuresis in rats following oral administration. The biological response was enhanced for AVP and LVP by the simultaneous administration of 1000 units of aprotinin, a protease inhibitor. The synthetic analogue DDAVP was more active than the natural hormones, but the effect of aprotinin with DDAVP was inconsistent. The relatively greater oral activity of DDAVP is caused by the unnatural D-arginine, which makes it resistant to attack by trypsin. Starchg- poly (acrylic acid) copolymers and starch/poly (acrylic acid) mixtures have been synthesized and may have potential for enabling oral peptide delivery because of their proteolytic enzyme inhibition activity and ion-binding capacity [20].

6.5 Carrier systems

Carrier systems such as nano-particles, microspheres, liposomes, or erythrocytes can also be used to improve the oral absorption of peptides and proteins. Emisphere Technologies, Incorporated (Tarrytown, NY) initiated clinical trials for oral delivery of insulin using its carrier eligen® technology. The company has also initiated oral delivery of recombinant human growth hormone in collaboration with Novartis and is currently in phase II clinical trials for oral delivery of calcitonin. These carrier molecules, in high concentrations, cause the protein to undergo a conformational change to a partially unfolded or molten globule state that has a higher oral permeability. The carrier molecules are small organic molecules with a molecular weight of about 200 to 400 Da. The protein is used in its native state rather than by a chemical modification approach, and the interaction between carrier and protein is non-covalent. Using cell mono-layers, it has been shown that the tight junctions between cells are not disrupted. The company has also used PYY 3-36 to demonstrate proof of concept for its oral delivery technology. PYY 3-36is a 34-residue gut hormone that physiologically inhibits food intake and has potential for treatment of obesity [12,45]. The use of different carrier systems to improve the absorption of insulin in anesthetized diabetic rats following intraduodenal administration from amid line incision has been evaluated. Several erythrocyte&ndashmembrane carrier systems were tested. These included erythrocyte ghosts (EGs) prepared by hemolysis of human red blood cells, erythrocyte vesicle (EVs) prepared by sonication of EG suspension, and liposome-incorporating ghosts or vesicles (LEGs and LEVs, respectively). Compared to a control group, these carriers enhanced oral absorption of insulin, with LEV the best carrier for more efficient delivery [45].

Uptake of liposomes by Payer's patches can increase the uptake of any entrapped drug. Negatively charged liposomes with at least 25 mol of phosphatidyl serine have been reported to be taken up readily by the rat Payer's patches following intra-luminal administration. Proteins such as albumin have also been used to prepare micro particles to improve the stability of drugs in the gastrointestinal tract. Thermally condensed amino acids (proteinoids) can spontaneously form microspheres when exposed to an acidic medium. Proteinoid microspheres have been used with positive results to deliver encapsulated calcitonin to rats and monkeys. In rats, the serum calcium levels decreased by 23 mg/ml 1 h after dosing encapsulated calcitonin. In contrast, rats receiving control calcitonin (no microspheres) had a decrease of only 6.5 mg/ml [1].

The advantage of using nano-particle formulations over other methods such as liposome formulations is the capability of controlled release in addition to the ability of improving drug stability, absorption and targeting [44]. The absorption and tissue distribution of 14C-labeled poly (D, L-lactide-co-glycolide) nanoparticles after oral administration to the mice has been determined in comparison to the intravenous route. The gastrointestinal transit of the nano-particles was very fast, with most of the radioactivity appearing rapidly in the colon 4 h after administration and in the feces 24 h after administration. Of the amount absorbed through the intestinal barrier (about 2%), most was found in the carcass and liver [16]. Nanocapsules may prefer initially absorbed through the Payer's patches and may be visible in M cells and intercellular spaces around lymph cells. It seems that this uptake by Payer's patches is especially important in the ileum. Absorption of nano-capsules in the jejunum may be by a paracellular pathway, possibly through the intercellular spaces formed by the desquamation of well-differentiated absorptive cells at the tip of the villi [16]. A palmitic ester prodrug of the model drug leucine-5-enkephalin was encapsulated within chitosan amphiphilic nanoparticles. Palmitic acid was used for increasing the lipophilicity of Leucine-5-enkephalin also stabilizing the peptide in the plasma and chitosan amphiphilic nano-particles were used to enhance gastrointestinal uptake. Via the oral route the nano-particle pro-drug formulation increased the brain drug levels by 67% and significantly increased leucine5-enkephalin&rsquos anti-nociceptive activity. The nano-particles facilitated oral absorption and the pro-drug prevented plasma degradation enabling brain delivery [46]. Free insulin did not affect glycemia when administered orally under the same experimental conditions. The intestinal absorption of insulin and calcitonin encapsulated in poly-isobutyl cyano-acrylate nano-particles has been investigated in rats, and the resulting pharmacokinetic profiles were characteristic of sustained delivery. A relatively higher plasma concentration was seen at the later time points, but was balanced by lower initial concentrations thus, there was no significant net enhancement of absorption. This suggests that the nano-capsules slowly released the peptide into the intestinal lumen, with small amounts absorbed [47]. Hydrogel nano-spheres composed of polymetha crylicacid-grafted-poly (ethylene glycol) have also been investigated for oral protein delivery and have been reported to be capable of opening the tight junctions between epithelial cells in Caco-2 cell mono-layers [48]. Thus, our current knowledge provides some promising approaches on how to deliver peptides based drugs not only to the site of disease but also inside the target cell for enhanced therapy. Traditional methods of intracellular delivery, such as electro-portion or microinjection are invasive and applicable for in vitro experiments, but not for clinical conditions. The use of various pharmaceutical nano-carriers, such as liposomes, possessing pH-sensitivity and being able to escape from the endosomes upon the endocytic uptake, or the modification of peptide and protein drugs with cellpenetrating peptides, can allow for efficient and noninvasive intracellular delivery. Although the majority of experiments with pH-sensitive pharmaceutical nano-carriers and cell-penetrating peptide-modified drugs and drug carriers are still in pre-clinical stage, we can expect the appearance of new drugs and treatment protocols based on these methods in the very near future (Figure 2).

6.6 Other formulation

Figure 2:(A) Transport mechanism of biodrug through the intestinal epithelium membrane, (B) Probable mechanism of penetration enhancer, and (C) enzyme inhibitors, (D) Representative mechanism of prodrug absorption and its activation.

Several formulations have been reported for the gastrointestinal absorption of peptides. The oil phase contains a lipid composition similar to those of chylomicrons. A protinin, a protease inhibitor, will prevent peptide degradation chylomicrons will improve absorption into the enterocyte. The emulsion is coated on carrier powders, which are then filled in hard gelatin capsules. The capsules are then enteric coated to prevent dissolution in the stomach. Another approach involves non-covalent linking of the peptide to phospholipids so that the complex can be absorbed into the enterocytes by endocytosis. These approaches are used to develop oral formulations for insulin, calcitonin, porcine somatotropin, erythropoietin, and an interferon [48]. Oral administration of insulin in solid form to nondiabetic and diabetic dogs has been attempted by mixing insulin with cholate and soybean trypsin inhibitor and delivering it orally as enterocoated micro tablets. Following administration of the drug, plasma insulin levels increased and plasma glucose levels decreased after a gap of about 60 to 140 min. Because delivery of insulin by the oral route leads to targeting of the entero-hepatic pathway, the authors of this study felt that this or a similar formulation may serve as an adjuvant treatment for patients with type II diabetes mellitus [34]. He et al. [49] stabilized biocompatible nano-emulsions by food proteins which could deliver fenofibric acid in vivo. In this study Food proteins (soybean protein isolate, whey protein isolate, &beta-lactoglobulin) were used as stabilizers for nano-emulsions to deliver hydrophobic drugs such as fenofibric acid. Food proteinstabilized nano-emulsions, with small particle size and good size distribution, exhibited good stability and bioavailability. The nano-emulsions enable the lipophilic drug to be absorbed more rapidly and better when compared with the oil solution also a much better stability was observed in protein-stabilized nanoemulsions relative to nano-emulsions stabilized with surfactants so Food proteins are viable replacements for traditional surfactants. It should be noted that the bioavailability of SPI-stabilized nano-emulsions were dramatically greater than that of nano-emulsions stabilized by &beta-lg and WPI [49].

7. Conclusion

Proteins and many peptides compound digested in alimentary system. Therefore, active proteins and peptides like hormones cannot be administered orally because of inadequate oral availability. Successful peptide delivery by the gastrointestinal route needs a succession of events to bypass the various penetration or enzymatic barriers at each stage. A site-specific delivery system and approaches to minimize proteolytic degradation are required. The use of penetration enhancers, carrier systems, especially new designed protease inhibitors, or chemical modification of the peptides offers promising approaches to enhance their oral delivery. Designing of absorbable small peptides that penetrate the intestinal mucosa by the paracellular pathway and absorbed to blood seems to be a possible approach. The peptide transporter1 (PEPT1) is primarily responsible for the absorption of dietary di- and tripeptides from the small intestinal lumen. Substrate type interactions by PEPT1 have been successfully exploited with pro-drugs that were designed to introduce peptide- and peptide bond like moieties on the parent molecule. It seems that in the future an important witness will be reported about the valuable effects of small peptides which using with protease inhibitors, and/or chemical modification could be easily absorbed at high levels from the gastrointestinal tract.


Farnesoid X receptor represses hepatic human APOA gene expression

1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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1 Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, and 2 Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University of Graz, Graz, Austria. 3 Department of Cardiology, University of Queensland, Mater Adult Hospital, Brisbane, Australia. 4 Laboratory of Metabolism, National Cancer Institute, NIH, Bethesda, USA. 5 Division of Gastroenterology and Hepatology, Department of Internal Medicine III, Medical University of Vienna, Vienna, Austria.

Address correspondence to: Gert M. Kostner, Institute of Molecular Biology and Biochemistry, Center of Molecular Medicine, Medical University of Graz, 8010 Graz, Harrachgasse 21, Austria. Phone: 43.316.380.4202 Fax: 43.316.380.9615 E-mail: [email protected]

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High plasma concentrations of lipoprotein(a) [Lp(a), which is encoded by the APOA gene] increase an individual’s risk of developing diseases, such as coronary artery diseases, restenosis, and stroke. Unfortunately, increased Lp(a) levels are minimally influenced by dietary changes or drug treatment. Further, the development of Lp(a)-specific medications has been hampered by limited knowledge of Lp(a) metabolism. In this study, we identified patients suffering from biliary obstructions with very low plasma Lp(a) concentrations that rise substantially after surgical intervention. Consistent with this, common bile duct ligation in mice transgenic for human APOA (tg-APOA mice) lowered plasma concentrations and hepatic expression of APOA. To test whether farnesoid X receptor (FXR), which is activated by bile acids, was responsible for the low plasma Lp(a) levels in cholestatic patients and mice, we treated tg-APOA and tg-APOA/Fxr –/– mice with cholic acid. FXR activation markedly reduced plasma concentrations and hepatic expression of human APOA in tg-APOA mice but not in tg-APOA/Fxr –/– mice. Incubation of primary hepatocytes from tg-APOA mice with bile acids dose dependently downregulated APOA expression. Further analysis determined that the direct repeat 1 element between nucleotides –826 and –814 of the APOA promoter functioned as a negative FXR response element. This motif is also bound by hepatocyte nuclear factor 4α (HNF4α), which promotes APOA transcription, and FXR was shown to compete with HNF4α for binding to this motif. These findings may have important implications in the development of Lp(a)-lowering medications.

Lipoprotein(a) [Lp(a)] is a plasma lipoprotein found in humans and Old World monkeys but is absent in conventional laboratory animals. Plasma Lp(a) concentrations are under strict genetic control and range from less than 1 mg/dl to more than 200 mg/dl, with medians of 8 to 9 mg/dl (reviewed in refs. 1 , 2 ). Lp(a) is a complex plasma lipoprotein formed through covalent binding of free APOA, which is synthesized predominantly in the liver, with apoB-100 of low-density lipoprotein ( 3 ). Although it has been known for many years that elevated plasma Lp(a) concentrations are associated with thrombo-atherogenic diseases ( 4 – 6 ), recent evidence from large cohorts has finally confirmed a causal relationship ( 7 – 11 ). Therefore, in a consensus report, the European Atherosclerosis Society recommended screening for Lp(a) in people at moderate to high risk of cardiovascular disease, in which the desirable cut-off for Lp(a) was set at less than 50 mg/dl ( 12 ).

The thrombo-atherogenic properties of Lp(a) have also been well documented in transgenic mice ( 13 , 14 ). Several hemostatic pathways have been attributed to the pathomechanisms of Lp(a) ( 15 , 16 ). Due to its high atherogenicity, several attempts were made to treat individuals with increased Lp(a) levels with either medication or diet ( 16 ), without success. Even though nicotinic acid and its derivatives lower Lp(a) levels by up to 30%, they are not widely used due to frequent side effects. Therefore, to date, there is no safe drug available for the treatment of individuals with elevated plasma Lp(a) levels, and the development of new drugs is hampered by a lack of detailed knowledge of both Lp(a) biosynthesis and catabolism.

Previous turnover studies in humans demonstrated that plasma Lp(a) levels strongly correlate with its rate of biosynthesis but not with the fractional catabolic rate ( 17 , 18 ). Thus, any attempt to control plasma Lp(a) levels should focus on an interference with APOA biosynthesis. This has been supported by in vivo studies using antisense strategies in which plasma levels of an N-terminal APOA fragment expressed in mice under the control of the CMV promoter were reduced to almost zero ( 19 ). However, small molecule medications are not yet available.

The farnesoid X receptor (FXR, also known as NR1H4) is a bile acid–activated receptor and belongs to the nuclear receptor superfamily of ligand-activated transcription factors ( 20 – 23 ). FXR is mainly expressed in the liver, intestine, kidney, and adrenal glands. FXR heterodimerizes with the retinoid X receptor (RXRα also known as NR2B1), binds to FXR response elements (FXREs) that are usually but not exclusively inverted repeat-1 (IR-1), and regulates transcription of target genes ( 24 ). A direct repeat (DR) with a similar core sequence is also compatible for binding of FXR, either as a monomer or heterodimer ( 24 – 27 ). FXR plays important roles in bile acid, cholesterol, lipoprotein, and triglyceride metabolism. Activation of hepatic FXR modulates the expression of many hepatic genes involved in lipid metabolism. Studies using Fxr –/– mice have illustrated the importance of this nuclear receptor in maintaining cholesterol and bile acid homeostasis ( 28 , 29 ).

In the present study, we report that transcription of the APOA gene is under strong control of FXR, which binds to a negative control element located at the –826-bp region of the human APOA promoter. FXR was found to interfere with the hepatocyte nuclear factor 4α–mediated (HNF4α-mediated) (HNF4α is also known as NR2A1) activation of APOA transcription.

Elevated bile acid levels drastically reduce plasma Lp(a) levels in humans. We consistently noticed in various clinical settings that patients suffering from obstructive jaundice exhibited very low or even undetectable levels of plasma Lp(a). To study this in a more systematic way, patients with obstructive jaundice were analyzed for markers of cholestasis, such as bilirubin, lipoprotein X (LP-X), and plasma bile acid concentrations, and the results were correlated with Lp(a) levels. Supplemental Table 1 (supplemental material available online with this article doi: 10.1172/JCI45277DS1) lists the results from 20 patients suffering from biliary obstruction due to pancreatic, gallbladder, or bile duct cancer. In addition, 1 patient with congenital biliary atresia and 5 patients with choledocholithiasis were included. All patients had elevated plasma bilirubin concentrations (316 ± 48 μmol/l) and were positive for plasma LP-X (370 ± 47.7 mg/dl). Notably, the patients had plasma total bile acid levels of 98.9 ± 9.2 μmol/l that were more than 10-fold higher when compared with those of healthy individuals (Figure 1A). In 13 out of 20 of these patients, the plasma Lp(a) concentrations before therapy were less than 1 mg/dl, which is the detection limit of the particular assay. The remaining 7 patients had very low Lp(a) levels in relation to their APOA isoform (K-IV repeats). After successful surgical or endoscopic treatment of biliary obstruction, bilirubin, LP-X, and total bile acid levels were normalized, and Lp(a) concentrations rose significantly to levels that correspond to those of healthy controls with the corresponding APOA isoforms. Mean plasma Lp(a) levels were 2.7 ± 1.1 mg/dl before therapy and 20.3 ± 4.4 mg/dl after therapy (Figure 1B).

Low plasma Lp(a) levels in patients with obstructive jaundice. Plasma samples from 20 patients suffering from obstructive jaundice of different etiology were assayed for (A) total bile acids and (B) Lp(a), before and after surgical or endoscopic treatment. Values are expressed as mean ± SEM. See Supplemental Table 1 for details.

A cholestatic mouse model with elevated bile acid levels exhibits very low plasma and hepatic expression of APOA. To determine the effects of obstructive cholestasis on plasma levels and hepatic APOA expression, mice transgenic for human APOA (tg-APOA mice) and tg-APOA mice that were Fxr deficient (tg-APOA/Fxr –/– mice) were subjected to biliary obstruction by common bile duct ligation (CBDL) for 3 days. CBDL in tg-APOA mice resulted in significantly elevated serum liver enzymes (Supplemental Table 2), total bile acids, and bilirubin (Figure 2, A and B). The accumulation of endogenous bile acids in tg-APOA mice led to dramatic reduction of plasma APOA levels by 87% (Figure 2C) and of hepatic APOA mRNA expression by 98% (Figure 2D). CBDL in tg-APOA/Fxr –/– mice showed a small but measurable reduction of plasma APOA by 15% and hepatic mRNA by 19% (Supplemental Figure 1), which might be due to inflammation and hepatic injury. In conclusion, low APOA levels found in mouse and human cholestasis suggested that APOA expression is regulated by bile acids in vivo.

Drastic reduction in plasma levels and hepatic mRNA expression of APOA in a mouse model of cholestasis. tg-APOA mice were subjected to biliary obstruction by CBDL (n = 3 per group) or sham operation (n = 4 per group) for 3 days. (A and B) Total bile acids and bilirubin were measured in serum. Data are presented as mean ± SD. ***P ≤ 0.001, when compared with sham-operated mice. (C) Plasma levels of APOA were measured by DELFIA and are expressed as mean ± SD (***P ≤ 0.001). (D) Liver APOA mRNA levels were analyzed by real-time quantitative PCR normalized to cyclophilin and are expressed relative to those of sham-operated mice. Results represent mean ± SEM (***P ≤ 0.001).

Cholic acid feeding reduces plasma APOA concentrations and hepatic APOA expression in transgenic APOA mice. To study regulation of the human APOA gene expression by bile acids in a non-cholestatic model, tg-APOA and tg-APOA/Fxr –/– mice expressing the human APOA gene were fed for 5 days with either a normal chow diet (control) or a chow diet supplemented with 0.2% cholic acid (CA) (w/w). No changes in body weight or food intake were observed between control and treated groups (data not shown). Plasma total cholesterol and triglyceride levels were reduced in tg-APOA mice upon CA feeding but remained unchanged in tg-APOA/Fxr –/– mice (Supplemental Figure 2). A 0.2%-CA supplementation led to a significant 72% decrease in plasma APOA levels in tg-APOA mice (Figure 3A). To evaluate whether the reduction of plasma APOA levels was due to decreased APOA mRNA levels in liver, real-time quantitative PCR analysis was performed. APOA mRNA levels were significantly decreased in the livers of the CA-fed tg-APOA mice (Figure 3B). Western blot analysis of liver homogenates confirmed that this repression also occurs at the protein level upon CA feeding in tg-APOA mice (Figure 3C). In tg-APOA/Fxr –/– mice, however, plasma APOA concentrations (Figure 3D), hepatic APOA mRNA levels (Figure 3E), and protein levels (Figure 3F) were comparable in control and CA-treated mice. Taken together, these data indicate that both plasma levels and hepatic expression of human APOA are downregulated by CA feeding in tg-APOA mice in an FXR-dependent manner.

CA decreases plasma levels and hepatic expression of APOA in tg-APOA mice but not in tg-APOA/Fxr –/– mice. tg-APOA mice (n = 8 per group) and tg-APOA/Fxr –/– mice (n = 8 per group) expressing human APOA were fed 0.2% CA (w/w) mixed in normal chow for 5 days. Control mice received normal rodent chow. (A and D) Plasma levels of APOA were measured by DELFIA and are expressed as mean ± SD (**P ≤ 0.01). (B and E) Mouse liver APOA mRNA levels were analyzed by real-time quantitative PCR and normalized to cyclophilin and are expressed relative to those of control mice. Results represent mean ± SEM (***P ≤ 0.001). (C and F) Western blot analysis and densitometric quantification of APOA levels in the protein extracts from liver tissue (expressed as mean ± SD relative to controls **P ≤ 0.01). The gene expression profile was analyzed in (G) tg-APOA mice and (H) tg-APOA/Fxr –/– mice by real-time quantitative PCR. mRNA expression in control mice was arbitrarily set to 1 and normalized to that of cyclophilin. Results represent mean ± SEM (***P ≤ 0.001, **P ≤ 0.01, *P < 0.05).

Subsequently, we profiled hepatic expression of known FXR target genes involved in bile acid and cholesterol metabolism after a 5-day feeding of tg-APOA and tg-APOA/Fxr –/– mice with CA (Figure 3, G and H). As expected, CA treatment of tg-APOA mice led to a strong inhibition of both Cyp7a1 and Cyp8b1 ( 30 – 32 ), 2.3-fold upregulation of small heterodimer partner (Shp, also known as NR0B2) ( 33 ), and induction of Bsep. No changes were observed in the hepatic mRNA expression of Lrh1 and Hnf4a. Fgf15 mRNA in the ileum was upregulated by 2.8 fold ( 34 ). CA feeding did not change hepatic expression of Cyp3a11, a target gene of pregnane X receptor (PXR also known as NR1I2) in tg-APOA mice, indicating that PXR was not activated by 0.2% CA in the diet. In contrast, CA treatment had no impact on mRNA levels of FXR target genes in tg-APOA/Fxr –/– mice, yet Cyp3a11 expression was induced ( 35 , 36 ).

Bile acids can induce inflammation in the liver and cause liver damage ( 37 ). Moreover, cholestasis in humans and mice is characterized by high inflammation ( 38 ). Therefore, we studied the hepatic expression of several proinflammatory genes. A 0.2%-CA feeding did not change the expression levels of proinflammatory cytokines such as Il6, Il1b, and Tnfa in tg-APOA mice (Supplemental Figure 3A), whereas Il6 expression was 2.6-fold increased in tg-APOA/Fxr –/– mice (Supplemental Figure 3B). Taken together, these results showed that bile acids repress APOA expression by an FXR-mediated mechanism.

CA and GW4064 decrease human APOA gene expression in primary hepatocytes. To further study the direct mechanism of the inhibitory effect of FXR on hepatic APOA expression, we then studied the influence of FXR agonists on APOA expression in mouse primary hepatocytes. For this purpose, primary hepatocytes were isolated from tg-APOA mice and incubated with different concentrations of the natural FXR ligand CA. Analysis of mRNA levels by real-time quantitative PCR revealed a significant dose- and time-dependent decrease in APOA transcript levels, suggesting a transcriptional effect (Figure 4, A and B). Western blot analysis confirmed that this CA-mediated repression also occurs at the protein level (Figure 4C). Cell viability was assessed with the trypan blue exclusion test, revealing that all concentrations of CA were well tolerated by the cells (data not shown).

FXR agonists downregulate APOA gene expression in a dose- and time-dependent manner in primary mouse hepatocytes. (A) Primary mouse hepatocytes from tg-APOA mice were incubated with increasing concentrations of CA (50, 100, and 200 μM) or vehicle (control) for 24 hours. APOA mRNA levels were analyzed by real-time quantitative PCR. These data are presented as mean ± SEM (**P ≤ 0.01, *P < 0.05). (B) Primary mouse hepatocytes were incubated with CA (200 μM) or vehicle for 12, 24, and 48 hours. APOA mRNA levels were measured by real-time quantitative PCR. Results represent mean ± SEM of 3 independent experiments (**P ≤ 0.01). (C) Western blotting and densitometric analyses of APOA expression in whole cell lysates from hepatocytes treated for 24 hours with increasing concentrations of CA (expressed as mean ± SD relative to controls *P < 0.05). (D) Primary hepatocytes were treated with GW4064 (5 μM) for 24 hours and analyzed for APOA mRNA levels by real-time quantitative PCR. These data are presented as mean ± SEM of 3 independent experiments (**P ≤ 0.01). (E) Western blotting and densitometric analyses of APOA expression in whole cell lysates from hepatocytes treated for 24 hours with GW4064 (5 μM) (expressed as mean ± SD relative to controls **P ≤ 0.01).

Since bile acids may exert FXR-independent effects by activating other signal transduction pathways ( 39 , 40 ), we additionally tested the influence of the synthetic nonsteroidal FXR agonist GW4064 on APOA gene expression. Treatment of primary hepatocytes with 5 μM GW4064 for 24 hours resulted in a significant decrease of APOA mRNA (Figure 4D) and protein levels (Figure 4E) when compared with those of vehicle-treated control cells. In addition, we measured the expression levels of control FXR target genes after treatment with CA and GW4064 and found that both ligands increased Shp and markedly decreased Cyp7a1 and Apoa1 mRNA levels (Supplemental Figure 4, A and B).

Overall, these results demonstrate that both natural and synthetic FXR agonists downregulate human APOA expression in cultured mouse primary hepatocytes via a transcriptional mechanism.

Mapping of an FXRE in the human APOA promoter. To provide direct evidence for the FXR-mediated inhibitory effect on the APOA promoter and to further identify relevant promoter element(s), a 2-kb fragment of human APOA promoter (from –1,952 bp to +52 bp, referred to herein as hAPOA –1,952/+52 promoter) was cloned into pGL3-luciferase reporter plasmid (Figure 5A). In addition a series of 5′ deletion constructs were generated, as shown in Figure 5D. Transient transfections were performed in HepG2 cells with the hAPOA –1,952/+52 promoter construct in the absence or presence of FXR and FXR agonists. FXR alone resulted in a 29% decrease in promoter activity, and this effect was further enhanced by the addition of chenodeoxycholic acid (CDCA) (63%) (Figure 5B). Likewise, incubation with FXR and GW4064 also strongly repressed the activity of the hAPOA –1952/+52 promoter by 57% (Figure 5C). In the absence of FXR overexpression, the hAPOA –1952/+52 promoter was inhibited by 25% or less by CDCA or GW4064 alone. This decrease likely resulted from the activation of the endogenous FXR that is expressed in HepG2 cells ( 27 ).

Bile acids and the nonsteroidal FXR agonist GW4064 downregulate human APOA promoter activity via FXR. (A) Scheme of the full-length hAPOA –1,952/+52 promoter–driven luciferase reporter system. (B and C) HepG2 cells were transfected with the hAPOA –1,952/+52 promoter reporter plasmid (150 ng) in the presence of either the pcDNA3 (control) or FXR expression vector (150 ng). Cells were subsequently treated with CDCA (100 μM), GW4064 (500 nM), or vehicle for 36 hours. Values are normalized to internal control β-galactosidase and expressed as percentages. Transfections were performed in triplicates, and each experiment was repeated at least 3 times. (D) Scheme of the deletion constructs of the human APOA promoter used in the luciferase reporter assay. HepG2 cells were transfected with the indicated human APOA promoter reporter plasmids (150 ng) in the presence of pcDNA3 empty or FXR expression vector (150 ng). Cells were then treated with CDCA (100 μM) or vehicle for 36 hours. Values are normalized to internal control β-galactosidase activity. (E) Scheme showing wild-type and mutant sequences. Mutations are indicated in bold lowercase letters. Underlined letters define the DR-1 element. (F) Mutational analysis of the human APOA promoter. HepG2 cells were transfected with the wild-type and mutant (M1, M2) human APOA promoter reporter plasmids in the presence of pcDNA3 empty or FXR containing expression vector (150 ng). Cells were then treated with CDCA (100 μM) or vehicle for 36 hours. Values are normalized to β-galactosidase activity and expressed as percentages. Data are presented as mean ± SD (**P ≤ 0.01, *P < 0.05).

To avoid endogenous FXR-mediated feedback inhibition, transient transfection experiments were performed in COS-7 cells, a nonhepatic cell line. Transfection of COS-7 cells in the absence or presence of FXR repressed the hAPOA –1,952/+52 promoter activity by 24%, an effect that was significantly enhanced by CDCA (43%) (Supplemental Figure 5). These experiments demonstrated that ectopic expression of FXR and a physiological concentration of CDCA are required to repress the APOA promoter activity in nonhepatic cells.

Since Shp was induced by CA treatment in vivo and in vitro, we subsequently studied the APOA promoter activity upon cotransfection of cells with increasing concentrations of a SHP expression plasmid. Surprisingly, SHP did not lower APOA promoter activity but further enhanced it (Supplemental Figure 6). Taken together, these results showed that FXR can regulate APOA promoter activity in a direct and SHP-independent manner.

Next, to identify promoter elements responsible for the observed effects of FXR, HepG2 cells were transfected with 5′ deletion constructs of the human APOA promoter in the absence or presence of FXR and/or CDCA. Reduced promoter activities were noted for both the –1,446-bp and –857-bp constructs (Figure 5D). However, the repression was relieved for –757-, –657-, –477-, and –148-bp promoter constructs, indicating that the region between –857 bp to –757 bp of the human APOA promoter contains a potential negative FXRE, which might be responsible for the observed bile acid response.

Notably, in silico Matinspector promoter analysis ( 41 ) and NUBIScan algorithm ( 42 ) suggested the presence of a DR-1 element located between nucleotides –826 and –814. Previous studies have already shown that the DR-1 element can function as an FXRE ( 24 , 43 , 44 ).

To test whether this DR-1 site could mediate FXR-dependent repression of the APOA promoter, we introduced mutations in the context of the full-length hAPOA –1,952/+52 promoter (WT) and generated 2 mutant constructs (M1 and M2), as shown in Figure 5E. Mutation (M2) of this site completely abolished the FXR-mediated repression of APOA promoter activity (Figure 5F), indicating the binding of FXR to the second half site of the DR-1 element.

Taken together, these results suggest that the DR-1 site, located between nucleotides –826 and –814, is a negative response element via which FXR represses human APOA promoter activity.

FXR binds to the DR-1 site of the APOA promoter in EMSA. To provide additional evidence that the DR-1 element at the –826-bp region of the human APOA promoter can function as an FXRE, gel shift assays were performed. Consensus IR-1 probe was used as a positive control. FXR bound the labeled IR-1 probe both in the absence (Figure 6A, lane 3) and presence (Figure 6A, lane 4) of RXR. In contrast, FXR bound as a monomer to the radiolabeled probe containing a wild-type DR-1 element (DR-1 WT) (Figure 6A, lanes 7 and 8) but not to the probe carrying the mutated DR-1 element (DR-1 M2) (Figure 6A, lanes 11 and 12). Formation of the FXR-DNA complex was specifically competed by cold DR-1 WT probe (Figure 6B, lanes 3–5), whereas the DR-1 M2 probe did not compete (Figure 6B, lane 6). Binding of FXR to the DR-1 WT probe was also competed by a cold IR-1 probe (Figure 6B, lane 7) notably, the cold IR-1 and cold DR-1 WT probes competed with a similar efficiency for the labeled DR-1 WT oligo. These results indicated that FXR binds specifically to the DR-1 site of human APOA promoter.

FXR binds to the DR-1 element of the human APOA promoter as a monomer. (A) EMSAs were performed with radiolabeled IR-1 consensus FXRE (lanes 1–4), DR-1 WT (lanes 5–8), and DR-1 M2 (lanes 9–12) probes using in vitro transcribed/translated RXR (lanes 2, 6, and 10), FXR (lanes 3, 7, and 11), both RXR and FXR (lanes 4, 8, and 12), or unprogrammed reticulocyte lysate (lanes 1, 5, and 9) as indicated. (B) Competition EMSAs on radiolabeled DR-1 WT probe were performed by adding 50-fold, 100-fold, 200-fold molar excess of the indicated cold DR-1 WT (lanes 3–5) and 50-fold molar excess of cold DR-1 M2 (lane 6) and IR-1 (lane 7) probes. Numbering indicates relative intensity of the bands.

FXR competes for HNF4α binding to the DR-1 element. DR-1 elements have been shown to function as HNF4α response elements ( 45 ). In order to investigate the regulation of APOA gene expression by HNF4α, we overexpressed HNF4α in cultured primary hepatocytes and studied the expression of the human APOA gene. As shown in Figure 7A, adenovirus-mediated overexpression of HNF4α in mouse primary hepatocytes from tg-APOA mice dose dependently induced the expression of APOA mRNA levels compared with LacZ-transfected cells.

Effects of hepatocyte nuclear factor HNF4α overexpression on human APOA. (A) Primary mouse hepatocytes were infected with either adenovirus coding for β-galactosidase (Ad-LacZ) or human HNF4A (Ad-HNF4α). Total RNA was extracted, and gene expression was measured by real-time quantitative PCR. Data represent mean ± SEM. (**P ≤ 0.01). (B) HepG2 cells were transfected with the hAPOA –1,952/+52 reporter plasmid (150 ng) in the presence of increasing amounts of HNF4α expression vector. Values represent mean ± SD (**P ≤ 0.01). (C) HepG2 cells were transfected with the hAPOA –1,952/+52 reporter plasmid in the presence or absence of FXR and HNF4α. Cells were then treated with CDCA (100 μM) or vehicle for 36 hours. Values are normalized to internal control β-galactosidase and expressed as percentages. Values represent mean ± SD (**P ≤ 0.01, *P < 0.05). (D) HNF4α binds to the DR-1 motif in the human APOA promoter. EMSAs with end-labeled DR-1 WT probe using in vitro transcribed/translated HNF4α (lanes 2). Competition analysis was performed by adding 50-fold (lane 3) and 100-fold (lane 4) molar excess of the indicated cold DR-1 WT probe. Underlined letters indicate the DR-1 element. (E) tg-APOA mice were fed normal chow or 0.2% CA chow for 24 hours, and livers were collected for ChIP analyses. For ChIP assay, sheared chromatin was immunoprecipitated with the indicated antibodies. The final DNA extractions were amplified by PCR using primer pairs covering the distal region and the DR-1 motif of the APOA gene promoter. As a positive control for FXR/RXR binding, the Shp gene promoter was amplified by PCR.

Next, we studied the effect of HNF4α overexpression on the activity of the hAPOA –1,952/+52 promoter. As shown in Figure 7B, overexpression of HNF4α in HepG2 cells dose dependently transactivated the human APOA promoter. However, additional cotransfection with FXR and/or CDCA treatment abolished the HNF4α-mediated transactivation (Figure 7C). This effect might be due to the occupancy of the HNF4α response element (DR-1) by FXR. HNF4α-mediated transactivation of the hAPOA –1,952/+52 promoter was also observed in the nonhepatic cell line, COS-7, which neither expressed FXR nor HNF4α. Cotransfection with FXR alone or with FXR and CDCA significantly inhibited HNF4α transactivation (Supplemental Figure 5), suggesting that FXR competes with HNF4α for the DR-1 binding motif. We then performed a mobility shift assay to check whether HNF4α binds to the DR-1 element at the –826-bp region of the human APOA promoter. HNF4α bound to the radiolabeled probe containing DR-1 WT, and the protein-DNA complex was specifically competed by cold unlabeled WT probe (Figure 7D).

Taken together, these results suggest that this response element at –826 bp might be occupied by HNF4α at the basal level, whereas bile acid activation leads to a switch of occupancy of this site by FXR.

To further confirm the interaction of FXR with the DR-1 element in the APOA promoter, in vivo ChIP experiments were performed with liver tissue isolated from tg-APOA mice fed for 24 hours with normal chow or with chow containing 0.2% CA (Figure 7E). In the control group, antibodies against HNF4α precipitated DNA encompassing the DR-1 element (–826- to –814-bp region) in the APOA promoter. By contrast, 0.2%-CA feeding led to occupancy of this response element by FXR alone without RXR. As a negative control, an equivalent amount of chromatin precipitated with a nonrelevant anti-IgG antibody did not result in any signal. The same DNA samples were PCR amplified by using primers covering the distal region of the APOA promoter, but no signal was observed, whereas 0.2%-CA feeding increased the occupancy of both FXR and RXR to the Shp promoter.

Taken together, these results prove that the DR-1 element at the –826- to –814-bp region of the human APOA promoter could mediate the FXR repression of APOA transcription by a competition between FXR and HNF4α (Supplemental Figure 7).

Meta-analyses from prospective and epidemiological studies demonstrated an association of elevated plasma Lp(a) levels with an increased risk for ischemic heart diseases and stroke ( 7 – 11 ). Lp(a) is causally associated with an increased risk for myocardial infarction and reported to increase the likelihood for major adverse cardiovascular events when plasma Lp(a) levels exceed 30 mg/dl by 2.3 fold ( 9 , 46 ). Therefore, unraveling molecular and pharmacological factors reducing Lp(a) constitutes a novel goal, with important therapeutic and pharmacological consequences in human population. In this study, we identified the bile acid–activated receptor FXR as a major repressor of Lp(a) levels in patients and mice having high bile acid levels. Notably, therapeutic normalization of bile acid concentrations lead to increased plasma Lp(a) levels. Similarly, bile duct ligation in transgenic APOA mice virtually abrogated APOA expression. Therefore, we hypothesized that high intrahepatic bile acid could suppress APOA expression. In order to firmly demonstrate that bile acid–activated FXR was repressing APOA expression in a more physiological condition, we fed transgenic APOA and Fxr –/– transgenic APOA mice with bile acids. Bile acid feeding lowered APOA plasma concentration as well as gene expression and protein levels in transgenic APOA mice, an effect abolished in Fxr –/– transgenic APOA mice. Furthermore, in vitro activation of FXR by bile acids or a nonsteroidal FXR agonist lowered APOA gene expression in a time and dose-dependent manner, due to a transcriptional mechanism.

In normal individuals, plasma Lp(a) levels have been shown to correlate significantly with the synthesis rate of APOA ( 17 , 18 ) and appear to be minimally affected by its catabolism. Thus, pharmacological FXR activation could constitute a novel and promising approach to treat hyper-Lp(a) individuals and significantly reduce adverse coronary events in a high-risk population. Interestingly, novel FXR agonists were shown already to display antiatherosclerotic effects in mice ( 47 ) and to normalize dyslipidemia ( 48 ) in rodent models lacking APOA expression. It will therefore be of interest to measure Lp(a) in ongoing human clinical trials using FXR agonists such as INT-747. Bile acid binding resins, probably by depleting FXR ligand and therefore lowering FXR activation, were shown to reduce the incidence of coronary disease ( 49 ), due to the lowering of cholesterol and glucosuria ( 50 ). Novel resins with higher bile acid affinity, specificity, and binding capacity are currently available and constitute a safe therapeutic option, either in combination with HMGCoA reductase inhibitors (statins) or in statin-resistant patients. However, our results allude to the fact that resins and intestinal bile acid uptake inhibitors should be carefully monitored in a patient population with dyslipidemia and potentially elevated Lp(a). Conversely, introducing Lp(a) as a screening parameter could lead to improved resins without side effects, with even more cardiovascular protective.

FXR was found to directly repress APOA promoter activity by binding to a DR-1 site shared with HNF4α, leading to suppression of transcription. Several molecular regulators were found to bind to and modulate the promoter region of APOA. These include binding sites for HNF1, HNF4α, RXR, and LINE among others ( 51 , 52 ). FXR is highly expressed in the liver and was found to bind to IR-1 response elements in promoters as a heterodimer with RXR as well as to various DR elements ( 24 ), thereby transactivating cognate target genes. In addition, FXR can bind monomeric response elements and hence directly repress gene transcription ( 24 – 27 ). Recently, the location and sequence of FXRE was systematically studied via ChIP and sequencing ( 53 ). In this work, Chong et al. identified 1,656 binding sites, including 10% located in the proximal 2 kb of the promoter. Moreover, up to 25% of these FXREs were not classical IR-1. In our study, by combining, reporter assays, site-directed mutagenesis, EMSA, and ChIP, we unambiguously identified a DR-1 located at –826-bp upstream of the transcription start site as what we believe to be a new negative FXRE in the promoter of APOA. This site is therefore compatible with the architecture of a bona fide FXRE. Since Chong et al. used a mouse liver not expressing APOA chromatin-enriched material, our response element could not be found in their database. However, the DR-1 located at the –826- to –814-bp region was found to be bound and activated by HNF4α as shown by transfection, EMSA, and ChIP. In addition, HNF4α was competitively displaced by FXR, as demonstrated in Figure 7E. HNF4α is well known to be involved in lipid, glucose, and bile acid homeostasis ( 54 ). A competition between FXR and HNF4α was previously found in the promoter of APOCIII ( 24 , 43 , 44 ). It is therefore tempting to speculate that the balance between FXR and HNF4α binding on gene promoters could coordinate a network of genes involved in lipid homeostasis (Supplemental Figure 7). The precise mechanism for this suppression, including the events involved in a nonproductive FXR binding to a response element, requires additional studies.

FXR also transactivates mouse Fgf15, a gene that is expressed almost exclusively in the terminal ileum, and its human ortholog, FGF19, a gene that is expressed in the small intestine as well as in the liver. FGF15/19 signals from intestine to the liver to repress the transcription of key enzymes of bile acid biosynthesis ( 34 , 55 ). Notably, FXR activation efficiently repressed APOA in vitro in primary mouse hepatocytes that do not express Fgf15, indicating that FXR can regulate the APOA gene in an FGF15/19-independent manner. Further studies will be required to clarify a possible additional role of FGF15/19 in APOA gene repression.

In addition, FXR can indirectly modulate gene expression via the induction of Shp in the liver ( 31 ). Although SHP is a transcriptional repressor, it has no DNA binding motif ( 56 ) but interacts with several nuclear receptors, such as LRH-1 or HNF4α, thereby interfering with gene transcription. Recently, the SHP/LRH-1/CYP7A1 signaling pathway was disproved, and LRH-1 was identified as a master regulator of Cyp8b1 ( 57 , 58 ). Since SHP is able to interact in vitro with multiple partners, the identification of the actual SHP targets is still an open quest. Our transgenic APOA mice fed with CA or primary hepatocytes incubated with FXR activators were found to have more Shp and less APOA gene expression. We therefore wondered whether Shp induction could repress APOA. However, dose response transfection experiments with SHP expression plasmid showed that SHP did not repress and instead increased the APOA promoter activity in HepG2 as well as in COS-7 cells (Supplemental Figure 6). Conversely, FXR directly repressed APOA promoter activity by binding to a DR-1 also recognized by HNF4α. This was verified by ChIP assay, which impressively confirmed that the DR-1 element at the –826- to –814-bp region of the APOA promoter is occupied by HNF4α, whereas CA activation leads to a switch of occupancy of the site by FXR (Figure 7E). Taken together, our data suggest that SHP does not regulate the APOA promoter in contrast to FXR.

In view of the present results, FXR agonist could constitute a new therapeutic avenue to treat hyper-Lp(a) states and may be useful in the treatment of atherosclerotic disease and myocardial infarction. In addition, these results suggest that present and future FXR partial agonists, also called bile acid receptor modulator (BARM), have to be monitored for possible adverse effects on plasma Lp(a) levels in human clinical trials.

Chemicals. CA and CDCA were purchased from Sigma-Aldrich. GW4064 was purchased from Tocris Bioscience. Collagenase was purchased from Worthington Biochemical Corporation.

Patients. Patients suffering from obstructive jaundice due to gallstones or malignancy were studied for markers of biliary obstruction and plasma Lp(a) concentrations. Blood of patients referred to surgery or endoscopy was analyzed immediately for plasma levels of Lp(a), bilirubin, total bile acids, and LP-X. After appropriate treatment, reversal of jaundice, and normalization of plasma bilirubin, plasma Lp(a) levels were measured again. All human studies were approved by the ethical committee of the Medical University of Graz and were performed in accordance with the Helsinki Declaration. Informed consent was received from all patients or their parents for drawing extra blood to perform lipid and lipoprotein analyses.

Analysis of plasma parameters and Lp(a) in patients. Lipids from human plasma were measured enzymatically using the assay kits from Roche Diagnostics. Lp(a) was quantified by an in-house DELFIA method. The preparation of Lp(a) and APOA and the standardization of the Lp(a) assay have been described in detail previously ( 59 ). The determination of APOA isoforms was performed by Western blotting as described previously ( 60 ). LP-X was measured by standard methods ( 61 ). Total plasma bile acids were measured enzymatically ( 62 ).

Animal experiments. All animal experiments were performed after approval of the protocol by the Austrian Federal Ministry of Science and Research, Division of Genetic Engineering and Animal Experiments (Vienna, Austria). Fxr –/– mice ( 28 ) were backcrossed for 5 generations with tg-APOA mice, carrying a 110-kb human APOA gene surrounded by more than 60-kb 5′- and 3′-flanking DNA in the YAC ( 63 ). Mice were hosted under a standard 12-hour-light/12-hour-dark cycle and fed standard rodent chow diet and water ad libitum. Female mice, between 10 and 12 weeks old, were used in all the experiments. For feeding studies, tg-APOA (n = 8) and tg-APOA/Fxr –/– mice (n = 8) expressing the human APOA were divided into 2 groups. Animals were randomized based on plasma APOA levels. One group received a normal rodent chow diet (control), whereas the other group received the same diet supplemented with 0.2% (w/w) CA for 5 days. At sacrifice, mice were fasted for 4 hours before blood samples were collected. Liver and ileum samples were harvested and stored at –80°C until further analysis. For the ChIP assay, female tg-APOA mice (n = 3) were fed with either normal chow (control group) or chow with 0.2% CA for 24 hours. Freshly isolated liver tissue was pooled and used to isolate chromatin for immunoprecipitation.

Plasma lipid parameters in mice. Blood was collected by retro-orbital bleeding and EDTA plasma was prepared within 20 minutes. Plasma concentrations of APOA were measured enzymatically by an in-house DELFIA method. Plasma triglyceride (DiaSys) and total cholesterol concentrations (Greiner Diagnostics AG) were determined enzymatically according to the manufacturer’s protocols.

CBDL. Twelve-week-old female tg-APOA mice (n = 3–4 per group) and tg-APOA/Fxr –/– mice (n = 3 per group) were subjected to CBDL as described previously ( 64 ). In brief, the common bile duct was ligated close to the liver hilus, immediately below the bifurcation, and dissected between the ligatures. Sham-operated animals were subjected to the same surgical procedure but without ligation of the common bile duct. Sera and livers were collected for analysis 3 days after surgery. Liver tissue was frozen in liquid nitrogen for further RNA preparations. Serum was stored at –80°C until analysis. Serum alanine aminotransferase, aspartate aminotransferase, alkaline phosphatase levels, and bilirubin were determined by routine testing on a Hitachi 917 analyzer (Boehringer Mannheim), as measures of the degree of cholestasis. Total serum bile acid levels were determined enzymatically using the Bile Acid Kit (Ecoline S+, DiaSys Diagnostic Systems).

Cell cultures. Mouse primary hepatocytes from tg-APOA mice were prepared and cultured as described previously ( 65 ), with minor modifications. The mouse liver was perfused with collagenase solution, and liver cells were collected. After filtration and centrifugation, the isolated hepatocytes were resuspended in DMEM (Invitrogen) supplemented with 20% (v/v) FCS (Sigma-Aldrich), 100 units/ml penicillin, and 100 units/ml streptomycin and placed in 6-well collagen-coated plates (BD Biosciences) at a density of 1 × 10 5 cells/well at 37°C in an atmosphere of 5% CO2 for 4 hours. Thereafter, cells were cultured in DMEM supplemented with 10% FCS and 100 units/ml penicillin/streptomycin for 16 hours. Experiments were performed in serum-free DMEM supplemented with various concentrations of the FXR ligands CA and GW4064.

The HepG2 and COS7 cells were obtained from ATCC. The cells were maintained in DMEM containing 10% FCS and 100 units/ml penicillin/streptomycin.

RNA extraction, reverse transcription, and real-time PCR. Total RNA from cells and mouse tissues was isolated using TRI zol (Invitrogen) according to the manufacturer’s protocol. Two micrograms of total RNA were reverse transcribed using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Quantitative real-time PCR was performed on a Light Cycler 480 instrument (Roche Diagnostics), using the QuantiFast SYBR Green PCR Kit (Qiagen). Primer sequences are listed in Supplemental Table 3. The gene expression values were normalized to cyclophilin A (Ppia) as a housekeeping gene. The data were analyzed by the public domain program Relative Expression Software Tool (REST http://www.gene-quantification.de/download.html#rest) ( 66 ). Values are presented as mean ± SEM.

Protein extraction and immunoblotting. Livers were homogenized or cells were lysed in an ice-cold RIPA buffer. The lysates were centrifuged (12,000 g) at 4°C for 10 minutes, and the supernatant was collected. Protein was quantitated using the Bradford protein assay (Bio-Rad). Equivalent amounts of protein homogenates were resolved by SDS-PAGE, transferred to a nitrocellulose membrane, and probed with rabbit polyclonal antibodies to human APOA (1:1,250) and a monoclonal anti-mouse β-actin (1:2,000) (Santa Cruz Biotechnology Inc.). The immunoblots were visualized by the Pierce ECL Chemiluminescence Detection System (Thermo Scientific). Densitometric analysis of the gels was carried out using ImageJ software ( http://rsbweb.nih.gov/ij/). Data are presented as mean ± SD.

Adenoviral infection of primary mouse hepatocytes. Primary hepatocytes from tg-APOA mice were isolated and maintained for 24 hours before infection with 50 and 100 MOI of adenovirus encoding LacZ or human HNF4A in serum-free DMEM. After a 4-hour infection, the cells were incubated in DMEM supplemented with 10% FCS for 24 hours, followed by cell harvesting for RNA analysis.

Plasmids. Expression plasmids encoding hFXR (pcDNA3-FXR), hRXRα (pSG5-RXR), SHP (pCDM8-SHP), and HNF4α (pSG5-HNF4α) were provided by Peter Young (Dupont, Oakley, California, USA), Philippe Lefebvre (Institut Pasteur de Lille, Lille, France), David D. Moore (Baylor College of Medicine, Houston, Texas, USA), and Mary C. Weiss (Institut Pasteur, Paris, France), respectively. The human APOA promoter construct (hAPOA –1,952/+52) was obtained by PCR amplification using human genomic DNA as a template. The PCR product was cloned into the pGL3 basic vector (Promega) as a MluI/BglII fragment to generate human APOA-Luc. (Primers used are as follows: for forward reaction, 5′-ACGCGTTCTGAGAGGGAGGTCAAAGTTTTC-3′, and reverse reaction, 5′-AGATCTCTTGAGAAAGCCAGCCCCAAAGGT-3′.) All constructs were verified by DNA sequencing (LGC Genomics).

Transient transfection and reporter gene assays. HepG2 cells were plated in 24-well plates over night before transfection. Cells at 60%–70% confluence were transiently transfected with the indicated reporter plasmid (150 ng), with or with out receptor expression plasmids (150 ng), using FuGENE 6 reagent (Roche Diagnostics) according to the manufacturer’s instructions. β-Galactosidase expression plasmid was cotransfected to assess the transfection efficiency. After 12 hours of transfection, medium was changed, and cells were exposed to the ligands (CDCA [100 μmol/l], GW4064 [500 nmol/l]) or vehicle. After 36 hours, cell extracts were prepared using passive lysis buffer (Promega) and assayed for luciferase and β-galactosidase activities using the Luciferase Assay System and the β-Galactosidase Enzyme Assay System, respectively (Promega). Luciferase activities were measured using Lumat LB9501 (Berthold) and normalized to β-galactosidase activities for each transfected well. For each experimental trial, wells were transfected in triplicate, and each well was assayed in duplicate. Data are presented as mean ± SD.

Site-directed mutagenesis. Mutagenesis was performed using the QuikChange Site-Directed Mutagenesis System (Stratagene), according to the manufacturer’s manual. The mutants were verified by sequencing. The oligonucleotides M1 (5′-GAGGGTTGGAAGCAAGAGGGGatCCAACGCGCACGGG­GAGGAAGC-3′) and M2 (5′-GAAGCAAGAGGGGGGCCAACatGCACGGGGAGGAAGCATTTGGGCAG-3′) were used to introduce mutations into the full-length hAPOA –1,952/+52. Mutated bases are indicated by bold, lowercase letters, and underlined letters define the DR-1 element.

EMSAs. Human FXR, RXR, and HNF4α proteins were synthesized in vitro using the TNT T7 Quick Coupled Transcription/Translation System (Promega). The sense and antisense oligonucleotide probes of DR-1 WT (5′-AGGGGGGCCAACGCGCACGGG-3′), DR-1 M2 (5′-AGGGGGGCCAACatGCACGGG-3′), and a FXR IR-1 consensus response element-containing oligonucleotide (IR-1, 5′-GATCTCAAGAGGTCATTGACCTTTTTG-3′) were annealed and radioactively labeled at the 5′ end using T4 polynucleotide kinase and γ- 32 P-ATP (Hartmann Analytic GmbH) (mutated bases are indicated by bold, lowercase letters, and underlined letters define the IR-1 element). Unincorporated nucleotides were removed by using Micro Bio-Spin 6 Columns (Bio-Rad). In vitro translated proteins (2.0 μl) were incubated for 20 minutes at room temperature in a total volume of 10 μl with binding buffer (Gel Shift Assay System, Promega) before the labeled probe was added. Binding reactions were further incubated for 30 minutes and resolved by 6% nondenaturing polyacrylamide gel electrophoresis in 0.25X Tris-Borate-EDTA buffer at room temperature and 120 V for 3.5 hours. The gel was dried and exposed to an X-ray film. For competition experiments, unlabeled probes were included in the binding reaction at the indicated excess concentrations.

ChIP assay. The in vivo ChIP assay was performed with freshly isolated mouse liver tissue using the EpiQuik Tissue ChIP Kit (Epigentek) according to the manufacturer’s instructions, with minor modifications. Liver tissue was fixed in formaldehyde for 12 minutes and then quenched for 5 minutes with glycine. The nuclei were extracted and sonicated to yield 500- to 1,000-bp DNA fragments. Aliquots of sheared chromatin were then immunoprecipitated using 4 μg anti-FXR (sc-13063 Santa Cruz Biotechnology Inc.), anti-RXR (sc-553 Santa Cruz Biotechnology Inc.), 2 μg anti-HNF4α antibody (sc-6556 Santa Cruz Biotechnology Inc.), or 1 μg anti-IgG antibody. Nonprecipitated chromatin (input) was used as a positive control. DNA extractions were PCR amplified using the following flanking primers, and the PCR products were analyzed by agarose gel electrophoresis: DR-1 element in the APOA promoter (DR-1 ChIP forward, 5′ TTGGCAGTGTTATTGGGAGAC 3′ DR-1 ChIP reverse, 5′ ACAGGCAGTTCCATCACTCC 3′), distal region of the APOA promoter (distal ChIP forward, 5′ TCTCCCCTTCATGTTTCCAG 3′ distal ChIP reverse, 5′ CCAGTGGCCGACATAGAGAT 3′), and Shp promoter (Shp ChIP forward, 5′GCCTGAGACCTTGGTGCCCTG 3′ Shp ChIP reverse, 5′ CTGCCCACTGCCTGGATGC 3′).

Statistics. Statistical analyses of the experiments were performed with GraphPad Prism 5.0. Two-tailed, unpaired Student’s t test was applied to determine statistical significance.

This work was supported by the Medical University of Graz (I. Chennamsetty and A. Baghdasaryan are funded by the PhD program “Molecular Medicine”), the Austrian Science Fund FWF (SFB-LIPOTOX F3004, F3008, and P19186), and the Austrian Federal Ministry of Science and Research (GEN-AU project Genomics of Lipid-associated Disorders — GOLD). The authors thank A. Ibovnik for excellent technical assistance.

Conflict of interest: The authors have declared that no conflict of interest exists.

Reference information: J Clin Invest. 2011121(9):3724–3734. doi:10.1172/JCI45277.


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Acknowledgements

We thank K Kourouniotis and G Fragiadakis for assistance in transgenesis and biochemical assays and C Pritchard, H Hilton and D Williams for technical support with microarrays and database submission and R Nilsson and L Swensson for bioanalytical chemistry of the sera. We also acknowledge the MRC Rosalind Franklin Centre (formerly UK HGMP Resource Centre) for provision of microarrays. This work was supported by GSRT and grants from EU (QLRT-2000-01513 and LSHG-CT-2004-502950).


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