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CELLULAR TRANSLOCATION MECHANISM OF RIBOFLAVIN
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Se-Ne Huang, B. S.
*****
The Ohio State University 2001
Dissertation Committee:
Dr. Peter W. Swaan, Advisor Approved by
Dr. William L. Hayton ^
Dr. Robert J. Lee Advisor
Dr. Martin E. Dowty Pharmacy Graduate Program UMI Number: 3031208
UMI*
UMI Microform 3031208 Copyright 2002 by Belt & Howell Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.
Bell & Howell Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, Ml 48106-1346 Copyright by Se-Ne Huang 2001 ABSTRACT
Riboflavin, also known as vitamin 82 , is essential for normal cellular growth and development. Humans cannot biosynthesize riboflavin, thus, they must obtain it from the diet or maternal sources. Despite its critical importance to cellular functions, the molecular mechanism of riboflavin translocation is still controversial and remains to be defined. The purpose of this thesis is to characterize and further elucidate the hypothesis that a receptor-mediated endocytosis (RME) component(s) is involved in cellular translocation of riboflavin. A two-tiered approach was employed to unravel the involvement of a RME pathway. First, using pharmacological agents known to alter the intracellular RME events, we demonstrated that transepithelial transport of riboflavin in
Caco-2 cell monolayers is mediated by microtubule-based movements and vesicular- sorting components (chapter 2). To obtain the morphological evidence of an endocytosis mechanism, we chose human trophoblast-derived BeWo cells as our model. Consistent with our finding in Caco-2 cells, a high affinity riboflavin transporter(s) was identified on the cellular membrane of Be Wo cells (chapter 3). Riboflavin structural analogs studies further indicated that isoalloxazine ring has a more essential role in ligand-transporter interactions than the D-ribose side chain. Based on this knowledge, a fluorescent-tag riboflavin conjugate was synthesized by specifically attaching rhodamine onto the ribityl moiety of riboflavin (chapter 4). The rational conjugation strategy resulted in a rhodamine-riboflavin conjugate with high specificity and affinity towards the riboflavin ii transpocter(&) comparable to unlabeled riboflaviiu Usings the conjugate as a probe together
with markers for cellular organelles and specific endocytic compartments, we further
identified the involvement of a clathrin-mediated endocytosis pathway in cellular
trafficking of rhodamine-riboflavin conjugate. Combined, our results demonstrated that
riboflavin is, in part, internalized via a receptor-mediated endocytosis process. The outcome of this research will aid in our understanding of the cellular absorption mechanism of riboflavin and will facilitate future evaluation of this vitamin transport system as a means to deliver imaging agents or therapeutic moieties of limited membrane permeability such as genes, enzymes and chemotherapeutics into cells.
Ill Dedicated to my parents and grandparents
IV ACKNOWLEDGMENTS
I would like to acknowledge my advisor, Dr. Peter W. Swaan for his guidance, and support. Without his instruction and help this thesis work would not be possible. I also want to express my gratitude to all faculty members in Division of Pharmaceutics, especially Drs. William L. Hayton, Robert J. Lee, and Martin E. Dowty for their invaluable advice and help.
I would like to acknowledge the enormous help and friendship given to me from my colleagues, Yongheng Zhang, Amy Foraker, Cheng Chang, Mitch Phelps, and Antara
Banetjee. I am very privileged to work with them all. My sincere appreciation to Karen
Lawler and Kathy Brooks, for their countless assistance during my study. Also, special thanks to Li-Fen Hsu, Mei-Hua Chuang, Teh-Min Hu, Shih-Jiuan Chiu, and Teng-Kuang
Yeh for their friendship and encouragements.
My heartfelt gratitude to Chih-Hsin Chen, my longtime best friend, for always being there and for inspiring me to be a better person.
Mostly, I am very grateful to my parents and grandparents, for their infinite caring and love, for letting me follow my heart desired and for believing in me more than I do myself. My deepest thanks to my brothers, Liang-Chin and Chi-Chun, for their support and for taking care of all my responsibilities while I am abroad. This research was supported by a grant from National Institute of Health (to P.W.
S).
VI VITA
December 25, 1972 ...... Bom - Tainan, Taiwan
1995...... B.S. Pharmacy, National Taiwan University Taipei, Taiwan 1997-present ...... Graduate Research Associate, The Ohio State University
PUBLICATIONS
Research Publications
1. Riboflavin uptake in human trophoblast-derived BeWo cell monolayers: cellular translocation and regulatory mechanisms. Huang, S. -N. and Swaan, P.W. Journal of Pharmacology and Experimental Therapeutics 298(1): 264-271, 2001.
2. Involvement of a receptor-mediated component in cellular translocation of riboflavin. Huang, S. -N. and Swaan, P.W. Journal o f Pharmacology and Experimental Therapeutics 294(I): 117-125, 2000.
3. Intracellular trafficking of riboflavin-rhodamine conjugates, Huang, S. -N., Swaan, P. W. PharmSci 2(4): S-4I49, 2000.
4. Mechanism and regulation of riboflavin uptake in a human placental choriocarcinoma cell line, Huang, S. -N., Swaan, P. W. PharmSci 2(4): S-2014,2000.
5. Involvement of a receptor-mediated endocytosis component in the intestinal transport of riboflavin. Huang, S. -N., Swaan, P. W. PharmSci 1(4): S-3472,1999.
6. Transepithelial transport mechanism of riboflavin in a human small intestinal cell line, Caco-2. Huang, S. -N., Swaan, P. W. FASEBJ 13(4 pt I): A73, 1999.
7. Transepithelial transport mechanism of riboflavin across Caco-2 cell monolayers. Huang, S. -N., Swaan, P. W. PharmSci 1:S453, 1998.
vu FIELDS OF STUDY
Major Field: Pharmacy
V lll TABLE OF CONTENTS Pages Abstract...... ii
Dedication ...... iv
Acknowledgments ...... v
Vita...... vii
List o Tables ...... xi
List of Figures ...... xii
Chapters: 1. Introduction ...... 1 1.1 Biomembrane transport ...... 1 1.1.1 Epithelial barriers for drug delivery ...... 1 1.1.2 Absorption enhancement via targeting to membrane transporters ...... 4 Active transport mechanisms ...... 5 Membrane transporter-targeting approach ...... 8 1.2 Riboflavin ...... 13 1.2.1 Biochemical function and clinical significance of riboflavin ...... 13 1.2.2 Absorption and disposition of riboflavin ...... 14 1.2.3 Riboflavin carrier protein ...... 17 1.2.4 Active transport mechanism ...... 19 1.3 Research objectives ...... 27 Reference ...... 28
2. Involvement of a receptor-mediated component in cellular translocation of riboflavin ...... 33 Abstract...... 33 Introduction ...... 35 Methods ...... 38 Results...... 43 Effect of culture time on riboflavin transport ...... 43 Metabolic stability of [^Hj-riboflavin ...... 44 Polarization of riboflavin transport in rat tissue ...... 45 Concentration dependence of riboflavin transport ...... 45 Effect of endocytosis inhibitors on riboflavin transport ...... 46
ix pH-dependent dissociation of surface-bound riboflavin ...... 48 Discussion ...... 50 Reference ...... 62
3. Riboflavin uptake in human trophoblast-derived BeWo cells monolayers; cellular translocation and regulatory mechanisms ...... 65 Abstract...... 65 Introduction ...... 67 Methods ...... 70 Results...... 74 Riboflavin uptake kinetics ...... 74 Substrate specificity of BeWo cell riboflavin uptake ...... 75 Temperature and energy dependence ...... 76 Ion-coupling properties of riboflavin uptake ...... 77 Riboflavin uptake in syncytiotrophoblasts ...... 80 Effect of cyclic nucleotide-dependent pathways on riboflavin uptake 82 Involvement of protein kinase C and calmodulin-mediated pathways 83 Discussion ...... 85 Reference ...... 104
4. Subcellular localization of riboflavin: involvement of endocytic organelles ...... 108 Abstract...... 108 Introduction ...... 110 Methods ...... 112 Results...... 117 Synthesis of rhodamine-riboflavin conjugate ...... 117 Substrate specificity of rhodamine-riboflavin conjugate ...... 118 Internalization and subcellular localization of rhodamine-riboflavin conjugate ...... 118 Colocalization studies with an acidotropic marker, clathrin-heavy chain, rab 5, and FITC-transferrin ...... 120 Colocalization studies with LAMP-1 proteins ...... 121 Discussion ...... 123 Reference ...... 138
5. Summary and prospects ...... 140 Reference ...... 146
Bibliography...... 147 LIST OF TABLES
Table Page
1.1 Ligands that are transported via carrier-mediated pathways ...... 10 1.2 Ligands that are entered via receptor-mediated pathways ...... 11 1.3 Examples of transporter-targeting approach ...... 12 1.4 Intestinal uptake of riboflavin ...... 22 1.5 Renal uptake of riboflavin ...... 24 1.6 Hepatic uptake of riboflavin ...... 24 2.1 Transport parameters of bi-directional transport of riboflavin in Caco-2 cells ...... 55 3.1 Influence of ions and pH on riboflavin uptake in BeWo cell monolayers ...... 93
3.2 Effect of anion-exchange inhibitors and organic anion transport inhibitors on riboflavin uptake in BeWo cell monolayers ...... 94
3.3 Effect of pCPT-cGMP on riboflavin uptake in BeWo cell monolayers ...... 95
3.4 Effect of protein kinase C and calmodulin-mediated pathway modulators on riboflavin uptake in BeWo cell monolayers ...... 96
XI LIST OF FIGURES
Figure Page
1.1 Transepithelial transport mechanism...... 2
1.2 Receptor-mediated endocytosis mechanism ...... 7
1.3 Metabolic pathway of riboflavin in mammalian tissue ...... 16
2.1 Effect of culture time on riboflavin transport in Caco-2 cell monolayers ...... 56
2.2 Stability of [^H]-riboflavin in 14-day postseeding Caco-2 cells ...... 57
2.3 Time course of riboflavin transport in rat ileum ...... 58
2.4 Concentration dependence of transport of riboflavin in Caco-2 cell monolayers 59
2.5 Effect of nocodazole and brefeldin A on transport and uptake of riboflavin, cholic acid and FITC-transferrin in Caco-2 cell monolayers ...... 60
2.6 pH dependence of riboflavin-specific binding to cell surface ...... 61
3.1 Time course of [^H]-riboflavin uptake in BeWo cell monolayers ...... 97
3.2 Concentration dependence of [^H]-riboflavin uptake in BeWo cell monolayers 98
3.3 Effect of riboflavin structural analogs on uptake of [^H]-riboflavin ...... 99
3.4 Time-dependent effect of forskolin and 8 -bromo-cAMP on [^H]-riboflavin in BeWo cell monolayers ...... 100
3.5 Forskolin-induced differentiation of BeWo cells ...... 101
3.6 Effect of isobutylmethylxanthine (IBMX) on [^H]-riboflavin in BeWo cell monolayers ...... 102
3.7 Effect of calmodizalium on [^H]-ribofIavin in BeWo cell monolayers ...... 103
xn 4.1 Synthesis of rhodamine-riboflavtn^ conjugate ...... 128
4.2 Effect of rhodamine-riboflavin conjugate and rhodamine on riboflavin uptake 129
4.3 Internalization of rhodamine-riboflavin ...... 130
4.4 Internalization of FITC-transferrin, rhodamine or riboflavin ...... 131
4.5 Specific binding of rhodamine-riboflavin ...... 132
4.6 Colocalization of rhodamine-riboflavin with acidotropic probe LysoTrackerBlue- White-DXD...... 133
4.7 Colocalization of rhodamine-riboflavin with clathrin ...... 134
4.8 Colocalization of rhodamine-riboflavin with rab 5 protein ...... 135
4.9 Colocalization of rhodamine-riboflavin with transferrin ...... 136
4.10 Colocalization of rhodamine-riboflavin with LAMP-1 proteins ...... 137
xm CHAPTER 1
INTRODUCTION
1.1 BIOMEMBRANE TRANSPORT
1.1.1 Epithelial barriers for drug delivery
To exert their desired pharmacological activities, drugs must reach their sites of
action with certain minimal effective concentration. With the exception of a few drugs
(e.g. general anesthetics, osmotic diuretics), most drugs produce their effects by acting on
specific membrane proteins or intracellular enzymes (Neal, 1992). Since most of the
human body surfaces are covered with epithelium, to gain access to systemic circulation
or their cellular targets, drugs must first penetrate the epithelial barriers.
In most biological epithelia, transport of drug molecules is confronted by two
obstacles; 1) a biochemical barrier resulted from enzymatic degradation; 2) a physical
barrier originated from the lipid bilayer. The first challenge can be significantly circumvented by changing routes of administration and/or formulations, e.g. encapsulating drugs in vehicles. However, for hydrophilic drugs and drugs with high molecular weight, especially macromolecules, epithelial membrane still imposes an ultimate obstacle for entry of drugs. The epithelium in different tissues^ may vary in thickness or functions, but the
general transepithelial transport mechanisms for drug molecules are the same (Fig. 1.1).
Based on the route that drug molecules penetrate, epithelial transport can be classified
into two pathways: paracellular and transcellular. In the paracellular pathway, molecules
move across the epithelium via the intercellular junctions between adjacent cells, wheras in the transcellular pathway, molecules cross the epitheliuam through the cells.
Depending on nature of the driving forces, transcellular pathways can be further categorized into passive diffusion and active transport. In passive diffusion, movement of drug molecules is drived by its concentration gradient. In active transport, molecules are transported against a concentration gradient using an energy supply provided by either
ATP hydrolysis or co-transport of ions.
rYY r m
@
ATP
Figure 1.1 Transepithelial transport mechanism.
1.Paracellular pathway 2-6. Transepithelial pathways: 2. Passive diffusion 3. Facilitated diffusion 4. Carrier-mediated transport 5. Receptor-mediated endocytosis/transcytosis
6. Efflux pump In general, drug transport across any epithelium is dictated by the characteristics of the cell membrane and the physicochemical properties of drugs. In absorptive epithelium such as enterocytes, intercellular space is sealed by tight junctions or zonula occludens. These Junctional proteins play two essential roles in the epithelial barrier functions. They not only maintain the cell polarity by confining surface proteins to their appropirate membrane domains, but also prevent diffusion of water-soluble molecules and backflow of the absorbed nutrients (Alberts et al., 1994). Studies in rat small intestines have shown that only water and small hydrophilic solutes with molecular radius smaller than 25 Â can move across the paracellular pathway (Pappenheimer, 1987).
The overall surface area available for transcellular transport is significantly larger than that of the paracellular pathway. Therefore, the transcellular pathway is naturally the preferred route of transport for most molecules. Lipophilic and small amphiphilic compounds can traverse the epithelium efficiently by partitioning into and out of the lipid bilayers. In constrast, large hydrophilic molecules cannot diffuse freely through the cells even when thermodynamic conditions e.g., concentration gradient, favor such action.
Factors influencing the transcellular passive diffusion of drugs have been thoroughly characterized. It is predicated that only drugs that have a molecular weight less than 500,
Log P less then 5, less than five hydrogen bond donors and ten hydrogen bond acceptors
(the rule o f 5), are more likely to permeate across the cell membrane passively (Lipinski etal., 2001). t.t.2 Absorption enhancement via targeting tn membrane transporters
Several strategies have been developed to enhance the bioavailability of drugs
with limited membrane permeability. These approaches can be categorized into methods
that manipulate the membrane barrier properties or increase drug solubility. Permeation- enhancing agents including compounds such as surfactants, bile salts, chelating agents or short-chained fatty acids, have been used to improve transport of poorly-absorbed drugs
(Lee, 1991). Most agents enhance uptake of drugs by compromising the integrity of cell membrane. Generally, the increased absorption results from either disrupting the tight junctions or altering the membrane fluidity or both. However, due to their nonspecific mechanisms of action and the safety concerns regarding mucosal damage, the widescale application of these penetration enhancers has not happened.
An alternative approach to increase drug permeation is to increase the lipophilicity of drugs through chemical modifications. In general, a prodrug is synthesized from the parent compound by converting its hydrophilic residues into less polar moieties. During or after absorption, the parent drugs are then released from their lipophilic derivatives by hydrolysis or specific enzymatic actions. This strategy has successfully improved absorption of numerous drugs, especially for compounds with relatively small molecular weight. However, not all poorly absorbed drugs can be subjected to structural modifications. Moreover, lipophilization could be potentially problematic for delivery of macromolecules such as polypeptides and proteins, whose structures are closely related to their biological activities.
Whereas charged, hydrophilic compounds and pharmaceutical macromolecules encounter difficulties in permeating the cell membrane, the systemic absorption of many water-soluble nutrients (e.g., sugars, vitamins), endogenous proteins (e.g., insulin, growtb factors), and toxins (e.g., ricin, cholera toxin) appears to be highly efficient. The effective transcellular movement of these molecules is facilitated by specialized transport processes in the epithelia, namely carrier-mediated transport and receptor-mediated endocytosis/transcytosis (Fig. 1.1). Table 1.1 and 1.2 summarized a variety of ligands that are taken up by these two pathways. Both processes are operated by specific membrane associated proteins and share common features of active transport mechanisms, i.e., concentration-, energy-, temperature-dependent, and subjective to structural analog inhibition. On the other hand, they also differ significantly in ways that transporter proteins are anchored in membrane and in their ligand internalization mechanisms.
1.1.2.1 Active transport mechanisms
Carrier-mediated pathways
Carriers are integral membrane proteins that associate with the lipid bi layer via multiple membrane spanning domains (usually ranging from seven to twelve). The molecular details by which a carrier protein operates the transcellular movement of a solute are unknown. Thermodynamically, due to their multipassed transmembrane features, carriers are highly unlikely to “carry’' ligands back and forth across the membrane (Alberts et al., 1994). It is generally believed that carrier proteins shuttle their substrates across the lipid bi layer by undergoing reversible conformational changes.
Briefly, ligand binding to the substrate-binding site exposed on the outside of the membrane induces a conformational change of carrier proteins, which leads to subsequent release of ligands to the opposite side of membrane. Most carriers are either directly energized by ATP hydrolysis (e.g., p-glycoprotein
pumps) or powered by the energy stored in ion gradients. In mammalian cells, uptake of
substrates is usually driven by the free energy released during the movement of Na^, fT
or OPT down an electrochemical gradient (Hedigeretal., 1995).
Receptor-mediated endocytosis pathways(RME)
In contrast, receptor proteins usually have far fewer membrane-spanning loops,
with some having only a single membrane anchorage domain (e.g., glycosyl-
phosphatidyl-inositol-anchored receptors). Compared with carrier-mediated pathways,
RME operates by a more complicated mechanism involving several cellular components
(Fig. 1.2) (Mukhetjee et al., 1997).
In mammalian cells, RME can occur via clathrin- dependent or clathrin-
independent pathways. In general, ligands bind to receptors located on specialized regions of the membrane (e.g., clathrin-coated pits or caveolae). Through a membrane invagination process, ligand-receptor complexes are delivered into the peripheral early sorting endosomes that are acidified by an ATP-dependent proton pump. Following the internalization process, the endocytosed cargo can proceed into one of these different scenarios. I) Many receptors and ligands dissociate from each other upon exposure to the acidic pH in the early endosome. The receptors are recycled back to the cell surface whereas the ligands are delivered to late endosomes or lysosomes for degradation (e.g.,
LDL receptor). 2) Some ligand-receptor complexes are unaffected by the acidification and both are sorted to the lysosomes (e.g., insulin receptor). 3) Some ligands remain associated with their receptors and both are returned to the cell surface (e.g., transferrin). 4) Some receptors and ligands are both transcytosed tathe opposite side of membrane
and released to the extracellular milieu (e.g., IgA) (Alberts et al., 1994; Swaan, 1998).
Endosomes are very dynamic entities. As shown in Figure 1.2, there are extensive
and rapid traffic among different endosomal organelles. Cytoskeleton and many cellular
proteins e.g., RAB and t-SNARE proteins are involved in regulation of endocytic
processes. In polarized cells such as enterocytes, the RME pathways are generally more complex than in the non polarized cells. It is known that distinct sets of endocytic components are involved in endocytosis from the apical and basolateral surfaces.
CM tM ptf
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■ wyi»
vscr vtmao vwiMe VBU99
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Figure 1.2 Receptor-mediated endocytosis. (from Sorkin A., 2CXX)) 1.1.2.2 Membrane transporter-targeting approach
Apart from naturally occurring substrates, it is now well recognized that many drugs can be selectively taken up by the active transport processes (Table 1.1). For pharmaceutical scientists, these membrane transporters provide alternative routes for the delivery of drugs that would normally be impermeable to the biological barriers. Utilizing a method similar to the conventional prodrug approach, absorption enhancement is pursed by formation of conjugates between drugs and the endogenous ligands of the membrane transporters. Consequently, via the specific interaction ligands between the moiety and its transporter, drug candidates can be shuttled across or into the cells and eventually be released from the ligands.
Taking advantages of recent advances in molecular biology and computer modeling, scientists are now able to design prodrugs based on the structural requirements of the transposer systems. Several successful examples targeting both types of active transport processes are summarized in Table 1.3. In general, prodrug strategies involving carrier-mediated pathways have the advantage of high uptake capacity. However, the size of drug conjugates is relatively limited (-600 Da) probably because larger conjugates fail to be shuttled through the restricted space within the carrier protein. For peptide and protein delivery, carrier-mediated pathways could only facilitate peptides up to four amino acids (Tamai and Tsuji, 1996; Chen et al., 1999).
Compared with carrier-mediated pathways, receptor-mediated endocytosis systems have a rather limited uptake capacity, which in some cases is insufficient to deliver enough drug to elicit pharmacological activities. Yet, because of the endocytic pit
8 formation (up to several hundred nanometers) and vesicular internalization mechanism,
RME pathways are perfectly suited to accommodate large molecular weight peptide
and/or protein conjugates. More importantly, recent success in transport of RME ligand-
drug vehicle conjugates (e.g., nanoparticles, liposomes) via RME pathways opens new
possibilities for macromolecular delivery across biological barriers. First of all,
formulating pharmaceuticals in drug vehicle systems compensates the limited capacity of
RME systems, resulting in 10^ to 10*^ -fold increase in uptake. Secondly, drug vehicle
systems also protect drug molecules from possible enzymatic degradation in the
biological membrane. Furthermore, this type of conjugation avoids direct chemical reaction between drug molecules and ligands, allowing incorporation of drugs with more diverse structural properties. Carrier name Natural substrate Known drug substrate
Hexose* D-glucose, D-galactose Phenyl-P-D-xylopyranoside, arbutin
Amino acid L-Amino acids L-dopa, L-a-methyldopa, baclofen, gabapentin Peptide** Di-, tri-peptide Penicillins, cephalosporins, ACE inhibitors, alafosfalin, bestatin Nucleoside* Nucleoside AZT, ddC, ddl
Monocarboxylic acid** Lactic acid, short-chain fatty Salicylic acid, p-aminobenzoic acid acid HMG-CoA reducase inhibitor Dicarboxylate Succinate, citrate, a- ketoglutarate Phosphate* Inorganic phosphate Foscamet, phosphomycin Biotin* Biotin Carbamazepin, primidone
Folic acid Reduced folates Methotrexate
Multi-vitamin* Panthothenate, lipoate, biotin
Nicotinic acid Nicotinic acid
Bile acid* Bile acid Somastatin analogs, cyclosporine, fusidic acid.
Table 1.1 Ligands that are transported via carrier-mediated pathways (Tamai and Tsuji, i996)
* Na^ dependent, ** H^-dependent Hormones and Toxins and lectins Viruses and Serum proteins Vitamins growth factors bacteria and antibodies Calcitonin Cholera toxin Adenovirus IgE Folate Catecholamines Concanavalin A Reovirus IgG, via Fc receptor Vitamin Epidermal growth factor Diphtheria toxin Rotavirus Low density Glucagon E. coli heat labile toxin Rous sarcoma virus lipoprotein Growth factor Pseudomonas toxin Semliki forest virus Maternal IgG Insulin Staphy. Enterotoxin A & B Varicella zoster Polymeric IgA Interferon Ricin Vesicular stomatitis Transcobalamin Luteinizing hormone Toxic shock syndrome toxin I virus Transferrin Nerve growth factor Yolk proteins Platelet derived growth factor £. coli Prolactin Enterobacter strains Thyroid stimulating hormone L piantarum Thyroid hormone Klebsiella strains Serratia strains V. cholerae
Table. 1.2 Ligands that are entered via receptor-mediated endocytosis (Pastan and wiiiingham, i98S; Swaan, i998) Transporter Conjugates Advantages Disadvantages Carrier-mediated pathways Peptide transporter Enalaprilat Broad substrate specificity Maximal size of transported (PepTI) L-a-methyldopa Parent compound can be liberated peptide: tripeptide by cytosolic enzymes Bile acid transporter Chlorambucil High uptake capacity (20g per day) Maximal size of transported HMG-CoA reductase Liver-specific targeting peptide: tetrapeptide; L-triiodothyronine Unable to release parent compound from conjugates
Receptor-mediated endocytosis pcdhways IV Folate receptor Toxins Tumor-specific targeting; Low capacity Radioimaging agent Able to transport very large Liposomes molecular weight Vitamin B,? receptor GCSF Transcyotsis, transport to systemic Low capacity (40 fmol per EPO circulation; day) LHRH agonists Able to transport very large Vasopressin molecular weight Nanoparticles
Table 1.3Examples of transporter-targeting approach (Swaan, 1996; Waiter ct at., 1996; Reddy and Low, 1998; Russell-jones, 1998) Note: G-CSF: granulocyte colony stimulating factor; EPO: erythropoietin;,LHRH :luteinizing hormone t.2 RIBOFLAVIN
Since its discovery in 1920s, riboflavin, also known as vitamin Bz, has been one
of the main focuses in vitamin research. Overwhelming numbers of studies have
contributed to substantial information in the chemistry, physiology, and medical
significance of this essential vitamin (Rivlin, 1975). Cellular homeostasis of vitamins is
tightly controlled in the body due to their unique and indispensable roles in normal
cellular function, growth and development (Ferraris, 1994). Thus, to address the topic on
the biological transport phenomenon of riboflavin, it is equally important to understand
its physiological properties and metabolic processes. In the following sections, a brief
overview of absorption and disposition of riboflavin is presented with a special emphasis
on the current knowledge of its cellular translocation mechanism.
1.2.1 Biochemical function and significance of riboflavin
In its coenzyme forms of flavin adenine dinucleotide (FAD) and riboflavin 5’-
phosphate (FMN), riboflavin performs as an electron transfer intermediary in biological oxidation-reduction reactions. Flavoproteins, enzymes containing FMN or FAD as cofactors, catalyze a remarkable spectrum of biological processes in both prokaryotic and eukaryotic cells. They are not only essential for biosynthesis and metabolism of carbohydrate, lipid, and amino acid, but also crucially involved in activation of other vitamins such as pyridoxine and folic acid (Cooperman and Lopez, 1991; Voet and Voet,
1995).
In human, riboflavin deficiency results in dermatitis, angular stomatitis, cheilosis, and neuropathy (Cooperman and Lopez, 1991). Despite of its important involvement in
13 metabolic pathways, these overt ctmicat signs of riboflavin deficiency are rare among
inhabitants of the developed countries. However, about 28 million Americans exhibit a
common “sub-clinical” deficiency stage, characterized by a change in biochemical
indices (e.g., erythrocyte glutathione reductase) (Lemoine et al., 1980). In this stage, such
person’s dietary uptake is sufficient to prevent clinical manifestations but adequate to
sustain flavoproteins for their optimal activities. Although the long-term effects of this
sub-clinical deficiency are unknown in adults, this deficiency results in growth
retardation in children (Goldsmith, 1975). Moreover, a severe primary riboflavin
deficiency is often followed by secondary deficiencies of other vitamin Bs (Pinto and
Rivlin, 1987).
1.2.2 Absorption and disposition of riboflavin
Unlike microorganisms, mammals cannot biosynthesize riboflavin. The only way to obtain this essential vitamin is from diet via intestinal absorption or in the cases of fetus, through maternal sources. Most dietary riboflavin is in the form of flavoproteins or its coenzyme forms, which must be first hydrolyzed to riboflavin before absorption can occur (Fig. 1.3). Digestion of flavoproteins, FAD, FMN occurs by both nonspecific and specific enzymes in the intestine, suggesting a physiologic preference of riboflavin as the absorptive species of dietary flavins.
Upon entry into the cell, most of the water-soluble vitamins are metabolically altered, and, as a consequence, become trapped within the cell (McCormick and Zhang,
1993). Many pieces of evidence indicate that a two-step phosphorylation process is involved in metabolic trapping of riboflavin (Jusko and Levy, 1975; Okuda et al., 1978).
14 Two cytosolic enzymes, fTavokinase andFAD synthetase, have been identified in
enterocytes and hepatocytes to catalyze these specific conversions of riboflavin into its
anionic coenzyme forms (Merrill et al., 1978; Yamada et al., 1990). Some studies also
further suggest this phosphorylation process is closely coupled to an absorption
mechanism of riboflavin and is necessary for its intestinal absorption (Kasai et al., 1988),
although direct proof of this mechanism remains elusive. Excessive riboflavin and
catabolic metabolites of intracellular flavoenzymes could exit the enterocytes via a
basoiateral-located transporter.
In humans, approximately 50% of total riboflavin in plasma is protein-bound.
Circulatory transport of riboflavin is known to involve both weak associations with
albumin and tight binding to a subclass of immunoglobulins (McCormick, 1989). In oviparous vertebrates, a class of pregnancy specific, estrogen-induced riboflavin-carrier proteins is found in transporting riboflavin to the embryonic tissue (Adiga, 1994).
Recently, a similar protein has also been identified in mammals including humans. A more detailed review of this protein will be presented in section 1.2.3.
After reaching systemic circulation, riboflavin is imported and converted into
FMN and FAD inside the tissues. The liver is the major storage site, containing about one-third of total body flavins. Excessive amounts of riboflavin are excreted primarily in the urine, mostly in its intact form. Both in vitro and in vivo studies have demonstrated that after glomerular filtration, riboflavin is secreted and reabsorbed by the renal tubules via saturable processes (Jusko and Levy, 1975; Yanagawa et al., 1997; Yanagawa et al.,
1998).
15 Dietary ______Gastric acidation S-cyteiny-flavins, N-histidyl-flavins flavoproteins protease FAD, FMN FAD pyrophosphase ^Alkaline phosphatase FMN phosphatase Alkaline phosphatase Riboflavin (Rf) FMN rrY rY Y ^ Intestinal mucosa
Flavokmase
Flavoenzymes
FAD synthetase
FAD > Flavoenzymes
Circulation Protein binding; Albumin & other Rf binding proteins Liver
Rf Ravokinasc ^
FMN
FAD synthetase ^
FAD
Figure 1.3 Metabolic pathway of riboflavin in mammalian tissue.
Effect o f drugs on disposition of riboflavin
The structure of riboflavin consists of two main parts: one isoalloxazine ring and a D-ribose side chain. Interestingly, several groups of therapeutic compounds are structurally related to riboflavin, for example, phenothiazine derivatives like
16 chiorpromazineand tricyclic antidepressant agents (TGAs>such as imipramine and
amitriptyline. Co-administration of these agents has been shown to enhance urinary
excretion of riboflavin and to accelerate tissue depletion of FAD levels in liver
(Pelliccione et al., 1983; Pinto and Rivlin, 1987). Since psychoactive drugs that are
structurally unrelated to riboflavin fail to produce comparable outcome, these studies
suggested that the phenothiazines and TCAs might exert their inhibitory effects on the
cellular disposition of riboflavin. Although the exact mechanism of these drugs remains
unclear, it is likely that these metabolic changes are caused by modulation of riboflavin-
specific enzymes and/or membrane translocating systems.
OH OH
OH .O
.NH
HjC
Riboflavin Chlorpromuzine
Amilriptyline Imipramine
1.2.3 Riboflavin Carrier Protein
During pregnancy, rapid embryonic development demands a continuous supply of nutrients including riboflavin from the maternal system. However, like most epithelium, the placental/vitelline membrane prevents the transit of free riboflavin into the fetus/egg.
17 In oviparous vertebrates, a specific riboftavin carrier protein (RCP) is involved in
transport of riboflavin from plasma to oocytes. The best characterized of the RCPs,
chicken RCP (cRCP), is a 37 kDa phosphogiycoprotein synthesized in liver and oviduct.
RCP is unique among flavoproteins because it preferentially binds to riboflavin (in 1:1
molar ratio) over its coenzyme forms (Adiga, 1994). Although attempts to locate the
cRCP-specific receptors have been inconclusive, cRCP-mediated delivery of riboflavin in
eggs has been shown to occur via endocytosis of the cRCP-riboflavin complexes through
the lipoprotein receptor (Mac Lachlan et al., 1994).
Using antibodies raised against cRCP, Adiga and coworkers are able to identify
RCP in the plasma of pregnant mice, rats, monkey, and women. Although these carriers
have not been characterized thoroughly, the rodent and primate RCP display marked similarities to cRCP with regard to size, affinity, isoelectric point, and immunological cross-reactivity. In humans, the distribution and role of an RCP is more elusive. One study reports the presence of a binding protein in the amniotic fluid of pregnant women.
RCPs have also been detected in plasma of male monkeys as well as rat testicular Leydig and Sertoli cells. Similar to their female counterparts, RCP production in male rats and monkeys was sensitive to estradiols-lVP and follicle stimulating hormone.
Recently, two separate groups have independently shown elevated serum RCP levels in breast cancer patients (Karande et al., 2001; Ramesh and Meenakshi, 1996). The rising serum RCP seems to correlate with the progression of the breast cancer with the advanced breast cancer patients harboring the highest RCP levels than the early breast cancer patients and normal disease-free women (Karande et al., 2001).
Immunohistochemical analysis in breast cancer biopsy specimen also detected the
18 presence of RCP in the cytoplasma of neoplastic cells. Although it remains to be verified
whether the elevated patient RCP is resulted from overexpression of RCP in breast cancer
cells, these studies suggest that circulatory RCP could present as a promising marker for
breast cancer diagnosis and prognosis.
In summary, although RCP have been identified in plasma and reproductive
organs of either sex, its function in mammalian riboflavin transport remains to be defined.
Nevertheless, based on the remarkably conserved features between cRCP and
mammalian RCPs, it is proposed that a similar RCP-mediated mechanism could also
exist in fetolplacental transport of riboflavin (Adiga, 1994).
1.2.4 Active transport mechanism
Riboflavin is the preferred absorption species among all flavins, but its low
partition coefficient (Log Poctanoi = -1.46) prevents its permeation across cell membrane
by simple passive diffusion. Contradictorily, when riboflavin is given orally in human
subjects, maximal plasma concentration and urinary excretion are observed within the
first 1.5 hr, indicating a rapid absorption and/or elimination process (Jusko and Levy,
1967; Zempleni et al., 1996). The efficient disposition together with a dose-dependent
absorption strongly suggests the involvement of a specialized pathway in cellular
translocation of riboflavin.
Several in vitro techniques such as everted sacs, membrane vesicles, and isolated
cell preparations have been applied to directly investigate the specialized transport process of riboflavin. An active transport mechanism has been observed in uptake of riboflavin in intestine, liver, kidney, choroid plexus of species rat, rabbit, and human.
19 Intestine
Because of its pivotal role in absorption of riboflavin, the intestinal uptake
mechanism of riboflavin is the most extensively studied among all tissues (Table 1.4).
Taken together, these results suggest that riboflavin is taken up by enterocytes via an
active, saturable, temperature-dependent carrier-mediated mechanism.
Affinity
The high affinity specialized intestinal uptake process has a Km value
within a lower pM range (<10 pM). The riboflavin transporter system in rat
intestine exhibits a lower capacity but an approximately 10-fold higher affinity
compared to that in rabbit intestine.
Site specificity and distribution
Early pharmacokinetics studies suggest riboflavin is absorbed more
rapidly in the proximal small intestine (Jusko and Levy, 1967). However, several
recent studies showed similar level of uptake activity in both jejunum and ileum
(Kasai et al., 1988; Said and Mohammadkhani, 1993).
Intestinal absorption of riboflavin involves vectorial transport across both
apical and basolateral cell membrane. Using rabbit intestinal membrane vesicles.
Said and co-workers reported the existence of a specialized carrier-mediated
process on basolateral membrane of enterocytes (Said et al., 1993a).
2 0 Co-transporter properties
The influence of on riboflavin uptake in intestine remains
controversial: although some studies report Na"^ dependency, most reports indicate
riboflavin transport to be sodium-independent. Furthermore, riboflavin uptake is
mostly insensitive to ouabain, which confirms the apparent Na^-independent
behavior of this transport system.
Regulation
In both rat intestine and Caco-2 cells. Said and co-workers have shown
that intestinal uptake of riboflavin is closely regulated by its dietary level (Said
and Mohammadkhani, 1993; Said and Ma, 1994). Riboflavin uptake in
riboflavin-deficient intestine and Caco-2 cells is up-regulated with significant
increase in their Jmax values without affecting Km of the systems.
In Caco-2 cells, treatment of intracellular cAMP modulators results in
significant inhibition of riboflavin uptake, suggesting possible involvement of a
protein kinase A (PKA)-mediated pathway(s) in regulation of the transporter
activity (Said et al., 1994).
2 1 Species Techniques Km Jnm Properties Rat BBMV 0.12 pM 0.36 pmol/5 sec /mg protein Na*-independent BBMV (jejunum) 0.38 nM 0.9 pmol/5 sec/mg protein Na*-dependent pH-dependent
Everted sacs 0.177 uM 25.8 pmol/min/lOOmg tissue Na*-dependent
Everted sacs 0.54 uM 0.182 pmol/min/cm" Na*-dependent Ouabain sensitive
Single pass perfusion 0.38 pM 12 pmol/min/g tissue Rabbit BBMV (jejunum) 7.24 pM 24.3 pmol/5 sec/mg protein Na*-independent BBMV (ileum) 8.88 pM 32.2 pmol/5 sec/mg protein pH-independent Probenecid, PAH insensitive
BLMV (jejunum) 5 pM 91.6 pmol/5 sec/mg protein Na*-independent BLMV (ileum) 4.4 pM 60.8 pmol/5 sec/mg protein pH-independent Probenecid, PAH insensitive
Human BBMV (jejunum) 7.26 pM 0.48 pmol/5 sec/mg protein Partial Ha* dependent Electrogenic uptake
Caco-2 cell line 0.3 pM 210 pmol/3min/mg protein Na*-independent Ouabain insensitive pH-independent Probenecid, PAH insensitive
Frog Xenopos oocytes 0.41 pM 2.86 fmol/hr/oocyte Na*-independent Ouabain insensitive Probenecid, PAH insensitive BBMV: brush border membrane vesicle; BLMV: basolateral membrane vesicle; PAH: p-aminhippurate
Table 1.4 Intestinal uptake of riboflavin (Daniel et al., 1983; Middleton, 1990; Feder et al., 1991; Said and Ananas, 1991; Daniel and Rehner, 1992; Said and Mohammadkhani, 1993; Said et al., 1993b; Casirola et al., 1994; Said and Ma, 1994; Dyer and Said, 1995)
2 2 Kidney
In renal tubules, riboflavin is imported bi-directionally via a carrier-mediated
process (Table 1.5). In term of Na^ and pH dependency, these two systems exhibit
different co-transporter properties (Yanagawa et al., 1997; Yanagawa et al., 1998).
Compared with that of intestine, the renal riboflavin transporter systems have relatively
lower affinity but higher capacity.
In early in vivo studies in human, probenecid, a substrate of organic anion
transporter, was shown to inhibit renal excretion in a dose-dependent fashion (Jusko et al.,
1970). Consistent with this result, renal uptake in most in vitro studies are found to be inhibited by probenecid and other organic acids such as p-aminohippurate.
Using HK-2 cells, a human renal proximal tubule epithelial cell line, Kumar and colleagues demonstrated renal uptake of riboflavin is also adaptively regulated according to riboflavin supplement (Kumar et al., 1998). However, up-regulation of riboflavin uptake in riboflavin-deficient HK-2 cells is accompanied by increase of both the apparent
Km and Jmax values. Moreover, unlike Caco-2 cells, riboflavin uptake in HK-2 is under the regulation of a Ca‘V calmodulin mediated pathway(s).
23 Species Techniques Km Jmax Properties Rat Isolated cells 8.4 pM 7 pmol/30sec/10 * cells Na*-dependent Ouabain insensitive PAH insensitive Rabbit BBMV 25.7 pM 76 pmol/IOsec/mg protein Partial Na*-dependent pH-independent Probenecid, PAH sensitive BLMV 8.3 pM 14.3 pmol/IOsec/mg protein Na*-independent pH-dependent Probenecid, PAH sensitive Human HK-2 cell line 0.67 pM 10 pmol/3min/mg protein Na*-independent Ouabain insensitive Probenecid, PAH sensitive
PAH: p-aminhippurate
Table 1.5 Kidney uptake of riboflavin (Bowers-komro and McCormick, 1987; Yanagawa et al., 1997; Kumar et al., 1998; Yanagawa et al., 1998)
Liver
The liver is the major storage site of riboflavin and plays an important role in
riboflavin homeostasis. Hepatic uptake of riboflavin proceeds by a carrier-mediated
process. Riboflavin import in Hep G2 cells, a human derived liver cell line, is regulated
by riboflavin levels in the growth medium and a Ca'Vcalmodulin-mediated pathway
(Said et al., 1998).
Species Techniques Km Jmax Properties Rat BLMV 3.55 pM 39.9 pmol/5sec/mg protein Na^-independent pH-independent Probenecid, PAH insensitive
Isolated hepatocytes 11.8 pM 81.7 pmol/min/ 10^ cells
Human Hep 0 2 cell line 0.41 pM 3.6 pmol/3min/ mg protein Na^-independent
Table 1.6 Hepatic uptake of riboflavin (Aw et al., 1983; Said et al., 1995; Said et al., 1998)
24 Choroid plexus and brain
Unlike in other tissues, the concentration of riboflavin in brain is maintained
relatively constant. Since this essential factor is not biosynthesized in brain cells, it must
enter brain and cerebrospinal fluid (CSF) from the blood. Choroid plexuses, which
comprise the blood and CSF barrier, together with blood brain barrier, are responsible for
controlling the nutrient homeostasis in the brain. A saturable, Na'^-dependent, ouabain-,
probenecid- sensitive carrier-mediated uptake process has been reported by Spector and
colleagues (Km: 78 pM; Jmax: 1.66 mmol/kg/15min) (Spector and Boose, 1979; Spector,
1980).
Summary
From the above studies, it is generally concluded that riboflavin enters cells via an
active carrier-mediated pathway. The mechanistic features of riboflavin uptake seem to be different among various epithelial types. Currently, based on these results it is unclear whether distinct carrier protein is responsible for the uptake process in different tissues.
The discrepancy in affinity and capacity of these uptake systems could be contributed by
1) species difference (as reflected in the intestine studies), 2) difference in models or tissue preparation methods. Interestingly, the riboflavin uptake systems in intestine and kidney are very different in its sensitivity towards organic ion transporter inhibitors e.g., probenecid and in its signal transduction regulatory mechanisms.
Most carriers are found to be directly coupled to an electrochemical gradient of secondary ion. Transport of acidic vitamin is also thought to be primarily via a sodium-
25 dependent process (see Table 1.1% Since nboflavin has an isoeiectric point of pH 6.0, a majority of riboflavin molecules are as neutral species at physiological pH. While conflicting results were obtained regarding the effect of Na^ and pH on riboflavin uptake, the Na^- and pH-independent feature obtained in most studies is consistent with this prediction.
Riboflavin-enhanced delivery of macromolecules
Recently, Low and co-workers reported the facilitated entry of bovine serum albumins (BSA) into several human pulmonary and ovary cell lines and lung epithelial preparations after they covalently coupled BSA to riboflavin (Wangensteen et al., 1996;
Holladay et al., 1999). In in vivo studies, after intratracheal instillation of BSA-riboflavin,
2.1 times more BSA-riboflavin than unconjugated BSA was found in the plasma. Similar extent of enhancement has also been shown in delivery of BSA-riboflavin conjugates in isolated perfused lungs. Furthermore, their studies suggest that BSA-riboflavin conjugates enter the cells via by an endocytosis/transcytosis mechanism, since I) the facilitated uptake was reduced to the control level with addition of transcytosis inhibitors, and 2) the conjugates were detected in endosomal-like compartments. Interestingly, they also found the cell association of BSA-riboflavin is inhibited by the unlabeled conjugates, but not by free riboflavin while the cellular uptake of riboflavin is only partially inhibited by riboflavin-BSA. These data further suggest that the endocytosis pathway may not be the sole mechanism of cell entry available to riboflavin.
26 1.3 RESEARCH OBJECTIVES
As discussed above, the complete active cellular translocation mechanism of riboflavin remains to be defined. The apparent uniport behavior (Na^, pH independency) of riboflavin transport system contradicts our current knowledge that all nutrient membrane carriers are co-transporters (Hediger et al., 1995). The involvement of receptor-mediated endocytosis in riboflavin absorption remains hypothetical and is yet to be elucidated. It is the aim of this thesis to unravel the receptor-mediated endocytosis components in riboflavin translocation and to further exploit the riboflavin transport system as potential means for drug delivery. Specially, major objectives of these thesis studies include to:
1. Characterize the transepithelial transport and uptake kinetics of riboflavin,
2. Define the structural requirements of the riboflavin transport system,
3. Assess the involvement of receptor-mediated endocytosis events and to
identify intracellular compartments.
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32 CHAPTER 2
INVOLVEMENT OF A RECEPTOR-MEDIATED COMPONENT IN THE
CELLULAR TRANSLOCATION OF RIBOFLAVIN
ABSTRACT
This study addresses the transport mechanism of riboflavin (vitamin Bi) across intestinal epithelium in the presence and absence of pharmacologically active compounds.
A polarized transport process with a 6-fold higher basolateral (BL)-to-apical (AP) flux was observed in both a human intestinal cell model (Caco-2) and rat intestinal tissue.
Riboflavin-specific translocation systems on both the apical and basolateral cell surfaces were saturable with affinity values close to most receptors (Km: 9.72 ± 0.85,4.06 ± 0.03 nM, respectively). Pharmacological agents known to alter intracellular endocytic events were used to examine the potential involvement of receptor-mediated events. Nocodazole significantly inhibited apical uptake (58.4%), BL-to-AP riboflavin (56.7%) and
FITC-transferrin (31.8%) transport without affecting mannitol or cholic acid transport, whereas AP-to-BL riboflavin (152.8%) and FITC-transferrin (45.1%) transport was increased. Brefeldin A significantly enhanced AP-to-BL riboflavin (37.1%) and bi-directional FTTC-transferr.n transport (AP-to-BL: 13 fold, BL-to-AP: 5 fold), without affecting BL-to-AP riboflavin transport. Combined, these data suggests an essential role of
33 microtubule-dependent movement and vesicular sorting component(s) in the bi-directional transport of riboflavin. Dissociation of riboflavin from the cell surface was pH-dependent with significantly higher substrate release at acidic pH, indicating the presence of riboflavin-specific cell surface receptors. In summary, our studies provide biochemical evidence of the involvement of a receptor-mediated mechanism in the cellular translocation of riboflavin.
34 INTRODUCTION
The cell membrane imposes a formidable absorption barrier to the translocation of
water-soluble vitamins. To accommodate the entry of these essential nutrients, the cell
expresses specific membrane proteins on the cell surface. These proteins are part of a
specialized uptake mechanism that can be broadly categorized as carrier-mediated and
receptor-mediated endocytosis. The first mechanism facilitates the movement of vitamin
molecules across the cell membrane via membrane carrier protein(s) energized by ATP
hydrolysis or the co-transport of ions moving down their electrochemical gradient. In fact,
transport of most B vitamins has been identified to occur via a Na^- or lU- dependent
carrier-mediated pathway (Dutta et a i, 1999). In the second model, the vitamin molecules
may first bind to an endogenous protein that in turn binds to surface receptors (e.g. vitamin
B 12) or directly binds receptors localized in specialized membrane regions (e.g. folate)
before they are internalized into endocytic vesicles (Antony, 1996).
Cellular uptake of riboflavin, also known as vitamin B?, has been extensively investigated in a variety of cell lines, and organs and tissues (intestine, liver, kidney) from several species (human, rat, and rabbit) (Rindi and Gastaldi, 1997, Said and Arianas,
1991). From these studies, it appears that riboflavin is taken up into most cells via an active, carrier-mediated mechanism that is pH-independent. It has been suggested that these mechanisms are present on both surfaces of epithelial cells (Said et a i, 1993; Said and Mohammadkhani, 1993). Contradicting results have been reported regarding the influence of Na^ on riboflavin uptake. Although some studies suggest no obvious requirement for Na^ or partial Na^-dependency (Middleton, 1990; Said and Arianas, 1991), most reports indicate riboflavin transport to be Na^-independent (Dyer and Said, 1995;
35 Safcf and Ma, 1994). Furthermore, rîbofTavîn uptake is nor sensiti ve to the Na*/K*-ATPase
inhibitor ouabain (Dyer and Said, 1995; Said and Ma, 1994), which confirms the
Na^-independent behavior of this transport system. The apparent Na^, K* and iF
(pH)-independency of the riboflavin transport system would classify this system as a
uniporter, which challenges the current paradigm that mammalian apical solute carrier proteins are predominantly co-transporters (Hediger et a i, 1995). The outlined controversy surrounding riboflavin absorption prompted us to explore alternative hypotheses to better rationalize the uptake mechanism of this important vitamin. Another possibility for the active cellular uptake of riboflavin is internalization by surface receptors via a process similar to the receptor-mediated endocytosis of other water-soluble vitamins such as folate and vitamin Bi?. In fact, a soluble high-affinity riboflavin-binding protein has been detected in plasma (Zheng et a i, 1988) and the reproductive organs of either sex
(Natraj et al., 1994) although its function in riboflavin transport remains to be defined.
Interestingly, Low and co-workers recently showed the facilitated entry of bovine serum albumin (BSA) into lung epithelial cells and other cell cultures after covalently coupling
BSA to riboflavin (Holladay, etaL, 1999, Wangensteen, et at., 1996). Conjugates were detected in endosomal compartments, suggesting that BSA-riboflavin conjugates enter the cell via endocytosis. However, direct evidence on the involvement of endocytosis and/or transcytosis in the uptake of riboflavin in any cell type has not been reported previously.
To elucidate the cellular translocation mechanism of riboflavin in the intestine and investigate the potential involvement of a receptor-mediated endocytosis component, we have determined its binding and transport in the absence and presence of pharmacologically active compounds that are known to affect vesicular trafficking
36 pathways. We used the wed-characterized Caco-2 celt line as a model for the small intestine. These cells, when grown to confluence on polymer membrane inserts, mimic the in vivo intestinal absorption process enabling us to characterize vectorial substrate movement across both cell membranes and the cytoplasm.
37 METHODS
Materials. [^H]-Riboflavin (20 Ci/mmol) and [‘‘*C]-mannitol (60 mCi/mmol) were
purchased from Sigma (St. Louis, MO). [^H]-ChoIic acid (25 Ci/mmol) was from
American Radiolabeled Chemicals Inc. (St. Louis, MO). Cell culture materials and buffer
solutions were obtained from Gibco BRL (Grand island, NY). Transwell^ inserts were
purchased from Costar (Coming, NY) and rat tail collagen (type I) was from Becton
Dickinson Labware (Bedford, MA). Human transferrin, FUC-conjugation kit, BCA
protein assay kit, nocodazole and brefeldin A were purchased from Sigma (St. Louis, MO).
All other chemicals were from Fisher Scientific (Pittsburgh, PA).
Cell culture. Caco-2 cells were obtained from American Type Culture Collection
(ATCC, Rockville, MD). Cells with passage numbers 23-38 were maintained at 37 °C,
under 5% CO?, in complete medium consisting of Dulbecco’s modified Eagle’s medium
with 10% heat-inactivated fetal bovine serum, 1% non-essential amino acids, 100 U/ml
penicillin, 100 pg/ml streptomycin, lOmM HEPES. Cells were plated at 6x10“* cells/cm" in
tissue culture treated flasks and used during the exponential growth phase (day 5). Cells
were propagated and cultured as described previously (Hidalgo and Borchardt, 1990). Cell
monolayers were grown on collagen-coated polycarbonate Transwell™ membrane inserts
(3.0 pm pore size) at a density of 63,000 cells/cm". The culture medium was changed every other day during the first week after seeding and daily thereafter. Protein content was determined by the BCA method using bovine serum albumin as standard.
Assessment of transepitheliai membrane resistance. An epithelial voltohmmeter with dual electrodes (World Precision Instruments, New Haven, CT) was used to measure transepitheliai resistances of monolayers grown on filters as described by Rindler and 38 Traber (Rindler and Trabcr, 1988>. Filters were used only if the electrical resistance of the
monolayer exceeded 250 Q cm". Coincubation with (‘'‘C]-mannitol (0.2 pCi/ml) was
performed as an independent confirmation that paracellular transport in monolayers with
electrical resistances higher than 250 Q cm" was indeed minimal.
Transepitheliai transport studies (Caco-2 cells). Prior to transport studies, cell
monolayers were washed twice with warm Dulbecco’s-Phosphate Buffered Saline
(D-PBS) and incubated for 30 min at 37°C with substrate-free bathing medium (Hank’s
Balanced Salt Solution containing 25mM glucose, lOmM HEPES, adjusted to pH 7.4).
Transport studies were initiated by adding 1.5-480 pi of 10 pCi/ml [^H]-riboflavin and 30
pi 10 pCi/ml ['’*C]-mannitol stock solutions to the donor compartment of pre-equilibrated
Caco-2 cell monolayers, thereby achieving final donor concentrations of 0.5-160 nM
[^H]-riboflavin and 3.6 pM [‘‘*C]-mannitol. The donor compartment volume was adjusted
accordingly to maintain a constant 1.5 ml apical bathing volume. The transepitheliai flux of
[^H]-riboflavin and [‘'‘CJ-mannitol from either direction was followed over time by withdrawing samples at t=0, 15, 30,45,60,90,120, 150, and 180 min. To maintain a constant volume, an identical volume of bathing medium was added back after every sample. Cell monolayers (14 days post-seeding) that exerted a mannitol flux larger than
0.18%/hr/cm^ were excluded from data analysis (Hidalgo, 1996). At the end of experiments, the apical and basolateral bathing media were removed. Cell monolayers were washed three times with ice-cold PBS, pH 7.4, and cells were scraped off the inserts and processed for liquid scintillation counting.
Transport studies with Rat Intestinal Tissue. Three-month old male
Sprague-Dawley rats fed ad libitum were used in all experiments. The small intestine was 39 removed after decapitation and tissue was prepared for mounting in side-by-side diffusion
chambers (Ussing chamber type) as described previously (Swaan et al., 1994). Briefly,
mucosa was stripped of underlying muscle, mounted in Ussing chamber (1 cm^ exposed
surface area) and bathed on both sides with Ringer’s buffer solution. Solutions were
circulated by gas lift with carbogen (95%02-5%C02; Liquid Carbonic, Columbus, OH)
and maintained at 38°C (rat body temperature) by water-jacketed reservoirs. Tissues were
equilibrated for 30 min before initiation of experiments. Potential difference (Pj) and
short-circuit current (Isc) were monitored during the entire transport experiments to
ascertain tissue viability. Tissue integrity was independently determined by assessing the
['^Cj-mannitol flux (Marks et ai, 1991). Tissue viability was further assessed at the end of
the study by spiking apical sides of tissue with IM glucose solution (100 pi) and the
intestinal tissues with a 2 to 4-fold Isc jump were considered viable. Lc values ranged from
10-25 pEq/h»cm^ to 40-125 pEq/h«cm^ after addition of D-glucose.
Binding studies. Caco-2 cells were seeded on 6-well plates and cultured for at least
14 days. Prior to experiments, cells were washed twice with ice-cold PBS. After incubation at 4°C for Ihr with 2nM [^H]-riboflavin, cells were washed twice with ice-cold
PBS (6 ml per well for each 2 minute wash) and surface-bound riboflavin was released by incubation with 1 ml ice-cold PBS with pH values ranging from 3.0 to 8.0 for 2 minutes.
After buffer collection, cells were washed again with 6 ml ice-cold PBS and finally lysed with 0.5 ml 1% Triton X-100 solution. To ascertain mass balance cell lysates and samples from each PBS washing step were analyzed for radiolabeled material. InM ['*C]-mannitol was incorporated in the incubation medium as a control for the specificity of the washing steps. Nonspecific binding and potential passive diffusion was determined in parallel 40 studies by measuring radioactivity bound in the presence of a 1000-fold excess
non-radiolabeled riboflavin. Riboflavin-specific binding was obtained by subtracting
nonspecific binding count from the total radioactivity.
Analytical Methods. Radioactivity: The amount of dual-labeled radioactivity in
the samples was quantitated using a Beckman liquid scintillation counter (model LS
6000IC) at a counting efficiency of 43% and 75% for and ' ‘‘C respectively. HPLC
analysis: A Beckman HPLC system (Fullerton, CA), consisting of a Model 166 UV
detector, a LPSX gradient pump, was coupled in series with a Packard radiometric flow
scintillation analyzer (Packard Instruments, Meriden, CT). A Merck reversed phase
RP-C18 column (10cm, 5 pm) was eluted at a 3ml/min scintillation cocktail and a Iml/min
with 87% lOmM ammonium phosphate (pH S.5)/acetonitrile (Pietta et ai, 1982).
Data analysis and statistics. Uptake of riboflavin or cholic acid by cell
monolayers was expressed as fmoles/mg protein. Unidirectional flux was estimated over a
period of 0-180 min with an observed lag time of approximately 10-15 min. Linearity was
observed up to at least 180 min. Values in figures and tables are means ± S.D. of at least
three different experiments with cells from different passages. The effect of sample
withdrawal was taken into account for the calculation of fluxes using the following
equation:
G = % (^C n-,) + VCn (Equation 1) n=i
Where Q is the total amount of radioactive ligand in the donor compartment, V, is the sample volume, Vt is the volume of Transwell and Ci,2 n represents the concentration of sample 1,2 n. Transport parameters (Michaelis-Menten constant (Km), maximum
41 velocity (Jm ax).and passive permeability coefficient ( P m ) ) were calculated using the
NONLIN module in S YSTAT (version 8.0, SPSS Inc.) by non-linear regression analysis of
the obtained data to the general expression:
J = ^ + Pm*C (Equation2) Km + C
Statistical analyses were performed by one-way analysis of variance and significant differences were reported with a confidence interval of 95%.
42 RESULTS
Effect of culture time on riboflavin transport
Within 10 days after achieving confluency, Caco-2 cells undergo a differentiation
program that involves formation of tight junctions (as assessed by changes in electrical
resistance across monolayers grown on filters) and marked elevations in the levels of
several brush border hydrolases and transport proteins (Delie and Rubas, 1997). When
grown on filters, this proliferation and differentiation program yields a functional
monolayer of cells strongly resembling the small intestinal epithelium. It has been shown
previously that the transepitheliai transport level of some nutrients such as biotin and
vitamin 8,2 across these monolayers can vary significantly with days in culture depending
on their specific membrane transport protein expression during differentiation (Dix et al.,
1990; Ng and Borchardt, 1993). Thus, we first investigated the optimal culture time for
maximal expression of riboflavin transport protein(s). In this experiment, we chose a
[^H]-riboflavin concentration in accordance with the baseline riboflavin concentration in
human plasma (-12 nM) (Zempleni et al., 1996). [‘‘‘CJ-mannitol was used as a control to
ascertain monolayer integrity. This compound uniquely diffuses across the cell membrane
via the paracellular pathway, providing a quantitative measure of tight junctional
development in maturing Caco-2 cell monolayers (Marks et al., 1991, Swaan et al., 1994).
A relatively high mannitol flux 7 days after seeding indicates that a cohesive
monolayer has not yet been established (Fig. 2.1). This supports the premise that
membrane leakiness accounts for the apparent maximal riboflavin flux in the first week post-seeding. After 7 days in culture, however, no significant difference can be observed in the transepitheliai flux of riboflavin nor mannitol in both transport directions with regard to
43 culture time (apical (AP)-to-basolateral (BL) or BL-to-AP; p> 0.05; one-way ANOVA).
Accordingly, all subsequent transport studies were performed at or after 14 days in culture.
The BL-to-AP riboflavin flux is consistently higher, approximately 6-fold, compared to the
AP-to-BL flux, regardless of culture time.
Metabolic stability of [^H]>riboflavin
In addition to formation of tight junctions, functional differentiation in Caco-2 cells
is accompanied by elevated expression of several metabolic enzymes. Multiple studies
have shown that enzymatic activities gradually increase during differentiation and reach
maximal levels 15-21 days after confluency (reviewed by Delie and Rubas, 1997). It has
also been suggested that phosphorylation of riboflavin inside enterocytes is an essential
step of its apical absorption (Kasai et al., 1988). To investigate whether putative enzymatic conversion and metabolic instability of [^H]-riboflavin (which contains a general tritium
label) could contribute to the observed transport polarity, we analyzed samples from transport experiments on an HPLC system tethered to a radiometric liquid scintillation detector. Chromatograms of samples after a 3hr transport study (Fig. 2.2) indicated that most radiolabeled riboflavin on the donor side remains unchanged (Fig. 2.2B, 2.2C), although we cannot exclude the possibility that any metabolites are below the limit of detection on this system. On the apical acceptor side, 85% of transported ligand remains in the form of unchanged riboflavin (Fig. 2.2A), whereas the remaining activity cannot be ascribed to the major coenzyme forms of riboflavin, FMN and FAD, which would elute at
2.5 and 3.0 min respectively.
44 Polarization of riboflavin transport in rat tissue
Though the Caco-2 cell culture system is known to closely mimic small intestinal
enterocytes, it has been reported that these cells exert slightly different biochemical indices
and constitute altered protein composition and expression when compared to human small
intestinal enterocytes. To further validate that the observed transport polarity of riboflavin
did not reflect a cellular aberration inherent to the biochemical differences between Caco-2
cells and human intestinal epithelial cells in vivo, we conducted transport studies using rat
intestinal tissues mounted in side-by-side diffusion chambers (Ussing chamber type).
Figure 2.3 shows that riboflavin transport in rat intestine from both directions is linear up to
180 min with a significantly greater BL-to-AP flux (approximately 5-fold, 577.5
fmol/hr/cm^; R‘= 0.99) over AP-to-BL flux (124.9 fmol/hr/cm"; R‘= 0.98).
Concentration dependence of riboflavin transport
The observed polarization in transepitheliai transport of riboflavin could be
attributed to the presence of two different transport systems located on the AP and BL side
of the membrane. To test this possibility, concentration dependency experiments were
carried out in both transport directions. Figure 2.4 demonstrates that riboflavin is
transported by a binary mechanism consisting of both a saturable and a linear (passive) component. At concentrations near basal plasma riboflavin concentrations, riboflavin is transported predominantly by the saturable component. Table 2.1 lists the kinetic parameters of the transport systems on both sides of the epithelium. Consistent with our findings (previous sections) that riboflavin transport is greater in the BL-AP direction, we determined a basolateral transport system with a two fold higher affinity (Km) compared to
45 the apical translocation system. These results suggest that the higher BL-to-AP riboflavin
transport is partly due to the higher affinity of Rf for the BL system.
Effect of endocytosis inhibitors on riboflavin transport
The affinity values (Km) for carrier-mediated transport pathways are generally in
the pM range, whereas most receptor-mediated processes exert substrate affinity values in
the low nM range (Feener and King, 1998). Concentration dependency studies reveal two high-affinity riboflavin transport systems with Km values in the low nM range (Table 2.1).
Therefore, our results suggest the involvement of a receptor-mediated mechanism in the transepitheliai transport of riboflavin. Moreover, most receptor-mediated transcytosis systems transport their substrates preferentially in the BL-to-AP direction (Okamoto,
1998). The interesting consistency between these observations and our current results led us to hypothesize that cellular uptake of riboflavin may use a receptor-mediated endocytosis/transcytosis mechanism similar to that of other vitamins in the B group, e.g. folate and vitamin Biz Without the availability of a cDNA clone of the postulated transporter/receptor, it is relatively difficult to distinguish between RME and carrier-mediated pathways using contemporary biochemical or electrophysiological techniques since both mechanisms reveal characteristics of active transport processes.
However, several inhibitors are known to affect specific processes and/or organelles involved in vesicle trafficking, protein sorting, RME and transcytosis. To directly test our hypothesis, two inhibitors with different mechanisms of action were chosen: nocodazole and brefeldin A (BFA). At the cellular level, nocodazole inhibits endocytosis by depolymerizing microtubules (Hamm-Alvarez and Sheetz, 1998), which are necessary for
46 endocytic vesicles to move within the cell and BFA induces missorting of vesicles in the
rran^-Golgi network (Wan etal., 1992).
To confirm the specificity of endocytosis inhibitors in distinguishing
carrier-mediated processes from receptor-mediated endocytosis, we also examined the
transepitheliai transport of cholic acid (negative control) and transferrin (positive control).
Cholic acid is a known substrate of the intestinal bile acid transporter, a well characterized
carrier-mediated transport system (reviewed by Swaan, 1996). In Caco-2 cells, it has been
demonstrated that cholic acid is absorbed via a bile acid transporter with a Km of 49.7 pM
(Hidalgo and Borchardt, 1990). Furthermore, the transepitheliai flux of cholic acid can
serve as an indicator for functional integrity and viability of Caco-2 cell culture system
(Hidalgo, 1996). Transferrin (Tf), an iron transport protein internalized by the transferrin
receptor (TfR), is used as a positive control for endocytosis since the RME mechanism of
TfR has been well-established (reviewed by Huebers and Finch, 1987). Endocytosis of
FITC-Tf has been characterized in different cell lines and it is widely used as an endosome
marker in microscopic analysis of receptor-mediated endocytosis event (Teter K et al.
1998). Thus to visualize and detect the transport of Tf across the Caco-2 cell monolayer, a
FTTC-Tf conjugate was synthesized.
Figure 2.5 shows the effect of endocytosis inhibitors on bi-directional transepitheliai transport of riboflavin, cholic acid, and FTTC-Tf. Compared to the control, nocodazole inhibits BL-to-AP transport of riboflavin and FTTC-Tf (56.7% and 31.8% respectively), but does not affect cholic acid transport from both directions (Fig. 2.5A).
AP-to-BL transport of riboflavin and FTTC-Tf is significantly increased after treatment with nocodazole (Fig. 2.5B). Corresponding mannitol fluxes indicate that nocodazole does
47 not compromise the intactnes&of Caco-2 ceii monolayers during these transport studies
(control: 5.31 ± 1.33 pmol/hr/cm^, nocodazole treated: 4.42 ± 1.55 pmol/hr/cm^). Fig. 2.5B
demonstrates that after exposure of cell monolayers to BFA, AP-to-BL riboflavin flux
increases 37.1% but the transport of cholic acid is not affected. Polarized transport of
FTTC-Tf is observed in the control Caco-2 cells with a 6-fold greater BL-to-AP transport.
BFA significantly enhances FITC-Tf transport in both directions (AP-to-BL: 13 fold,
BL-to-AP: 5 fold). Elevated riboflavin fluxes are not the result of monolayer leakiness as
evidenced by constant mannitol fluxes (control: 5.31 ± 1.33 pmol/hr/cm", BFA treated:
6.19 ± 1.33 pmol/hr/cm^). Compared to control, the BL-to-AP transport of riboflavin is
slightly decreased although not statistically different (p> 0.05) while cholic acid transport
is not affected by BFA treatment (Fig. 2.5A). To investigate the effect of nocodazole on
apical uptake of riboflavin without the interference of BL-to-AP backflux, we performed
uptake studies using tissue culture plates. As shown in Fig. 2.5C, 58.4% apical uptake of
riboflavin is blocked by nocodazole within 20 minutes. The combined effects of
nocodazole and BFA on both the transport and uptake of riboflavin demonstrated the
involvement of a receptor-mediated endocytosis mechanism. This observation is further
substantiated by the effects of these compounds on FTTC-Tf and cholic acid. pH dependent dissociation of surface-bound riboflavin
An additional distinct mechanistic feature of ligands for receptor-mediated endocytosis pathways is pH-dependent dissociation of these molecules from their receptors
(reviewed by Mukheijee et a i, 1997). Consequently, we performed surface binding experiments at 4 °C using wash buffers with pH 3 to 8. At 4 °C the cellular internalization
48 of riboflavirv i& significanUy blocked (data not shown). Nonspecific binding i& determined
from parallel studies in the presence of 1000-fold excess unlabeled riboflavin, and
[‘‘*C]-mannitol binding serves as a negative control. A decrease in pH from 8 to 3 resulted
in elevated dissociation of riboflavin from its specific binding site(s) with significantly greater amounts of riboflavin released at pH 3 and 4, whereas the release of mannitol is pH-independent (Fig. 2.6). At physiological pH of the small intestinal lumen (pH 6-7), riboflavin shows higher binding interaction to its cell surface receptors.
49 DISCUSSION
Although multiple studies characterized the uptake mechanism of riboflavin in a
variety of tissues and species, there is surprisingly little information on the transepitheliai
transport of riboflavin across different cell types. In this study we find that, at
concentrations around its Km, riboflavin transport from the BL to the AP surface is 6-fold
greater compared to the flux in AP-to-BL direction. This elevated transport from the
systemic circulation into gut lumen has not been observed previously in either cell culture
systems (Fig. 2.1) or intestinal tissue preparations (Fig. 2.3). The fact that transport polarity
is conserved between two different species (rat vs. human) further confirms the
physiological and biological relevance of this observation. It also validates that the striking
polarity of riboflavin transport observed in Caco-2 cells is not due to a transport anomaly
specific to this particular cell line.
Interestingly, a similar transport polarity has been reported for intestinal transferrin absorption (Shah and Shen, 1994) with a 40-fold difference in BL: AP translocation activity and receptor expression. Enterocytes are known to obtain iron from dietary iron uptake and, during periods of low dietary iron intake, also from body iron stores.
Analogous to intestinal iron homeostasis, we anticipate that intracellular riboflavin levels in the gut are tightly regulated as well. Therefore, the putative physiological function of the riboflavin receptor on the basolateral surface of enterocytes is to participate in the supply of riboflavin to maturing epithelial cells. Since the intestine has the highest cell turnover rate of any tissue, continuous supply of riboflavin will be required to support cell growth and differentiation as well as the synthesis of essential flavoproteins such as digestive enzymes.
50 The Km we reported from transport experiments (Table 2.1 > are lOO-fold lower than
those reported by Said and co-workers, who found values in the low range (0.3 |xM)
via uptake studies (Said and Ma, 1994). It is essential, however, to point out two
consequential differences between the previous studies and our data. First, transepitheliai
transport in polarized cells comprises a series of sequential events, namely ligand-binding,
internalization, ligand movement through the cytoplasm, and translocation across two cell
membranes with different lipid compositions. Uptake experiments can only discern
binding and movement through the apical membrane. Second, we selected riboflavin concentrations according to the reported basal human serum level (12 nM), whereas earlier
studies cover higher concentrations in the pM domain. If more than one type of riboflavin transport system exists in Caco-2 cells, the higher affinity system would not be detected at
ligand concentrations ten-fold over the Km value of this system. Therefore, the possibility exists that small intestinal cells adopt two different membrane transport mechanisms for riboflavin with unequal substrate affinity. Interestingly, this observation correlates with the previous discovery of two unique co-existing transport systems for folic acid, another member of the B-vitamin group. It is now widely accepted that the cellular uptake of this essential molecule is mediated by both a GPI-anchored receptor with a Km of O.InM and a transmembrane carrier protein with an apparent Km of I^M (reviewed by Reddy and Low,
1998). These, and our current observation may suggest that more unique high affinity transport mechanisms could exist for other members of the B vitamin group.
In intestinal epithelial cells, microtubules extend in straight arrays from the apical to the basolateral cell membrane providing an essential network for many membrane-trafficking events including endocytosis and transcytosis (Hamm-AIvarez and 51 Sheetz, 1998). Disruption of microtubules with nocodazole significantly inhibits
BL-to-AP transport of riboflavin (Fig. 2.5A). These results are in good agreement with the data by Maple et al. (1997), who reported a 50-75% inhibition of the BL-to-AP transcytosis of pIgA in MDCK cells in the presence of nocodazole. More importantly, nocodazole inhibition of the BL-to-AP flux of FTIC-labeled transferrin, a well-characterized RME substrate, suggests that these two compounds share similar microtubule-dependent intracellular trafficking pathways. Specificity of nocodazole for
RME processes, but not carrier-mediated events, was shown by the apparent lack of reduction in cholic acid flux. This observation, in turn, rules out the possibility that our data are a reflection of cellular damage caused by the adverse effects of nocodazole on protein synthesis and signal transduction (Hamm-Alvarez and Sheetz, 1998).
It has been reported that polarized epithelial cells have spatially separated early endosomal systems governing distinct bidirectional endocytosis/transcytosis processes
(Mukheijee et ai, 1997). These two pathways are also functionally distinct in their sensitivities towards pharmacological agents (Okamoto, 1998). Interestingly, nocodazole treatment results in increased AP-to-BL transport of both riboflavin and FTTC-Tf (Fig.
2.5B). Currently, the factors underlying this phenomenon are not completely understood and needs further investigation.
Even though AP and BL endosomal systems are separately regulated in epithelial cells, studies in Caco-2 cells have revealed an interconnection via intermediate endosomes and common recycling compartments (Knight et ai, 1995). BFA is known to cause intracellular missorting of ligand-receptor complex within the endosome-rrans Golgi network, a region closely involved in vesicular sorting and bidirectional
52 endocytosis/transcytosis (Wan é ta t, 1992). hr oorstudies, BFA results in a significant
increase in AP-to-BL transport of riboflavin and FTTC-Tf without perturbing the cholic
acid flux. BL-to-AP riboflavin transport is slightly, but not significantly reduced (p> 0.05).
These results are consistent with previous studies showing that BFA induces TfR-mediated
transcytosis of ['‘^I]-Tf across Caco-2 cell monolayers in both transport directions (Shah
and Shen, 1994). These observations are also in agreement with the findings that BFA
treatment leads to enhanced transcytosis of ricin and Botulinum neurotoxin
(Maksymowych and Simpson, 1998; Prydz et a t, 1992). Shen and colleagues
demonstrated that enhancement of Tf transcytosis by BFA is accompanied by a significant
increase in the number of TfR on the AP cell membrane, and a decrease in the number of
TfR on the BL cell membrane without affecting total TfR expression (Shah and Shen,
1994; Wan et a t, 1992). Whether a similar mechanism is responsible for increased
riboflavin transport remains to be investigated.
Our finding that dissociation of riboflavin from its specific surface binding site(s) is
pH dependent (Fig. 2.6) provides additional biochemical evidence corroborating our
hypothesis. This observation is consistent with a hallmark feature of receptor-mediated
endocytosis in which acidification of endosomes during the endocytic sorting process
causes dissociation of ligands from receptors. pH sensitive binding has been reported
previously for folate and vitamin B,2, both well-characterized ligands utilizing RME
mechanisms (Kamen and Capdevila, 1986).
In summary, our experiments demonstrate that a specialized, high-affinity riboflavin translocating system(s) exists in the intestinal cells with affinity values and biochemical characteristics similar to most receptor-mediated systems reported in the
53 literature. Thtssystetitisprimariljr expressed o i t the basolateral surface. Although the
physiological role behind this type of polarity needs to be further investigated, our findings
suggest that the basolateral uptake system could play an important role in supplying
nutrients to the highly proliferative intestinal epithelium. These data aid in our
understanding of the transepitheliai transport mechanism of riboflavin and the proteins
involved in its uptake and contribute to our knowledge of human riboflavin homeostasis
and the physiology of B vitamins in general.
ACKNOWLEDGMENTS
The authors would like to thank Dr. Michael Darby at the OSU comprehensive cancer center for his kind assistance with the use of the radiometric HPLC system.
54 Km(nM) Jmax (fmol/hr/cm^) BLto AP 4.06 ± 0.03 168.2 ±0.38
AP to BL 9.72 ± 0.85 192.98 ±0.85
Table 2.1 Transport parameters of bi-directional transport of riboflavin in Caco-2 cell monolayers.Transport parameters were calculated by nonlinear regression analysis of data obtained from concentration dependence study to equation 2. Parameters are presented as estimates ± asymptotic standard error.
55 240 40
220 Rf (AP-to-BL) Rf (BL-to-AP) 35 200 fWla(AP-to-BL) Ma(BL-to-AP) *£ 180 3 0 Î | . e o I J 140 X I I 120 “I (0 % 100 20 I 80 15 U • t - 60
40 10 20 0 I m « 7th 14th 21th 28th
Fig. 2.1. Effect of culture time on riboflavin transport in Caco-2 ceil monolayers.On 7,14,21,28 days post seeding, transpoit studies were initiated by spiking apical or basolateral sides of Caco-2 cell monolayers with [^H]-Rf and [‘‘*C]-mannitol (final concentration 5 nM and 3.6 pM respectively). Flux values were calculated according to methods described in the experimental procedures. Each bar/point represents the mean ± s.d. of triplicate cell monolayers.
56 310 dpm
210
140
S. 10 70
0 0
10616
170
3603
9623
Fig. 2.2. Stability of [^H] riboflavin in 14 day post-seeding Caco-2 cells.Apical (AP) and basolateral (BL) samples after 3hr transport study were analyzed by HPLC with radiometric detection. [^H]-riboflavin elutes at 6.0-6.2 min and [‘‘‘C]-mannitol has a retention time of 1.6-1.7 min. A, AP sample chromatogram from 3 hr BL-to-AP transport study. B, BL sample chromatogram from 3hr BL-to-AP transport study. A yet to-be-defined peak was detected in apical samples at 5.1 min. C, AP sample chromatogram from 3 hr AP-to-BL study.
57 2500
• Rf (AP-to-BL) o Rf (BL-to-AP)
2000 -
1500 - c
11000 -
500 -
0 30 60 90 120 150 180 Time (min)
Fig. 2.3. Time course of riboflavin transport in rat ileum.Rat ileal mucosa was mounted in diffusion chambers, circulated with 95% 02-5%C02 with Ringer solution containing 0.1 pM non-radioactive riboflavin on both sides. Transport studies were initiated by spiking the mucosal (#) or serosal (O) bathing solution with [^H]-Rf and [*‘*C]-mannitol (final concentration 5 nM and 3.6 pM respectively).
58 3000
2500
^ 2000
£ 1500
i 1000
500
0 20 40 60 80 100 120 140 Rf concentration (nM)
750
600 %
450
? 300 E.
150
0 20 40 60 SO 100 120 140 180 Rf concentration (nM)
Fig. 2.4. Concentration dependence of transport of riboflavin (Rf) in Caco-2 cell monolayers. Radiolabeled RF (0.5-160 nM) and mannitol (3.6 pM) were added to the basolateral (4A) or apical (4B) side of Caco-2 cell monolayers on the 14“' day post-seeding. Each point represents the mean ± s.d. of triplicate cell monolayers. The solid line represents the calculated fit of equation 2 as described in the experimental procedure. The estimated active transport component (Jacuve) to the total Rf flux (Jtotai) is indicated with a dotted line; the estimated passive component (Jpa^ive) is indicated with a dashed line.
59 1600 NCDZ NCDZ "S 500 -
S wo
i CA FITC-Tf
2 100
S 80
Fig. 2.5. EfTect of nocodazole and brefeldin A on transport and uptake of riboflavin, cholic acid, and FITC transferrin in Caco-2 cell monolayers. BL-to-AP transport (5A), AP-to-BL transport (5B) oFRf, cholic acid (CA), andFTTC-Tf in nocodazole (3.3 pM)- or brefeldin A (1.65 pM)-treated Caco-2 cells. 5nM [^H]-Rf or 60pg/ml FITC-Tf was added to the basolateral or apical side of cell monolayers. 8nM [^H]-cholic acid was applied to basolateral or apical side of cell monolayers with 50 pM non-radioactive CA on both sides. C, Uptake of Rf, CA, mannitol in nocodazole-treated Caco-2 cells. InM [^H]-Rf or [^H]-CA and [*‘*C]-mannitol were added to nocodazole pre-treated (3.3 pM, 15 min) Caco-2 cells seeded in 12-well plates. Cells were washed three times with ice-cold PBS, lysed with 0.1 % Triton XlOO and measured for radioactivity after 20 min uptake study. Each bar represents mean ± s.d. of triplicate cell monolayers (** p< 0.01 ; * p< 0.05).
60 Riboflavin M annitol - 45 - 40
- 35 a. oi • 30 % ■ 25
- 20 I 0.5 -
0.0 pH 8 pH 7 pH 6 pH 5 pH 4 pH 3
Fig. 2.6. pH dependence of riboflavin-specific binding to cell surface.Cells were incubated with 2nM [^H]-Rf and InM ['^C]-mannitol at 4 °C for Ihr. After removal of free [^H]-Rf with two ice-cold PBS washes, cells were incubated for 2 min at 4 °C with 1ml PBS at various pH. Total ligand-binding activity was determined by measuring radioactivity in PBS release buffers. The nonspecific binding component was determined in parallel studies in which cells were incubated with 1000-fold excess non-radioactive Rf for Ihr. A specific binding component was calculated by subtracting the nonspecific component from total binding values. Each point (# , Rf; A, mannitol) represents mean ± s.d. of three experiments.
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64 CHAPTER 3
RIBOFLAVIN UPTAKE IN HUMAN TROPHOBLAST-DERIVED BEWO
CELL MONOLAYERS: CELLULAR TRANSLOCATION AND
REGULATORY MECHANISMS
ABSTRACT
Riboflavin (vitamin 83) is essential for fetal development and must be acquired from maternal sources. The uptake mechanism of riboflavin and the major regulatory pathways involved were characterized in a model for the placental barrier, the human choriocarcinoma cell line, BeWo. Uptake of [^H]-riboflavin was saturable {Km = 1.32 ±
0.68 nM, J„tax = 266.63 ± 26.89 fmol/mg protein/20 min) and was significantly reduced at low temperature and in the presence of metabolic inhibitors (azide, 2-deoxyglucose) or structural analogs. Ouabain, amiloride, sodium-free buffers, and medium with pH values ranging from 3 to 8 did not affect uptake of [^H]-riboflavin. In contrast, substitution of Cl with other monovalent anions significantly inhibited its uptake. Induced differentiation of
65 BeWo cells into syncytiotrophoblasts by forskolin or 8-Br-cAMP introduced a time- dependent decrease of riboflavin uptake. Preincubation of cells with activators of cyclic nucleotide dependent protein kinase pathways (3-isobutyl-1-methylxanthine and pCPT-cGMP) and calmodulin antagonists (calmidazolium and W-13) resulted in a concentration-dependent reduction of [^H]-riboflavin uptake, whereas specific modulators of protein kinase C pathways did not have significant effects.
3-isobutyl-1-methylxanthine exerted its regulatory effect on riboflavin uptake via decreasing both AT„, and Jmax of the riboflavin uptake process {Km = 6.32 ± 1.29 nM, Jmax =
135.57 ± 10.42 fmol/mg protein/20 min). In summary, we report the presence of high affinity riboflavin transporter(s) on the microvillous membrane of BeWo cells, that appears to be modulated by cellular cyclic nucleotides levels and calmodulin.
66 INTRODUCTION
During pregnancy, the placenta not only provides a barrier separating the maternal and fetal compartments but also serves as a transport organ supplying nutrition from the mother to the developing fetus. In humans, the maternal and fetal circulations are physically divided by a placental barrier containing the trophoblasts, villous stroma, and the fetal capillary endothelium (Rama Sastry, 1999). Transport of hydrophilic nutrients and drug molecules across the placenta is mainly controlled by the two membranes of the polarized trophoblasts, with the apical microvillous membrane in direct contact with the maternal circulation and the basal membrane facing the fetal side (van der Aa et al.,
1998). To facilitate efficient passage of crucial nutrients to the fetus, trophoblasts are known to express specific nutrient transporter systems on their apical cell surfaces (Knipp et al., 1999).
Riboflavin, also known as vitamin B?, is essential for normal cellular function.
Maternal intake of riboflavin has been shown to associate positively with fetal growth and development (Badart-Smook et al., 1997). Animal studies suggest that riboflavin deficiency during pregnancy leads to congenital abnormalities (Rivlin, 1975). Despite its critical importance to the developing fetus, the molecular mechanism and regulation of
67 riboflavin translocation across the trophoblast is still poorly understood. Studies with
perfused human placenta tissues and in vivo analysis of maternal and umbilical cord
plasma riboflavin levels identified a saturable uptake process on the maternal surface of
the placenta putatively responsible for a four-fold elevation in free riboflavin concentrations of fetal plasma (Dancis et al., 1985; Danois et al., 1988; Zempleni et al.,
1992; Zempleni et al., 1995). Moe and colleagues showed that riboflavin is taken up into human syncytiotrophoblasts-derived membrane vesicles by a high-affinity membrane component via a concentration- dependent, Na^-independent mechanism (Moe et al.,
1994). However, the overall internalization process was found to be insensitive to temperature, a result inconsistent with general criteria for active transport processes and contradictory to previous mechanistic studies on riboflavin uptake reported in other tissues (Said and Ma, 1994; Kumar et al., 1998; Said et al., 1998; Huang and Swaan,
2000).
The objective of this study is to disseminate the uptake mechanism of riboflavin in the human placenta and to document its intracellular regulatory pathway(s). We used a human choriocarcinoma- derived cell line, BeWo, as a model for trophoblasts. Compared to isolated membrane preparations, an intact cell line system provides mechanistic model
68 to integrate cellular uptake processes and intracellular events associated with modulation of transporter function. Under normal conditions, BeWo cells have been shown to exhibit morphological and biochemical features that strongly resemble proliferative cytotrophoblasts; furthermore, these cells can be induced into differentiated syncytiotrophoblasts in vitro using pharmacological agents (Wice et al., 1990; Liu et al.,
1997). More importantly, several studies have shown that the characteristics of membrane transport systems expressed in BeWo cells are highly similar to those reported in normal human trophoblasts (Cool et al., 1991; Prasad et al., 1997).
69 METHODS
Materials. [^H]-Riboflavin (20 Ci/mmol) and l’ C]-mannitoI (60 mCi/tnmol) were
purchased from Sigma (St. Louis, MO) and Moravek Biochemicals (Brea, CA),
respectively. Cell culture materials and buffer solutions were obtained from Gibco BRL
(Gaithersburg, MD) and Lab Tek Chamberslide System was from Nalge Nunc
(Naperville, IL). Rat tail collagen (type I) was from Becton Dickinson Labware (Bedford,
MA) and BCA protein assay kit was purchased from Sigma (St. Louis, MO). All other
chemicals were from Fisher Scientific (Pittsburgh, FA) and Sigma (St. Louis, MO).
Cell culture. BeWo cells were obtained from American Type Culture Collection
(Manassas, VA). Cells with passage numbers 191-230 were maintained at 37 °C, under
5% CO2, in complete medium consisting of F-12K medium with 10% fetal bovine serum,
1% non-essential amino acids, 100 U/ml penicillin, 100 pg/ml streptomycin. Cells were
routinely maintained in tissue culture treated 175 cm" flasks and the culture medium was replaced every other day. The cells were harvested at 80% confluency (day 4-5) by exposure to a trypsin-EDTA solution (0.25% trypsin and 0.002% EDTA in HBSS). Cell monolayers were grown on rat tail collagen-coated 12- or 24 well- plates (3.8 and 2.0 cm^, respectively) at a density of 5x10“* cells/cm". Confluent monolayers were formed between
70 3-5 days after seeding and were used for experiments at that time.
Uptake Experiments. Confluent BeWo cells were washed twice with warm (37°C)
Dulbecco’s PBS (pH 7.4) before studies were initiated. Riboflavin uptake studies were
performed at 37°C in bathing medium (Hank’s balanced salt solution containing 25 mM
glucose and 10 mM HEPES, adjusted to pH 7.4) with a final concentration of 5 nM
[^H]-riboflavin. ['‘*C]-mannitol (0.37 pM) was incorporated in the incubation medium as
to determine the specificity of the washing steps. Mannitol has been shown to diffuse
across the cell membrane solely by passive diffusion and, thus, can serve as a control to
determine the respective effect of physical or pharmacological cell perturbations on
passive and active transport process (Huang and Swaan, 2000). After 20 minutes, bathing
medium was aspirated, and cells were washed twice with ice-cold Dulbecco’s PBS, pH
3.0 (2 ml per well for each 2-min wash) to remove free and surface-bound riboflavin
(Huang and Swaan, 2000). Finally, cells were lysed with 1% Triton X-iOO solution and
the amount of dual-labeled radioactivity in the cell lysates was quantitated using a
Beckman liquid scintillation counter (model LS 60(H)IC). Cellular protein content was determined by the BCA method using bovine serum albumin as a standard. Modulators of signal transduction pathways were prepared in either DMSO or absolute ethanol (fînal
71 concentrations of the organic solvent vehicle (DMSO or EtOH) was incorporated in the bathing media of the control experiments to determine the effect of these solvents on untreated cells. Viability of cells under all treatment regimens was monitored by the trypan blue exclusion method and was routinely between 92 and 96%. In vitro dlfTerentiation experiments.Forskolin or 8-bromo-cAMP was added to complete cell culture medium at a final concentration of 100 pM and 250 pM, respectively, followed by filter sterilization (0.22 pm). BeWo cells were grown in drug-free culture medium for two days to allow cell proliferation. Regular medium was replaced with drug-containing medium on the 3'^'' day and the medium was changed daily to ensure sufficient supply of nutrients. Parallel sets of control and differentiated BeWo cells grown on the ChamberSlide were processed with Giemsa stain (3%) for 2h at room temperature to visualize nuclei under a phase contrast light microscope. Data analysis and statistics. Kinetic uptake parameters such as the concentration at half-maximal transport velocity (Kr, Michaelis-Menten-type constant), maximum uptake velocity (Jmax), and passive permeability coefficient (Pm) were calculated using the NONLIN module in SYSTAT (version 8.0, SPSS Inc.) by non-linear regression analysis 72 of the obtained data to the general expression: J ^ Jj^ * C ^ ^ (Equation 1) Km + C where 7, is the total flux and C is the riboflavin concentration. All results are expressed as means ± standard deviation (SD). Statistical analyses between two groups were performed using Student’s r-test and one-way analysis of variance (ANOVA) was used for single and multiple comparisons. Significant differences were reported with a confidence interval of 95%. 73 RESULTS Riboflavin uptake kinetics The presence of a riboflavin transport system in the BeWo cell line was first determined by assessing the kinetics of riboflavin uptake. A [^H]-riboflavin concentration in accordance with the free riboflavin concentration in human maternal blood (~ 5 nM) (Zempleni et al., 1995) was used to examine the time course of riboflavin uptake. As shown in figure 3.1, uptake increased linearly up to 20 min ( r = 0.99, rate = 10.91 fmol/min/mg protein) and approached equilibrium at 40 min. Based on these experiments all subsequent uptake studies and kinetic analyses were performed from data collected through 20 min. Fig. 3.2 shows the relationship between the uptake of [^H]-riboflavin at 20 min and its concentration (1.0 nM to 1.0 ^M) in the bathing solution. Analysis of total [^H]-riboflavin uptake data reveals an absorption mechanism consisting of two pathways: a saturable pathway at low concentrations and an apparently non saturable (passive) pathway that dominates at concentrations above 250 nM (Fig.2). [*‘*C]-mannitol uptake ranged from 1.20 to 1.61 pmol/mg protein at all studied [^H]-riboflavin concentrations. After fitting the riboflavin data to equation 1, an uptake system with an apparent K, of 74 1.32 ± 0.68 nM and a Jmax of 266.63 ± 26.89 fmol/mg protein/20 min was identified. The dotted line represents the uptake for the saturable component calculated from the kinetic parameters. The dashed line represents the uptake for the non-saturable flux generated from multiplying the riboflavin concentration with the passive diffusion coefficient (Pm, 1.19 ± 0.05 fmol/mg protein/20min). Based on these data, subsequent experiments aimed to characterize the saturable riboflavin uptake component were conducted at 5 nM [^H]-riboflavin; at this concentration, the non saturable transport component contributes merely 3.81 ± 0.66 % to the total riboflavin uptake process (Fig. 2, inset). Substrate specificity of BeWo cell riboflavin uptake To investigate the substrate specificity of the saturable uptake process, we established the effects of various riboflavin coenzymes and structural analogs on [^H]-riboflavin uptake in BeWo cells (Fig. 3.3). 85.6 % of [^H]-riboflavin uptake was blocked in the presence of lOOO-fold unlabeled riboflavin. The major coenzyme forms of riboflavin, FMN (flavin mononucleotide) and FAD (flavin adenine mononucleotide), significantly inhibited uptake of [^H]-riboflavin (73.5 and 49.5% respectively). It should be noted that FAD has significant molecular bulk attached to the 5’-hydroxy moiety on the ribose chain, namely a trihydrogendiphosphate-adenosine group (M*= 409) which 75 doubles the molecular weight of this molecule compared to riboflavin. Interestingly, lumiflavin, a riboflavin analog with a methyl group substituted for the ribose side chain, was equally effective in inhibiting [^H]-riboflavin uptake (48%) compared to FAD; on the other hand, lumichrome, an analog that completely lacks the D-ribose side chain showed limited affinity for the riboflavin transport system (28.4% inhibition). Addition of D-ribose to the incubation medium did not significantly affect [^H]-riboflavin uptake. The molecular structures of folates and flavins reveal the presence of a pterin (pyrazino[2,3-d]pyrimidine) ring system and several tricyclic antidepressants such as chlorpromazine and amitriptyline are structurally related to riboflavin. Co-administration of these agents has been shown to enhance urinary excretion of riboflavin and accelerate tissue depletion of FAD levels in the liver (Pinto and Rivlin, 1987). However, at the cellular level, neither folate, nor chlorpromazine, nor amitriptyline had a significant effect on [^H]-riboflavin uptake in BeWo cells. Temperature, energy dependence The riboflavin uptake pathway in BeWo dels was further characterized using two additional criteria for active membrane transport, namely temperature and energy dependence. To determine temperature dependence of the transport pathway, uptake of 76 [^H]-riboflavin was performed at decreased temperatures. The rates of uptake were 3.217, 1.591, and 1.213 fmol/min at 37,20, and 4 °C, respectively, indicating that riboflavin uptake is temperature-dependent. At these conditions, [14C]-mannitol uptake was 0.014 (± 0.002), 0.010 (± 0.001), and 0.010 (±0.001) pmol/min at 37, 20, and 4 °C, respectively). Riboflavin uptake was also significantly reduced in BeWo cells pretreated with metabolic inhibitors (50 mM sodium azide/10 mM 2-deoxyglucose: 114.98 ± 11.8 fmol/mg protein, control: 184.25 ± 4.25 fmol/mg protein; 15 min preincubation). ['‘*C]-mannitol uptake was not significantly affected (p>0.05) in cells exposed to metabolic inhibitors (treated: 0.734 ± 0.10 pmol/mg protein; control: 0.916 ± 0.08 pmol/mg protein). These results imply that the riboflavin transport process is dependent on metabolic energy. Taken together, these results provide additional support for the presence of a carrier system that specifically mediates the uptake of riboflavin into BeWo cells. Ion-coupling properties of riboflavin uptake Active transport processes are generally energized by co-transport of ions (solute transport family) or ATP hydrolysis directly coupled to the transport system (ATP binding 77 cassette [ABC]-transport family). In mammals, transport of organic solutes is primarily coupled to the electrochemical gradients of Na^ or (Hediger et al., 1995). To investigate the role of sodium in riboflavin uptake in BeWo cells we replaced Na^ ions in the bathing media with choline, or Li* (Table 3.1). None of the aforementioined substitutions led to significant inhibition of riboflavin uptake. Preincubation of BeWo cells with 1 mM ouabain, a specific Na*-K*-ATPase inhibitor, for ih did not affect riboflavin uptake corroborating the previous results with Na*-free and K*-enriched media. To test the potential involvement of a hydrogen-coupled transport pathway, bathing solutions with H*-concentrations ranging from 10 ^ to 10 ® M were prepared by adjusting the pH of incubation media. Table I shows that uptake of riboflavin is not affected by solution pH over the hydrogen concentration range tested, suggesting that the uptake process is not driven by an inwardly directed proton gradient (Table 3.1). Pretreatment of cells with various concentrations of amiloride, a specific Na*/H* exchanger inhibitor, failed to block uptake of riboflavin (Table 3.2), which was taken as an additional validation for both the Na* and H* dependency experiments. Several studies have demonstrated that internalization of dopamine and other neurotransmitters is markedly affected by Ca‘* concentration (Uchida et al., 1998). To 78 assess the calcium dependency of riboflavin uptake, studies were performed in Ca^^-free buffer or incubation medium with 1 mM EGTA. At cellular level, neither treatment significantly inhibited riboflavin uptake. Chloride is the major physiological anion with a twenty- fold higher concentration in the extracellular fluid (103 meq/1) relative to the intracellular environment (4 meq/1) (Guyton and Hall, 20(X)). Studies have shown that active transport of neurotransmitters including serotonin, GAB A and some P-amino acids requires specific coupling of a downhill Cl gradient into the cells (Griffith and Sansom, 1998). To test the hypothesis that riboflavin uptake depends on selective co-transport of C l, experiments were carried out in bathing media containing salts of alternative organic and inorganic monovalent anions (gluconate, iodide and isocyanate). Substitution of Cl ions with I , SCN or gluconate significantly reduced the uptake of riboflavin in BeWo cells (21.5,33.2, and 15.6% respectively; Table 3.1). Analysis of concomitant [''*C]-mannitol uptake reveals that substitution of Cl ions does not significantly affect the passive diffusion of this marker molecule, indicating the specificity of this treatment regimen on the riboflavin transport system. Effects of anion-exchange inhibitors and general organic anion transporter inhibitors on riboflavin uptake in BeWo cells were also investigated (Table 3.2). Pre-treatment of BeWo cells 79 with various concentrations of probenecid (a general organic anion inhibitor) or furosemide (a Na^-K'^-2Cr cotransporter inhibitor) did not decrease riboflavin uptake, whereas 4 ,4’-diisothiocyanostilbene-2,2’-disulfonic acid (DIDS), a membrane impermeable anion-exchanger inhibitor, constantly reduced riboflavin uptake 20 to 30%. Control [’'*C]-mannitol data indicate DIDS inhibition to be specific for the riboflavin uptake system. Riboflavin uptake in syncytiotrophoblasts Syncytiotrophoblasts, the differentiated phenotype of cytotrophoblasts, are the main functional units of the term placenta (Knipp et al., 1999). More importantly, several studies have demonstrated that induction of cell differentiation was frequently accompanied with changes in membrane transporter activities (Furesz et al., 1993; Ogura et al., 2000). The majority of BeWo cells in culture express the cytotrophoblasts phenotype; however, they can undergo in vitro differentiation into syncytiotrophoblasts upon stimulation with cAMP modulators (Wice et al., 1990; Liu et al., 1997). To examine the uptake activities of riboflavin in the syncytiotrophoblast, two cAMP modulators with different mechanisms of action were used to induce differentiation of BeWo cells. Forskolin activates adenylyl cyclase and treatment with this drug has been 80 shown to increase linoleic acid transport in BeWo cells (Liu et al., 1997). 8-bromo-cyclic AMP (Br-cAMP), a membrane permeable cAMP analog, has been demonstrated to stimulate the expression of the placental facilitative glucose transporter-1 (GLUT 1) (Ogura et al., 2(HK)). Incubation of BeWo cells with 1(X) pM forskolin or 250 jiM Br-cAMP for 24 h resulted in significant decrease in [^H]-riboflavin uptake (Fig. 3.4) although noticeable features of morphological differentiation were not found under light microscope (results not shown). Continuous drug exposure induced a time-dependent reduction in [^HJ-riboflavin uptake. After 72h treatment, both agents induced 21% reduction in riboflavin uptake that was accompanied with dramatic morphological changes. Consistent with previous reports (Wice et al., 1990; Liu et al., 1997), differentiated BeWo cells exhibit an increased number of enlarged cells with fused nuclei and large intercellular openings closely resembling syncytiotrophoblasts (Fig.3.5). Parallel sets of cells were further used in these studies to assess whether the specificity of riboflavin uptake was modified during in vitro differentiation. Competitive studies were carried out by measurement of [^H]-riboflavin uptake in the presence and absence of l(XX)-fold of unlabeled riboflavin. Compared with the control cytotrophoblasts, no significant difference was found in the differentiated BeWo cells during various time 81 periods (control: 26.83 ± 4.72% and drug-treated cells: 26.92 ± 5.47%, p = 0.974). Effect of cyclic nucleotide-dependent pathways on riboflavin uptake The reduction of riboflavin uptake by cAMP modulators suggests a role of cAMP- dependent protein kinase A pathways (PKA) in the regulation of membrane riboflavin transport. To directly test this hypothesis, riboflavin uptake studies were conducted in the presence of 3-isobutyl-1-methyl-xanthine (BMX), a PKA pathway activator that prevents degradation of camp by inhibiting cyclic nucleotide phosphodiesterase (Shafer et al., 1998). Short-term IB MX incubation (ih) resulted in immediate and significant inhibition of [^H]-riboflavin uptake in BeWo cell monolayers (Fig. 3.6A). To further investigate the mechanism involved in the IBMX-mediated reduction, the riboflavin transporter activity in control and IBMX-treated BeWo cells was analyzed (Fig. 3.6B). Kinetic analysis revealed that IBMX-induced inhibition of riboflavin uptake is accompanied by changes in K, as well as in Jmax- IB MX treatment increased the K, from 1.10 ± 0.05 nM to 6.32 ± 1.29 nM. Furthermore, the J,mx was markedly reduced from 281.80 ± 2.99 fmol/mg protein/20 min to 135.57 ± 10.42 fmol/mg protein/20 min in IBMX-treated cells. 82 Since IB MX exhibits broad-spectrum activity towards several subtypes of cyclic nucleotide phosphodiesterases (PDEs) (Schudt et al., 1996), IBMX-mediated reduction of riboflavin uptake may originate from its effect on modulation of the cGMP-dependent protein kinase (PKG) pathways. To examine the involvement of PKG pathway, we carried out uptake studies in the presence of pCPT-cGMP, a membrane permeable analog of cGMP. Pretreatment with pCPT-cGMP caused significant attenuation of riboflavin uptake in BeWo cell monolayers (Table 3.3). Co-incubation of pCPT-cGMP and IB MX potentiated the uptake reduction to 43.7% (p <0.001), 15.3% (p <0.05) compared with that of cells treated with pCPT-cGMP or IB MX alone, respectively. Analysis of concomitant ['‘*C]-mannitol uptake reveals that treatment of BeWo cells with the aforementioned pharmacological agents does not significantly affect the passive diffusion of this marker molecule, implying specificity of this treatment regimen on the riboflavin transport system. Involvement of protein kinaseC- and calmodulin* mediated pathways The apparent PKA- and PKG-dependent uptake of riboflavin suggested a potential involvement of multiple signal transduction pathways in the regulation of membrane riboflavin transporter activity. Therefore, we further assessed the influence of protein 83 kinase C- (PKC) and calmodulin-mediated pathways on riboflavin uptake in BeWb cell monolayers. No significant change was found in riboflavin uptake upon treatment of BeWo cells with phorbol-12- myristate-13-acetate (PMA), a well-known protein kinase C activator or chelerythrine, a selective protein kinase C inhibitor (Table 3.4). Mixed effects were observed in Be Wo cells exposed to modulators of calmodulin (CaM)-mediated pathways. Pretreatment of cells with calmidazolium and W-13, two specific but structurally different CaM antagonists, resulted in marked concentration- dependent reduction on riboflavin uptake (Table 3.4). In contrast, neither KN-93, a selective Ca^*/CaM-dependent protein kinase II inhibitor, nor KN-92, negative control of KN-93, exhibited significant effect. Further kinetic analysis of riboflavin uptake in the presence and absence of calmidazolium was performed to elucidate the underlying mechanism of drug-mediated inhibition. A reduction in both K, and Jmax was observed (Fig 3.7). The maximal uptake velocity dropped 59.8% (control: 295.42 ± 0.03 fmol/mg protein/20 min; calmidazolium-treated cells: 118.80 ± 5.31 fmol/mg protein/20min), whereas the riboflavin affinity to its transporters changed from 1.35 ± 0.01 nM to 5.31 ± 0.01 nM. 84 DISCUSSION The placental membrane mediates the entry of the maternal nutrients into the fetal circulation. Although transplacental transport of riboflavin has been extensively studied in the perfused placental tissues, little is known regarding its cellular translocation and regulation mechanism across the cytotrophoblasts. The current study reports the existence of high affinity riboflavin transporter(s) on the microvillous membrane of the BeWo human placental choriocarcinoma cell line. Supporting evidence for this active transport mechanism include i) a significant dependency of riboflavin uptake with temperature, ii) reduction of uptake in the presence of metabolic inhibitors, iii) a saturable uptake component, and iv) inhibition of riboflavin uptake in the presence of structural analogs. Riboflavin has relatively high affinity for its uptake pathway (K, = 1.32 nM) suggesting the presence of a receptor-based translocation mechanism, which may resemble the receptor-mediated riboflavin pathway in Caco-2 cells we recently documented (Huang and Swaan, 2000). The essential structural requirements for specific interaction between riboflavin and its placental transporters were deduced from inhibition studies with structural analogs. The isoalloxazine ring appears to have an essential role in ligand-transporter interactions, 85 as evidence by the finding that D-ribose does not affect riboflavin uptake (Fig.3.3). This interpretation is further substantiated by significant uptake inhibition of lumiflavin and lumichrome, two riboflavin analogs that lack a D-ribose chain. Moreover, the lack of affinity of tricyclic antidepressants and folate demonstrates that the transporter uniquely recognizes flavin isoalloxazine moieties. Compared with unlabeled riboflavin, analogs with extension on N-10 side chain (phosphate in FMN and adenosine in FAD) exhibit lower but noticeably significant affinity to the transport system. Combined, these findings indicate that the D-ribose chain of riboflavin can serve as a potential modification site that least compromises ligand-transporter interactions. Consistent with previous studies in human syncytiotrophoblast microvillous membrane preparations (Moe et al., 1994) and other tissues (Said and Ma, 1994; Kumar et al., 1998; Said et al., 1998), no obvious requirements for N a\ Ca"^ or were found on the uptake of riboflavin in BeWo cell monolayers. Furthermore, riboflavin uptake is not sensitive to the Na^/K^-ATPase inhibitor (ouabain) and the Na*/lf^ exchanger inhibitor (amiloride), confirming the sodium and proton gradient- independent behavior of this transport system. 8 6 Interestingly, substitution of chloride ions in bathing media with other monovalent anions resulted in incomplete, yet significant reduction of riboflavin uptake. Comparable levels of riboflavin uptake attenuation by DIDS, a well-documented anion exchanger/ chloride channel inhibitor, further corroborates the potential role of Cl ions in the epithelial uptake of riboflavin. Unlike in other epithelia. Cl transfer across the placental membrane is Na* independent (Illsley et al., 1988). Consequently, this explains our observation that furosemide, a Na*-K^-2 Cl cotransporter inhibitor, has no effect on riboflavin uptake in BeWo cells. Three dominant mechanisms of Cl transfers across the placental microvillous membrane have been identified, namely an electroneutral DIDS- sensitive anion exchanger, a DIDS-sensitive Cl conductance, and a DIDS-insensitive voltage dependent Cl conductance (Stulc, 1997). The Cl dependence of riboflavin uptake in our studies is not attributed to the placental DIDS-sensitive anion exchanger. This is evidenced by i) capability of the placental anion exchanger to transfer other monovalent anions including f and SCN'(Stulc, 1997) and ii) the insignificant inhibition of riboflavin uptake by probenecid. Taken together, our results suggest for the first time that placental riboflavin uptake may be linked to a DIDS-sensitive Cl conductance. It should be noted that riboflavin uptake is not solely coupled to an electrochemical Cl 87 gradient, since removal of Cl- does not result in complete loss of uptake activity. The underlying mechanism behind Cl -coupled riboflavin uptake requires further investigation. It has been shown that differentiation of cytotrophoblasts is accompanied by changes in membrane transporter activities (Furesz et al., 1993; Liu et al., 1997). Our data show that riboflavin uptake in differentiated syncytiotrophoblasts induced by forskolin or 8-bromo-cAMP is significantly lower relative to proliferating cytotrophoblasts. Previously, Furesz and colleagues have reported a decreased alanine uptake via sodium-dependent amino acid transporters (ASC systems) in differentiated BeWo cells (Furesz et al., 1993). The reduced activities were attributed to apparent loss of the transporter proteins during extensive membrane remodeling processes throughout syncytiotrophoblast maturation and polarization. Currently, we cannot rule out the possibility that the diminished riboflavin uptake is a consequence of drug-induced modification of other cellular events. The specificity of riboflavin transporters in both undifferentiated and differentiated cell types remains unaltered; thus, reduced uptake could be attributed to decreased expression of the transport system during differentiation as a result of diminishing metabolic demands. Alternatively, higher riboflavin 8 8 requirements in cytotrophoblast stem cells may simply reflect a greater demand of nutrients for cellular proliferation. Several studies have demonstrated that activities of membrane transporter systems are rapidly regulated by the major signaling pathways, namely protein kinase A-, C-, and Ca'Vcalmodulin- mediated pathways (Racke et al., 1998; Braiman et al., 1999). Our results show that riboflavin uptake in BeWo cells appears to be modulated by cellular levels of cyclic nucleotides and calmodulin (CaM) but not protein kinase C pathway. Pretreatment of cells with PKA pathway stimulants or CaM antagonists cause marked decreases in uptake capacity and affinity (K,) of riboflavin absorption. The precise nature of the target sites in these signaling pathways is unknown, since the molecular identities of the riboflavin transporters are yet to be defined. The observations that short-term incubation (1 h) resulted in pronounced effects on riboflavin uptake eliminate the possible involvement of de novo biosynthesis of transporter mRNAs or proteins. Protein kinase A phosphorylation has been shown to reduce the Cl" conductance across placental microvillous membrane (Placchi et al., 1991), which allows us to speculate that PKA modulators exert their effect by changing the Cl'-coupling properties of riboflavin uptake. 89 Most signai transduction pathways are involved in diverse and critical functions of cells (Alberts et al., 1994). Using two CaM antagonists of different structural classifications enabled us to identify the specific involvement of CaM. Ineffectiveness of KN-93, a selective CaM-kinase H inhibitor, on blockage of riboflavin uptake further excludes the role of this ubiquitous CaM dependent kinase in the CaM- mediated signaling processes. The findings that activators targeting three distinct components of the protein kinase A cascade all lead to reduction of riboflavin uptake strongly suggests a significant relationship between intracellular levels of cAMP and cellular translocation of riboflavin. Interestingly, elevation of cGMP via pCPT-cGMP also results in a decrease of riboflavin uptake. Synergistic inhibition from co-incubation of pCTP-cGMP and IBMX further supports the involvement of both cyclic nucleotide secondary messengers in riboflavin uptake in BeWo cells. In contrast to cAMP, relatively little is known about the mechanism of action of cGMP on function of membrane transporters (Foreman and Johansen, 1996). Further studies are underway to disseminate the precise role of the PKG pathway and its involvement in the regulation in riboflavin uptake. In a series of studies. Said and coworkers found that riboflavin uptake in various tissues is modulated by different regulatory mechanisms. In Caco-2 cells, activation of 90 the PKA pathway leads to downregulation of riboflavin uptake (Said et al., 1994), whereas inhibition of the CaM- mediated processes attenuates uptake activities in renal, hepatic cell lines (Kumar et al., 1998; Said et al., 1998). Currently, the physiological significance behind the multiple- signaling regulatory mechanism involved in placental riboflavin uptake is not understood. Extensive cross-talk between cAMP and calmodulin- mediated signal transduction pathways exists at several levels of cellular control mechanism (Alberts et al., 1994); therefore, the observed cyclic nucleotide- and CaM- mediated reduction of riboflavin uptake in BeWo cells might be a manifestation of intertwined regulation of these processes. A similar phenomenon has been observed in the modulation of serotonin and dopamine transporter activities (Cool et al., 1991; Jayanthi et al., 1994; Ramamoorthy et al., 1995; Zhu et al., 1997; Batchelor and Schenk, 1998). In both cases, activation of PKA pathway increases the mRNA transcripts of transporters, but stimulation of PKC pathway impairs transporter activities without influencing the mRNA levels and membrane densities of transporters. In summary, our experiments demonstrate the presence of a high affinity riboflavin transporter system exists in the BeWo cell line. This active transport system does not require coupling to an electrochemical Na"^ gradient, corroborating previous observations 91 in microvillous membrane vesicles isolated from human syncytiocytotrophoblasts (Moe et al., 1994). We report, for the first time, that riboflavin uptake in BeWo cells is partly associated with a DIDS-sensitive Cl' conductance. Furthermore, riboflavin uptake appears to be modulated by cellular levels of cyclic nucleotides and calmodulin and cytotrophoblast differentiation induced by PKA activators is accompanied by decrease in riboflavin uptake activities. Combined, our data suggest that BeWo cells, which constitutively express a specific riboflavin transport system, may serve as a useful experimental model for understanding the human placental translocation of a vitamin that is essential for fetal development. 92 [■*H]-Rf uptake [“*C]-Ma uptake (% of control) (% of control) Control 100.00 ±5.15 100.00 ± 8.48 Choline -> Na* 91.33 ±4.33 99.66 ± 14.22 K+->Na+ 101.73 ±3.89 111.24 ±21.78 Li^ -> Na^ 101.97 ±5.79 105.65 ±23.62 1 mM Ouabain 93.08 ±8.61 101.74 ± 13.70 Ca"* free buffer 100.09 ± 2.45 109.89 ± 8.93 1 mM EGTA 95.67 ±5.12 91.63 ± 10.06 Control 100.00 ± 8.73 100.00 ± 18.66 I -> Cl 78.49 ±6.18*** 91.18 ±8.21 SCN -> Cl 66.85 ± 5.03*** 93.32 ± 13.16 Gluconate > Cl 84.39 ± 5.23** 80.54 ±9.32 [^H]-Rf uptake ["*C]-Ma uptake (% of pH 7) (% of pH 7) pH 3 93.94 ± 1.40 95.06 ±11.35 pH 4 96.95 ±4.00 88.24 ± 10.82 pH 5 97.12 ±3.72 92.49 ±5.13 pH 6 94.26 ±4.59 98.98 ± 12.72 pH 7 100.00 ±3.51 100.00 ± 11.85 pH 8 96.86 ±4.93 94.90 ±5.75 Table 3.1 Influence of ions^ and pH^ on riboflavin uptake in BeWo cell monolayers. ^ 5-day old BeWo cells were incubated with 5 nM [^H]-riboflavin (Rf) and 0.37 pM ['■*C]-mannitol (Ma) for 20 min either in control bathing medium (137 mM NaCl, 5.4 mM KCl, 5.4 mM CaCli, 1 mM MgSO^, 25 mM glucose, 10 mM HEPES) or in buffers in which NaCl was replaced with 137 mM of various inoiganic salts. Ca"* free buffer contains same composition of salts as in the control buffer except 0.1 mM EDTA and no CaCli were incorporated After a 20-min uptake study, cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100, and measured for radioactivity. Each value represents mean ± standard deviation of four experiments, p <0.001 versus control. ^ [^H]-riboflavin uptake studies were performed in bathing medium with pH values ranging from 3.0 to 8.0. 93 Inhibitor Concentration [^H]-Rf uptake ['“C]-Ma uptake (% of control) (% of control) Ami loride 0.5 mM 90.03 ±9.31 86.85 ±9.36 1.0 mM 105.02 ± 8.45 99.86 ± 17.84 Probenecid 0.5 mM 109.48 ± 18.03 87.98 ± 2.60 1.0 mM 121.89+ 15.17 96.83 ± 11.73 DIDS 0.1 mM 79.85 ±11.35* 87.24 ± 15.70 0.5 mM 74.47 ±6.61* 83.63 ± 16.76 1.0 mM 67.52 ± 12.99* 88.96 ± 17.63 Furosemide 0.5 mM 96.46 ± 10.24 95.46 ± 22.09 1.0 mM 101.69 ± 14.35 95.51 ±7.79 Table 3.2 Effect of anion exchange inhibitor and organic anion transporter inhibitors on riboflavin uptake in BeWo cell monolayers. 5 nM [^H]-riboflavin (Rf) and 0.37 pM ['‘*C]-mannitol (Ma) were added to 5-day old confluent BeWo cells pretreated with these inhibitors (10 min, 37°C). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100, and measured for radioactivity after a 20-min uptake study. Each value represents mean ± standard deviation of four experiments. * p< 0.05 versus untreated control 94 Inhibitor [^H]-Rf uptake [“*C]-Ma uptake (% of control) (% of control) PKG pathways pCPT-cGMP (250 pM) 76.88 ±5.14*” 109.24 ± 16.75 pCPT-cGMP (500 pM) 73.44 ± 4.64** 101.50 ±7.59 pCPT-cGMP (250 pM) + IBMX (2.5 mM) 43.28 ±6.21*** 84.73 ± 12.14 IBMX (2.5 mM) 51.07 ±6.31*** 94.38 ± 15.28 Table 3.3 Effect of pCPT cGMPon riboflavin uptake in BeWo cell monolayers. 5 nM [^H]-riboflavin (Rf) and 0.37 pM ['‘*C]-mannito! (Ma) were added to confluent BeWo cells pretreated with these inhibitors (ih, 37°C). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-1(X), and measured for radioactivity after a 20-min uptake study. Each value represents mean ± standard deviation of four experiments. p< 0.01, *** p< 0.001 versus untreated control 95 Inhibitor [^H]-Rf uptake ["C]-Ma uptake (% of control) {% of control) PKC pathways PMA (I pM) 99.45 ± 1.12 95.83 ± 16.71 PMA (5 pM) 99.18 ±3.70 116.61 ±12.71 PMA (10 pM) 97.05 ± 10.43 94.04 ± 8.92 PMA (50 pM) 97.86 ± 19.95 113.44 ± 18.47 Chelerythrine (1 pM) 112.41 ±14.12 97.29 ± 10.61 Chelerythrine (10 pM) 99.20 ± 14.12 95.06 ± 23.82 Chelerythrine (50 pM) 87.99 ±11.35 114.35 ± 12.92 Calmodulin pathways Calmidazolium (50 pM) 73.63 ± 10.99“ 84.58 ±11.66 Calmidazolium (125 pM) 19.69 ±4.31“ * 85.02 ± 7.37 W-13 (50 pM, 2h) 93.12 ±7.61 93.30 ± 17.41 W-13 (125 pM, 2h) 86.90 ±5.28“ 104.59 ± 17.03 W-13 (250 pM, 2h) 78.85 ±4.25*“ 97.06 ± 23.36 KN-93 (125 pM) 103.89 ± 12.05 120.61 ±4.05 KN-93 (250 pM) 94.21 ±6.67 103.32 ± 14.94 KN-92 (125 pM) 112.41 ±4.02 109.64 ± 12.42 KN-92 (250 pM) 110.68 ± 5.89 103.59 ± 13.60 Table 3.4 Effect of protein kinase C- and calmodulin- mediated pathways modulators on riboflavin uptake in BeWo cell monolayers. 5 nM [^H]-ribofIavin (Rf) and 0.37 pM ['‘*C]-mannitol (Ma) were added to confluent BeWo cells pre treated with these inhibitors (Ih, unless specified). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100, and measured for radioactivity after a 20-min uptake study. Each value represents mean ± standard deviation of four experiments. **p<0.01, ***pc0.001 versus untreated control 96 100 2.0 Q. I cç 1 t I 0 - 0.8 - 0.6 1 - 0.4 Dt - 0.2 0.0 0 5 10 15 20 25 30 35 40 Time (min) Fig. 3.1 Time course of [’H]*riboflavln (Rf) uptake in BeWo cell monolayers.5-day old BeWo cells were incubated with 5.0 nM [^H]-Rf (circles) and 0.37 pM [‘'‘C]-mannitol (diamonds). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-1(X) and measured for radioactivity at specified time points. Each value represents mean ± standard deviation of four experiments. 97 2000 I 300 i “ ° i I 200 1500 ISO I 100 Q. 1 50 25 f 1000 pHl-Rf concentration (nM) i 500 *5 i 3 0 100 200 300 400 500 600 700 800 900 1000 1100 concentration (nM) Fig. 3.2 Concentration dependence of [^H] riboflavin (Rf) uptake in BeWo cell monolayers.5-day old BeWo cells were incubated with [^H]-Rf (1-1000 nM) and [‘‘‘CJ-mannitol (0.37 jiM). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100 and measured for radioactivity after a 20-min uptake study. Each value represents mean ± standard deviation of four experiments. The solid line represents the calculated fit of the data to Eq. 1 as described in Methods. The estimated active component (Tactivc) to the total Rf flux (ytoui) is indicated with a dotted line; the estimated passive component (Vpassivc) is indicated with a dashed line. Inset shows the enlarged portion of the figure at concentrations below 20 nM. 98 120 I X % i i i # i# •uTT: T- MT r" "T Rf FMN FAD LF LC D-ribose Folate Chipmz Amtryp Fig. 3.3 EfTect of riboflavin structural analogs on uptake of [^H] riboflavin (Rf). Uptake of 5 nM [^H]-Rf and 0.37 pM ['^C]-mannitol were measured in the presence of 5 |xM various analogs (1000-fold excess). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100 and measured for radioactivity after a 20-min uptake study. Each value represents mean ± standard deviation of four experiments. LF: lumi flavin, LC: lumichrome, Chipmz: chlorpromazine, Amtryp: amitriptyline. 99 Cytotrophoblast Forskolin 8-Br-cAMP 24 hr 48 hr 72 hr Fig. 3.4 Time 100 Fig. 3.5 Forskolin induced differentiation of BeWo cells.2 day-old BeWo cells were grown in the presence of 100 pM forskolin (B, D) for 72h. Panels A, C showed the morphology of the control BeWo cells. Cells were processed with 3% Giemsa stain for 2 h at room temperature. Arrows indicated the multi-nucleated BeWo cells. 101 120 ■ 1 100 - I I “ ■ Of t “ ■ B S 40 ■ L. 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 IBMX concentration (mM) 400 Conlrol 350 • IBMX 300 - 250 ■ 200 - 150 • 100 ■ SO - 0 S 10 IS 20 [’H]-R( concentration (nM) Fig. 3.6 Effect of isobutylmethyixanthine (IBMX) on [^li]>rtboflavin (Rf) in BeWo cell monolayers. A. BeWo cells were incubated with varying concentrations of IBMX for ih, after which a 20-min uptake was performed with 5 nM [^H]-Rf and 0.37 jaM ['‘*C]-mannitol. B. Concentration dependent uptake of [^H]-Rf uptake in IBMX-treated cells. [^H]-Rf (1-20 nM) and [‘‘*C]-mannitol (0.37 iiM) were added to 5-day old control cells (circles) or cells pre-treated with 5 mM IBMX for Ih (diamonds). The passive diffusion component obtained using Eq. 1 in Methods was subtracted from total uptake values and only active uptake was shown. Each value represents mean ± standard deviation of four experiments. 102 3 450 1 • Control I ^0 A Calmidazolium ^ 350 Q. 0 » 300 Io E 250 I 200 I 150 100 *5 50 Î 0 20 40 60 80 100 [®H]-Rf concentration (nM) Fig. 3.7 Effect of calmodizalium on [^H]«riboflavin (Rf) in BeWo cell monolayers. [^H]-Rf (1-100 nM) and [‘‘*C]-mannitol (0.37 jiM) were added to 5-day old BeWo cells pre-treated with 125pM calmodizalium for Ih. Concentration dependent uptake of [3H]-riboflavin is presented in control (circles) and calmidazolium- treated (diamonds) cells. Data points are corrected for passive diffusion component obtained using Eq. 1 in Methods by subtraction from total uptake values to present only the saturable uptake process. Each value represents mean ± standard deviation of four experiments. 103 REFERENCE Alberts B, Bray D, Lewis J, Raff M, Roberts K and Waston JD ( 1994) Molecular biology of the cell. Garland Publishing, Inc. New York. 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Wice B, Menton D, Geuze H and Schwartz AL( 1990) Modulators of cyclic AMP metabolism induce syncytiotrophoblast formation in vitro. Exp Cell Res 186:306-316. Zempleni J, Link G and Bitsch 1 (1995) Intrauterine vitamin B2 uptake of preterm and full-term infants. Pediatr Res 2%:5%5-59\. Zempleni J, Link G and Kubler W (1992) The transport of thiamine, riboflavin and pyridoxal 5 -phosphate by human placenta. IntJVttam Nutr Res 62:165-172. Zhu SJ, Kavanaugh MP, Sonders MS, Amara SG and Zahniser NR (1997) Activation of protein kinase C inhibits uptake, currents and binding associated with the human dopamine transporter expressed in Xenopus oocytes. J Pharmacol Exp Ther 282:1358-1365. 107 CHAPTER 4 SUBCELLULAR LOCALIZATION OF RIBOFLAVIN: INVOLVEMENT OF ENDOCYTIC ORGANELLES ABSTRACT Previous studies by our laboratory have suggested the potential role of receptor- mediated endocytosis components in the cellular translocation of riboflavin. To delineate the intracellular compartments and events involved in the internalization of riboflavin, we synthesized rhodamine-riboflavin conjugate to monitor its movement via fluorescent microscopy. Cellular uptake studies in BeWo cells show that rhodamine-riboflavin conjugate exhibits similar ligand affinity towards the putative riboflavin transporters as [^H]-riboflavin, whereas rhodamine does not significantly interfere the ligand-transporter interaction. Microscope analysis reveals rapid internalization of the rhodamine-riboflavin conjugate via a riboflavin-specific process into acidic vesicular compartments throughout the cells. The intracellular punctate distribution is comparable to that of unconjugated riboflavin and FITC-transferrin, a well-characterized receptor-mediated endocytosis substrate. Double-labeling fluorescent studies further confirm that with 10 min of internalization rhodamine-riboflavin conjugate substantially concentrates within vesicular 108 stractures associated with clathrin, tab 5 and FFTC-transferrin. Following prolonged incubation, most rhodamine-riboflavin conjugate is distributed to LAMP-1 containing organelles whereas a small subpopulation of conjugates remains in LAMP-1- negative regions. In summary, our studies provide morphological evidence of the involvement of endocytosis machinery in the intracellular trafficking of riboflavin. The subcellular localization of rhodamine-riboflavin conjugate suggests that the internalization of riboflavin is very likely to follow a classical receptor-mediated endocytosis pathway. 109 INTRODUCTION Riboflavin, also known as vitamin Bi, plays an essential role in normal cellular growth and development (Rivlin, 1975). Like most water-soluble nutrients, studies have shown that specific transporter systems are involved to facilitate efficient entry of riboflavin across the cell membrane (Said and Arianas, 1991; Said et al., 1998; Huang and Swaan, 2001). Since the molecular identity of the riboflavin-translocating protein(s) remains elusive, most mechanistic studies focused mainly on biochemical characterization of the putative transporter(s) through determinants such as energy- or ion-dependence. Based on these studies, the generally accepted model is that riboflavin is taken up into the cells via an active carrier-mediated pathway (Said and Arianas, 1991; Rindi and Gastaldi, 1997). Although these biochemical criteria are very useful in identifying an active transport process, it is known that they cannot reliably discriminate the underlying mechanisms between carrier-mediated transport and receptor-mediated endocytosis pathway (Spinella et al., 1995). Recently, we have demonstrated that riboflavin transport in the human small intestinal Caco-2 cell line is sensitive to pharmacological agents known to alter endocytic pathways (Huang and Swaan, 20(X)). The results indicated that microtubule-based movements and vesicular-sorting components are essential for cellular transfer of riboflavin. Previously, Low and coworkers have showed that internalized bovine semm albumin-riboflavin conjugate has a distribution pattern reminiscent of endosomal compartments (Wangensteen et al., 1996; Holladay et al., 1999) although in these studies riboflavin failed to inhibit surface binding of BS A-riboflavin. The finding that BSA- riboflavin only partially blocked riboflavin uptake further suggested a nonspecific 110 internalization mechanism might exist to accommodate this macromolecolar riboflavin adduct (Holladay et al., 1999). Direct evidence of an endocytosis mechanism usually entails microscopic analysis of subcellular distribution of ligands and receptors (Pastan and Willingham, 1985). Since the riboflavin transporters have yet to be identified, to elucidate the involvement of receptor-mediated endocytosis we aim to monitor the cellular trafficking of riboflavin itself. Although riboflavin is intrinsically fluorescent, its emission spectrum is difficult to distinguish from cellular autofluorescence (Andersson et al., 1998). Moreover, endogenous riboflavin and riboflavin-related coenzymes might further complicate signal interpretation. To avoid these complications, in the present study, we develop a strategy to conjugate riboflavin with rhodamine, a widely used red fluorescent probe. The intracellular itinerary of riboflavin is investigated by following the movements of rhodamine-riboflavin conjugate under fluorescent microscope. We choose human choriocarcinoma-derived cell line, BeWo, as our system. Recently, we have reported the existence of high-afUnity riboflavin transporter(s) on the microvillous membrane of this polarized trophoblast model (Huang and Swaan, 2001). Compared to Caco-2 cells. Be Wo cells have a better-defined nucleus and cytoplasmic boundary together with a more homogenous cell population. More importantly, our studies showed that at physiological riboflavin plasma concentration (~ 5 nM), riboflavin uptake activity in BeWo cells is three-fold higher than that in Caco-2 cells (unpublished data, Huang and Swaan). I ll METHODS Materials Tetramethyl-rhodamine-carbonyl-azide, LysoTracker™ Blue-White-DPX, DAPI and SlowFade® light anti fading kit were purchased from Molecular Probes (Eugene, OR). Cell culture materials and buffer solutions were obtained from Invitrogen Life Technology (Carlsbad, CA). Monoclonal antibodies to human clathrin heavy chain, rabS, and LAMP-1 were from BD Biosciences Transduction Laboratories (San Diego, CA). FTTC- and TRUC- conjugated, Fc specific, goat anti-mouse IgG antibodies, FTTC conjugation kit, human transferrin, [^H]-ribofIavin (20 Ci/mmol) and [‘'‘C]-mannitol (60 mCi/mmoI) were purchased from Sigma (St. Louis, MO). All other chemicals were from Fisher Scientific (Pittsburgh, PA). Cell cultures BeWo cells were obtained from American Type Culture Collection (Manassas, VA). Cells were maintained at 37 °C, under 5% CO2, in complete medium consisting of F-12K medium with 10% fetal bovine serum, 1% non-essential amino acids, 100 U/ml of penicillin, and ICO pg/ml of streptomycin. The culture medium was replaced every other day. The cells were harvested at 80% confluence (day 4 to 5) by exposure to a trypsin- EDTA solution. Synthesis of rhodamine-riboflavin coi^ugates 6 mg Riboflavin and 0.8 mg tetramethyl-rhodamine-carbonyl-azide (10:1 molar ratio) was dissolved in DMSO and heated at 80 °C for Ih with gentle stirring. After Ih, 40 ^1 absolute ethanol was added in the reaction mixture and reacted for an additional 20 112 min to terminate residual isocyanates. Rhodamine-riboftavin conjugates were further purified by a Beckman System Gold HPLC system (Fullerton, CA) using a Beckman RP C-18 preparative column (ODS 5^tm, 10 mm x 25 cm). Crude reaction mixtures were eluted using a linear gradient of 0-100% acetonitrile over 30 min (3 ml/min) and the eluents were collected by a fraction collector (1.5 ml/min). A Beckman 166P detector operating at 267 nm and an Applied Biosystems 980 programmable fluorescence detector (Xe*: 545 nm, Xen,: 580 nm; Foster City, CA) were used to detect riboflavin and rhodamine respectively. Comparison of the aligned UV and fluorescence chromatograms revealed rhodamine-riboflavin conjugate and riboflavin have a retention time around 6 and 11 minute respectively. Other rhodamine byproducts are eluted after 14 minute (data not shown). Rhodamine-riboflavin conjugate was subjected to mass spectrometry and fluorescent absorption scanning spectrum analysis using a Perkin-Elmer Luminescence spectrometer LS 50 (Foster City, CA). Mass spectrometry indicated a Mr of 804.3. Compared to spectra of standard compounds, rhodamine-riboflavin shows combined riboflavin and rhodamine absorbance patterns with a slight right-shift in Xabs for the rhodamine fluorophore (15 nm). Uptake studies BeWo cell monolayers were grown on rat-tail collagen-coated 12-well plates at a density of 5 x 10“* cells/cm^. Confluent monolayers were formed between 3 to 5 days after seeding and were used for experiments at that time. Before studies were initiated, BeWo cell monolayers were washed twice with warm (37 °C) PBS (pH 7.4). Riboflavin uptake studies were performed at 37 °C in bathing medium (Hank’s balanced salt solution 113 containing 25 mM glucose and 10 mMHEPES, adjustedto pH 7.4) with a final concentration of 5 nM [^H]-riboflavin in the absence (control) or presence of rhodamine- riboflavin conjugate. ['"^Cj-mannitol (0.37 nM) was incorporated in the incubation medium to determine the specificity of the washing steps. After 20 min, bathing medium was aspirated, and cells were washed twice with ice-cold PBS (pH 3.0) to remove free and surface-bound riboflavin. Finally, cells were lysed with 1% Triton X-100 solution, and the amount of dual-labeled radioactivity in cell lysates was quantitated using a Beckman liquid scintillation counter (model LS 6000IC). Subcellular localization of rhodamine-riboflavin conjugate For internalization experiments, BeWo cells were grown on Becton Dickinson Falcon CultureSlides (Bedford, MA) at a density of 5,000 cells/cm^. Subconfluent BeWo cells (3 day old) were briefly rinsed with ice-cold PBS, and incubated with fluorescent ligands (500 nM of rhodamine-riboflavin, riboflavin or rhodamine; 25 pg/ml iron- saturated FTTC-transferrin in bathing medium) at 4 °C for 2 h to allow equilibrium binding. After preequilibration, cells were washed three times with ice-cold PBS to remove unbound substrate and chased with prewarmed ligand-free bathing medium at 37 °C for 10 min to allow internalization. Cells were then fixed with 4 % paraformaldehyde (in PBS with 4 % sucrose) at room temperature for 20 min, incubated with 300 nM DAPI for 10 min, and mounted in SlowFade anti fading reagent. Samples were sealed with nail polish and examined with a Nikon Eclipse 800 fluorescent microscope equipped with FTTC (Xex: 460-500 nm, Xem: 505-560 nm, dichroic splitter: 505 nm), rhodamine (Xe*: 530-560 nm, Xem: 590-650 nm, dichroic splitter: 570 nm) and DAPI (Xex 340-380 nm, Xem: 114 435-485 nm, dichroic splitter: 40&ran> filters, bnages were captmed with a Micromax cooled CCD camera (Roper Scientific Inc., Trenton NJ) and IPLab software (Scanalystics, Inc. Fairfax VA) using a constant exposure time at each filter combination. Composite images were colored and assembled in Adobe Photoshop 5.5 (Adobe Systems Inc., Mountain View, CA) with no alterations in the relative gray scale levels. Colocalization studies For colocalization with LysoTracker™, BeWo cells were pulsed with 500 nM rhodamine-riboflavin at 4 °C for 2 h. After three-time ice-cold PBS wash, cells were incubated with prewarmed bathing medium containing 300 nM LysoTracker™ Blue- White-DPX at 37 °C for 10 min. Cells were then immediately fixed with 4 % paraformaldehyde and mounted in SlowFade. Before immunoflourescent studies the specificity of all primary antibodies were confirmed by Western blot analyses. Compared with the positive control cell lysates (supplied with the monoclonal antibodies by Transduction Lab), BeWo cells express high levels of clathrin heavy chain, rab5, and LAMP! proteins and all antibodies were proved to be highly specific (data not shown). The absence of cross-reactivity of the secondary antibodies was also verified by omitting the primary antibodies during immunofluorescent studies. For colocalization with clathrin or rabS protein, BeWo cells were incubated with 500 nM rhodamine-riboflavin or 25 pg/ml iron-saturated FTTC-transferrin at 4 °C for 2 h and chased with prewarmed ligand-free bathing medium at 37°C for 10 or 30 min. After paraformaldehyde fixation, cells were washed twice with 25 mM glycine, and 115 pcrmeabKzed with PBS containing (h2%^TritoitX-tGO and 1% BSA for 2d min. Clathrin or rabS was visualized using a monoclonal mouse anti-human clathrin heavy chain or anti-human rabS antibody (1:250,1 h) followed by Fc specific, FTTC-or TRTTC- labeled goat anti-mouse IgG (1:200, ih). Cells were then counterstained with 300 nM DAPI, and mounted in SlowFade. To visualize LAMP-1 proteins, a monoclonal mouse anti-human LAMP-1 antibody (1:200, Ih) was used. For colocalization with FTTC-transferrin, cells were incubated with 500 nM rhodamine-riboflavin and 25 pg/ml iron-saturated FTTC-transferrin simultaneously at 4 °C for 2 h. After ice-cold PBS wash, cells were incubated with ligand-free medium at 37 °C for 15 min. Cells were immediately fixed with 300 nM DAPI for 10 min, and mounted in SlowFade. 116 RESULTS Synthesis of rhodamine-riboflavin coiyugate Compared to bioconjugation of macromolecules, direct fluorochrome labeling of small chemical entities such as riboflavin (Mw 376.4) is more challenging. The limited molecular size not only reduces flexibility for adduct linkage, but increases the risks of modifying the biological affinity of ligands due to impending steric hindrance. Potential conjugatable moieties in riboflavin include the hydroxyl groups of the D-ribose chain and the isoalloxazine nitrogen groups (Fig.4.1). Based on our previous studies with riboflavin-structural analogs, the D-ribose side chain plays an insignificant role in the interaction between riboflavin and its placental transporter when compared to the isoalloxazine ring (Huang and Swaan, 2001). Thus, conjugation to the ribose side chain would have a minimal effect on ligand-receptor interactions. Rhodamine was linked to the D-ribose chain of riboflavin using tetramethyl- rhodamine-5-carbonyl-azide (TMRCA). Upon heating of TMRCA (Fig 4.1) its acyl azide group undergoes a Curtius rearrangement to yield an isocyanate group, which reacts with primary hydroxyl groups to form a carbamate linkage (Adams, 1942; Takadate et al., 1985). Since the remaining secondary nitrogen and hydroxyl groups on riboflavin are less reactive, the only primary-OH on Cs of D-ribose is expected to be the preferred target for nucleophilic attack of isocyanate. Indeed, after a Ih synthesis reaction only one predominant peak with significant absorbance under both riboflavin and rhodamine wavelengths was detected in our HPLC analysis (data not shown). The chemical identity of the particular eluent was further verified as rhodamine-riboflavin conjugate utilizing fluorescent scanning spectrum and mass spectrometry analyses described in methods. 117 Substrate specifTcfty orrhodamine To examine the biological specificity of rhodamine-riboflavin conjugate, we used BeWo cells, a human choriocarcinoma cell line. Previously, we have reported that high- affinity riboflavin transporter(s) are functionally expressed on the microvillous membrane of these cells (Huang and Swaan, 2001). Uptake experiments in BeWo cell monolayers revealed that 75.6% of [^Hj-riboflavin uptake was blocked in the presence of 1000-fold rhodamine-riboflavin, whereas rhodamine did not show significant effect (Fig. 4.2A). Rhodamine-riboflavin conjugate was equally effective in inhibiting [^H]-riboflavin uptake compared with unlabeled riboflavin at various concentrations (Fig 4.2A, B), indicating that the conjugates have similar ligand affinity towards riboflavin transporter(s) comparable to that of riboflavin. Internalization and subcellular localization of rhodamine-riboflavin conjugate To analyze the internalization of rhodamine-riboflavin, cells were preincubated with the conjugate at 4 °C for 2 hr then chased with ligand-free medium at 37 °C. The subcellular distribution of rhodamine-riboflavin was visualized by fluorescent microscopy. Compared with nuclear DAPI stain, after 10 min the rhodamine signals resided intracellularly in the perinuclear punctate regions indicating efficient internalization and possible cytosolic accumulation of the conjugate (Fig. 4.3). FTTC-transferrin, a well-characterized receptor-mediated endocytosis substrate, was used to identify the characteristic morphology of endosomal compartments in BeWo cells. Unlike other polarized epithelia, BeWo cells express high numbers of transferrin receptor on their apical cell surface and a specific endocytic pathway has been identified for internalization of transferrin (Cemeus and van der Ende, 1991). After 10 min, FFFC- 118 transfemn (Fig. 4.4A) exhibited a comparable punctate distribution as observed previously with rhodamine-riboflavin conjugate (Fig. 4.3A). In a parallel study, Internalization of rhodamine alone was also examined to assess the basal level of nonspecific endocytosis processes in BeWo cells under the experimental condition. Contrary to the distinct localization patterns of rhodamine-riboflavin conjugate or FTTC- transferrin, the majority of rhodamine signal was randomly distributed in the cells (Fig 4.4D). A generally weak and hazy staining was observed throughout the cells indicating nonspecific background adsorption. Moreover, when BeWo cells were incubated with the same concentration of riboflavin, a distinct intracellular punctate staining pattern (Fig. 4.4G) was detected resembling that of rhodamine-riboflavin conjugate (Fig. 4.3A). The specificity of rhodamine-riboflavin conjugate binding to the surface riboflavin transporter was further investigated under microscope. In these experiments BeWo cells were preincubated with rhodamine-riboflavin conjugate in the presence of 100 pM riboflavin at 4 ®C for 2h. Compared with the cells labeled with rhodamine- riboflavin conjugate alone (Fig. 4.5A), significantly lower fluorescent intensities were detected in cells preincubated with both riboflavin and the conjugate (Fig. 4.SD). Since excess riboflavin competed for limited riboflavin binding sites on the BeWo cell membrane (Huang and Swaan, 2(X)1), these results provided additional support for the presence of a transporter system that specifically mediates the internalization of rhodamine-riboflavin conjugate into BeWo cells. 119 ColocaHzation studies^ with ait acidotroptc marker, cfathrin-heavy chafai, rah 5, and FITC-transferrln To examine the cellular identity of the vesicular structures containing rhodamine- riboflavin, cells were treated simultaneously with rhodamine-riboflavin conjugate and LysoTracker Blue-white DPX, a membrane diffusible probe accumulating in acidic organelles. This acidotropic marker has been used successfully to label compartments involved in endocytosis processes (Bucci et al., 2000). After 10 min, significant overlap of rhodamine-riboflavin and LysoTracker signals (as indicated by pink color) revealed that the vesicular structures associated with rhodamine-riboflavin were acidic (Fig. 4.6). In most mammalian cells, uptake of receptor-bound ligands results mainly from clathrin-dependent endocytosis (Mellman, 1996). To investigate the potential role of clathrin in rhodamine-riboflavin internalization, we performed dual-labeling studies with monoclonal antibodies against human clathrin heavy chain. The signals of clathrin were visualized with fluorochrome-conjugated secondary antibodies. As expected, clathrin was present from the cell periphery and throughout the cell interior (Fig. 4.7 C, G) (Tolbert and Lameh, 1996). After 10 min, a high degree of colocalization of rhodamine-riboflavin conjugate with clathrin in the cytoplasma was observed (Fig. 4.7D). In a parallel study, FTTC-transferrin, a substrate internalized via the clathrin-mediated endocytosis pathway (Welch, 1992), was incorporated as a positive control for the immunofluorescent detection. As shown in Fig. 4.7 H, significant colocalization of FTTC-transferrin and clathrin verified the specificity of our immunostaining. In addition to clathrin, we also used anti-rabS antibody and FTTC-transferrin to further delineate the involvement of clathrin-mediated endocytosis in internalization of 120 rhodamfne-riboftavîn conjugate. Rab 5, a smairGTP binding protein, resides only in clathrin-coated vesicles (Fig. 4.81) and early endosomes (Rhodaman and Wandinger-Ness, 2000). Throughout its entire endocytic cycle, transferrin remains bound to transferrin receptor and majority of the complex is known to confine in early endocytic compartments (Welch, 1992). Consistent with the literature report (Vandenbulcke et al., 2000), rab 5 signals were concentrated in punctate vesicular compartments (Fig. 4.8C, G, K). After 10 min of internalization, almost all rabS-labeled regions exhibited strong rhodamine-riboflavin conjugate staining (Fig. 4.8D). In line with our previous findings (Fig. 4.3,4.4,4.8), in cells preincubated simultaneously with rhodamine-riboflavin conjugate and FITC-transferrin, extensive colocalization of signals was also observed in the perinuclear endosomes after IS min incubation (Fig. 4.9). Overlapped staining of FTTC-transferrin and rab 5 further corroborated these findings (Fig. 4.8E-H). Colocalization studies with LAMP-1 proteins To characterize the late intracellular routing of rhodamine-riboflavin conjugate, immunofluorescent studies were performed using antibodies against LAMP-1, a marker for late endocytic compartments including late endosomes and lysosomes (Mellman, 1996; Mukhetjee et al., 1997). As shown in Fig. 4.10B, E, H, late endocytic compartments of BeWo cells not only concentrate in the juxtanuclear area of the cytoplasm but also spread out towards the cell periphery. Following 30 min of internalization, majority of the conjugate is colocalized with LAMP-1 (solid arrow. Fig. 4. IOC, 10. However, few vesicles consistently appeared red in cells (lined arrows), suggesting that a subpopulation of rhodamine-riboflavin conjugate exists in LAMP-1- 121 negative organelîes. In contrast, after 30 min o f uptake, internalized FTTC-transferrin signals were evidently excluded from the LAMP-1 positive vesicles (Fig 4.101). 122 DISCUSSION The present study demonstrates that the endocytic machinery is involved in the internalization of rhodamine-riboflavin conjugate via a highly specific riboflavin internalization mechanism. This is evidenced by: 1) riboflavin-specific binding of rhodamine-riboflavin conjugate, 2) a punctate endosome-like subcellular distribution of the conjugate, 3) selective sequestration of rhodamine-riboflavin conjugate into acidic vesicular organelles, 4) colocalization of conjugate within early endocytic compartments containing FTTC-transferrin and immunostained positively for clathrin and rabS, and 5) sorting of conjugate into LAMP-1 -immunoreacti ve late endocytic components following prolonged internalization. In contrast to the nonspecific internalization of BSA-riboflavin reported by Low and coworkers (Holladay et al., 1999), our results show that rhodamine-riboflavin conjugate significantly blocks [^H]-riboflavin uptake (Fig. 4.2). Specific membrane binding of rhodamine-riboflavin conjugate is also substantially reduced in the presence of riboflavin (Fig. 4.5). The mutual inhibition between riboflavin and rhodamine-riboflavin conjugate strongly indicates that the internalization of conjugate is mediated by membrane surface riboflavin receptors. Importantly, the preserved specificity and high affinity of rhodamine-riboflavin conjugate towards the riboflavin transporters further confirms the insignificant role of D-ribose chain in the ligand-transporter interactions (Huang and Swaan, 2001). Receptor-mediated endocytosis plays an essential role in selective uptake of nutrients, growth factors and hormones into most cells (Mukheqee et al., 1997). In most cases, the endocytosis process is initiated from clathrin-coated pits and subsequently the 123 tigands/receptors are internalizedin clathrin-coated vesicies (Mellman, 1996). Our results show that rhodamine-riboflavin conjugate is detected in clathrin-positive clusters after short-term incubation (Fig. 4.7) suggesting the formation of clathrin-coated vesicles in the internalization of riboflavin. Colocalization of rhodamine-riboflavin conjugate with FITC-transferrin, a well-characterized substrate of clathrin-mediated endocytosis, also supports this finding (Fig. 4.9). Clathrin-mediated endocytosis usually involves concentration of receptors in the clathrin-coated pits with most receptors containing a conserved internalization motif in their cytoplasmic domain (Hirst and Robinson, 1998). Currently, it is unknown if the present results reflect that the putative riboflavin receptor shares such a structural feature and is localized in coated pits. Further investigations are necessary to elucidate the role of clathrin in the endocytosis of riboflavin. Interestingly, previously we have shown that [^H]-riboflavin uptake was not affected by phorbol-12- myristate acetate, a protein kinase C activator known to specifically block caveolae- dependent endocytosis (Smart et al., 1994; Huang and Swaan, 2001). Whether riboflavin is internalized exclusively from the clathrin-coated membrane awaits further verification Following the clathrin-mediated endocytosis pathway, internalized ligand/receptors are delivered rapidly into early endosomal compartments (Mellman, 1996). Our results show that within 10 min of incubation, signals of rhodamine-riboflavin conjugate are noticeably accumulated within punctate perinuclear vesicular organelles reminiscent of endosomal compartments (Fig. 4.3). The acidic nature of these structures also supports our morphologic observations (Fig. 4.6). More importantly, colocalization of conjugate with FITC-transferrin and rab-S further confirms the molecular identities of these compartments as early endosomes (Fig. 4.8,4.9). 124 After 30 min of intemaUzatfon, the majority of rhodamine-ribofiavin conjugate is detected in LAMP-l-positive organelles (Fig. 4.10). The late endosomal sorting of conjugate is evidently in contrast to the itinerary of FITC-transferrin, in which most transferrin signals are retained exclusively in early endosomes. The distinct staining patterns of FITC-transferrin and LAMP-1 further confirm the existence of separated early and late endocytic compartments in BeWo cells, and validate our analysis. Currently, the molecular identities of those LAMP-1-negative vesicles are unknown and require further investigation. After ligand-receptor complexes are internalized into endocytic vesicles, they might undergo several sorting scenarios. The fates of ligands and receptors can vary significantly according to the specific type of receptors (Mukheijee et al., 1997). Most G- protein coupled receptors will release their ligands in the early endosomes and then recycle back into the cell membrane, whereas receptors like neurotension receptor will be delivered to lysosomes together with their ligands (Vandenbulcke et al., 2000). Most clathrin-mediated endocytosis ligands are generally released from their receptors in early endosomes. The dissociation of ligands and receptors is resulted from pH-dependent comformational change of receptors induced by the acidic climate of early endosomes. Previously, we have shown that dissociation of surface-bound riboflavin in Caco-2 cells is pH-dependent with significantly higher riboflavin release at acidic pH (pH 3-5) (Huang and Swaan, 2000). Taken together, since rhodamine-riboflavin conjugate is not designed to bind to the putative riboflavin transporters irreversibly, it is likely that upon entry of early endosomes, the rhodamine-riboflavin conjugate will dissociate from the transporter. 125 Accordîngfy, riboflavin transporter mighrnot travef in parattef with the conjugate into the late endosomes. The current observation that rhodamine-riboflavin conjugate is sorted from early endosomes to late endocytic compartments is also in line with our recent findings with endocytosis perturbants (Huang and Swaan, 2(XX)). Previously, we showed that nocodazole, a microtubule-depolymering agent, significantly inhibited the apical uptake of [^H]-riboflavin. Since microtubules are required for the maturation of early endosomes to late endosomes, disruption of microtubular network is expected to deter the endosomal movement (Mukheijee et al., 1997). Consequently, intracellular accumulation of riboflavin in nocodazole-treated cells would be substantially reduced. For growth factors such as gastrin releasing peptide, late endosomal sorting, subsequently lysosomal degradation of ligands is essential for termination of the triggered cellular response (Grady et al., 1995). At physiological concentrations (~ nM range) riboflavin itself does not exhibit any intrinsic activity, however, studies have reported that massive riboflavin supplement can protect against cerebral ischemic damage (Hultquist et al., 1993; Betz et al., 1994). Using cell culture and rat tissues, Daly and colleagues further demonstrated that riboflavin, at low pM concentrations, significantly inhibits adenylate cyclase and guanyl nucleotides turnover of G-proteins and is an antagonist of A,-adenosine receptors (Daly et al., 1997). Currently, it is unknown whether the late endosomal sorting of riboflavin represents a regulatory mechanism against its potential pharmacological effects. It should be noted that our present studies with riboflavin-rhodamine conjugate may not reflect the intracellular metabolic fate of native riboflavin. Like other B-vitamins, riboflavin exerts its biological functions via its coenzyme forms flavin mononucleotide 126 (FMN) and flavin adenine dinucteotide (FAD) (Rivlin, 1975). As documented in the hepatocytes, upon entry of the cells, riboflavin will be converted into FMN and FAD by cytoplasmic flavokinase and FAD synthetase (McCormick and Zhang, 1993). Because these biotransformation processes require a free 5’ hydroxyl group on the D-ribose chain of riboflavin, the rhodamine-riboflavin conjugate is likely to bypass the endogenous metabolism. In summary, our studies present a novel method to synthesize a rhodamine-tagged riboflavin. Since the strategy preserves the essential isoalloxazine moiety of riboflavin, the rhodamine-riboflavin conjugate successfully retains its specificity and affinity toward the putative riboflavin transporters. Using the conjugate as a probe, we report, for the first time, morphological evidence of the involvement of a classical endocytosis mechanism in the internalization of riboflavin. Our current finding, though, does not rule out the existence of a carrier-mediated transport mechanism in cellular translocation of riboflavin, it clearly identified the endocytic compartments involved in the intracellular trafficking of riboflavin. This subcellular localization and distribution pattern will aid in our understanding of the cellular riboflavin disposition and future development of drug targeting approach via the riboflavin transporter(s). 127 CHj .CHa .CHa HaC' h e a t ao®c HN. N-C“ ° HOHaC^ Tetramethyt-rhodamine-carboylazide Rhodamine-Riboflavin conjugate Ritx>navin Figure 4.1 Synthesis of rhodamine-riboflavin conjugate 128 120 C 100 *** *** Q . 20 Control Rho-Rf Rhodamin 22 " 2 20 : Rho-Rf 16 - eo 10 - B DI 0 1000 2000 3000 4 0 0 0 5000 [Inhibitor] (nM) Figure 4.2 Effect of rhodamine-riboflavin coi\jueate and rhodamine on riboflavin uptake. A, Uptake of [^H]-riboflavin (5 nM) and [ C]-mannitoI (0.37 pM) were measured in the presence of 5 pM of riboflavin (Rf), rhodamine-riboflavin conjugate (Rho-Rf) or rhodamine (Rho). Cells were washed twice with ice-cold acidic PBS, lysed with 1% Triton X-100, and measured for radioactivity after a 20 min uptake study. B, concentration-dependent inhibition of rhodamine-riboflavin conjugate on [^H]-riboflavin uptake in BeWo cell monolayers. Uptake of [^H]-riboflavin (S nM) was measured in the presence of various concentrations of rhodamine-riboflavin conjugate or riboflavin. Each value represents the mean ± s.d. for four experiments. *** p < 0.001 versus controls. 129 Figure 4.3 Internalization of rhodamine riboflavin.Rhodamine-riboflavin conjugate were incubated with BeWo cells at 4 °C for 2h. Unbound conjugates were washed off, and cells were warmed to 37 °C for 10 min. Cells were then fixed immediately, incubated with nuclear DAPI stain and processed for fluorescence microscopic analysis. Panels show distribution patterns of rhodamine-riboflavin conjugate (A), and nuclear stain (B) in the identical optical field. Panel C displays the superimposed images from panel A and B. 130 Figure 4.4 Internalization of FITC-transferrin, rhodamine or riboflavin.BeWo cells were incubated with FITC-transferrin (Tf) or rhodamine or riboflavin at 4 °C for 2h. Unbound ligands were washed off, and cells were warmed to 37 for 10 min. Cells were then fixed immediately, incubated with nuclear DAPI stain and processed for fluorescence microscopic analysis. Panels show distribution patterns of riboflavin (G), FTTC-Tf (A), rhodamine (D), and riboflavin (G) and nuclear stain (B, E, H) in the identical optical field. Column 3 displays the superimposed images from column 1 and 2. 131 Figure 4.5 Specific binding of rhodamine-riboflavin.BeWo cells were incubated with 500 nM rhodamine-riboflavin conjugate in the absence (A) or presence (D) of 100 pM riboflavin at 4 °C for 2h. Unbound conjugates were washed off, and cells were warmed to 37 °C for 10 min. Cells were then fixed immediately, incubated with nuclear DAPI stain and processed for fluorescence microscopic analysis. Panels show distribution patterns of rhodamine-riboflavin conjugate (A, D), and nuclear stain (B, E) in the identical optical field. Column 3 displays the superimposed images from column 1 and 2. 132 Figure 4.6 Colocalization of rhodamine-riboflavin with acidotropic probe LysoTrackerBlue-White-DXD. BeWo cells were incubated with rhodamine-riboflavin conjugate at 4 °C for 2h. Unbound conjugates were washed off, and cells were labeled with LysoTrackerBlue™ at 37 °C for 10 min. Cells were then fixed immediately and processed for fluorescence microscopic analysis. Panels show distribution patterns of rhodamine-riboflavin conjugate (A), and LysoTrackerBlue™ (B) in the identical optical field. Panel C displays the superimposed images from panels A and B. Arrows indicate spots containing internalized rhodamine-riboflavin conjugate and LysoTracker signals. 133 Fig 4.7 Colocalization of rhodamine*riboflavin with clathrin.BeWo cells were incubated with rhodamine-riboflavin conjugate or FTTC-transferrin at 4 °C for 2h. Unbound ligands were washed off, and cells were warmed to 37 °C for 10 min. After fixation and permeablization, cells were sequentially stained with anti-clathrin heavy chain monoclonal antibody followed by FTTC- or TRTTC- labeled secondary antibody and DAPI nuclear stain. Panels show distribution patterns of rhodamine-riboflavin conjugate (A) or FTTC-transferrin (E) with DAPI (B, F), and clathrin (C, G) in the identical optical field. Panel D, H display the superimposed images from columns 1 to 3. Arrows indicate spots containing internalized rhodamine-riboflavin conjugate and clathrin signals. 134 Fig. 4.8 Colocalization of rhodamine-riboflavin with rabS protein.BeWo cells were incubated with rhodamine-riboflavin conjugate or FITC-transferrin at 4 °C for 2h. Unbound ligands were washed off, and cells were warmed to 37 °C for 10 min. After fixation and permeablization, cells were sequentially stained with anti-rabS monoclonal antibody followed by FTTC- or TRTTC-labeled secondary antibody and DAPI nuclear stain. Panels show distribution patterns of rhodamine-riboflavin conjugate (A) or FITC- transferrin (E) with DAPI (B, F), and rabS (C, G) in the identical optical field. Panel D, H, L display the superimposed images from columns I to 3. Double immunostaining reveals the extensive colocalization of clathrin (I) and anti-rab S (K). Arrows indicate spots containing internalized rhodamine-riboflavin conjugate or FITC-transferrin and rabS signals. 135 Fig. 4.9 Colocalization of rhodamine-riboflavin with transferrin (Tf).FTFC-Tf and rhodamine-riboflavin conjugate were bound to BeWo cells at 4 °C for 2h. Unbound ligands were washed off, and cells were warmed to 37 °C for 15 min. Cells were then fixed immediately, incubated with nuclear DAPI stain and processed for fluorescent microscopic analysis. Panels show distribution of rhodamine-riboflavin conjugate (A), DAPI nuclear stain (B), and FTTC-Tf (C) in the identical optical field. Panel D displays the superimposed images from panels A to C. Panel D displays the superimposed images from panels A to C. Arrows indicate spots containing both internalized rhodamine- riboflavin and FITC-transferrin. 136 Fig. 4.10 Colocalization of rhodamine-riboflavin with LAMP-1 proteins.BeWo cells were incubated with rhodamine-riboflavin conjugate or FTTC-transferrin at 4 °C for 2h. Unbound ligands were washed off, and cells were warmed to 37 °C for 30 min. After fixation and permeablization, cells were sequentially stained with anti-LAMP-i proteins monoclonal antibody followed by FTTC- or TRTTC- labeled secondary antibody. Panels show distribution patterns of rhodamine-riboflavin conjugate (A, D), FTTC-transferrin (G), with LAMP-1 proteins (B, E, H) in the identical optical field. Panel C, F, I display the superimposed images from columns 1 to 2. Solid arrows (-+) and lined arrows ( ^ ) indicate the colocalization spots and the separated stains, respectively. 137 REFERENCE Adams R (1942) Organic reactions. John Wiley & Sons,New York. Andersson H, Baechi T, Hoechi M and Richter C (1998) Autofluorescence of living cells. J Microsc 191:1-7. Betz AL, Ren XD, Ennis SR and Hultquist DE (1994) Riboflavin reduces edema in focal cerebral ischemia.Acta Neurochir Suppl 60:314-317. Bucci C, Thomsen P, Nicoziani P, McCarthy J and van Deurs B (2(XX)) Rab7: a key to lysosome biogenesis. Mol Biol Cell 11:467-480. Cemeus DP and van der Ende A (1991) Apical and basolateral transferrin receptors in polarized BeWo cells recycle through separate endosomes. J Cell Biol 114:1149-1158. Daly JW, Shi D, Padgett WL, Ji X-D and Jacobson KA (1997) Riboflavin: Inhibitory effects on receptors, G-proteins, and adenylate cyclase. Drug Dev Res 42:98-108. Hirst J and Robinson MS (1998) Clathrin and adaptors. Biochim Biopliys Acta 1404:173 193. Holladay SR, Yang Z, Kennedy MD, Leamon CP, Lee RJ, Jayamani M, Mason T and Low PS (1999) Riboflavin-mediated delivery of a macromolecule into cultured human cells. Biochim Biophys Acta 1426:195-204. Huang SN and Swaan PW (2000) Involvement of a receptor-mediated component in cellular translocation of riboflavin. J Pharmacol Exp Ther 294:117-125. Huang SN and Swaan PW (2001) Riboflavin uptake in human trophoblast-derived bewo cell monolayers: cellular translocation and regulatory mechanisms. J Pharmacol Exp TTter 298:264-271. Hultquist DE, Xu F, Quandt KS, Shlafer M, Mack CP, Till GO, Seekamp A, Betz AL and Ennis SR (1993) Evidence that NADPH-dependent methemoglobin reductase and administered riboflavin protect tissues from oxidative injury. A m J Hematol 42:13-18. McCormick DB and Zhang Z (1993) Cellular assimilation of water-soluble vitamins in the mammal: riboflavin, B6, biotin, and C. Proc Soc Exp Biol Med 202:265-270. Mellman I (1996) Endocytosis and molecular sorting. Anna Rev Cell Dev Biol 12:575- 625. Mukheijee S, Ghosh RN and MaxHeld PR (1997) Endocytosis. Physiol Rev 77:759-803. 138 Pastan IH and Willingham MC (t9S5yEndocytosis. Plenum Press,New York. Rhodaman JS and Wandinger-Ness A (2000) Rab GTPases coordinate endocytosis. J Cc//Sc/113:183-192. Rindi G and Gastaldi G (1997) Measurements and characteristics of intestinal riboflavin transport. Methods Enzymol 280:399-407. Rivlin RS (1975) Riboflavin. Plenum press,New York. Said HM and Arianas P (1991) Transport of riboflavin in human intestinal brush border membrane vesicles. Gastroenterology 100:82-88. Said HM, Ortiz A, Ma TY and McCloud E (1998) Riboflavin uptake by the human- derived liver cells Hep G2: mechanism and regulation. J Cell Physiol 176:588-594. Smart EJ, Foster DC, Ying YS, Kamen BA and Anderson RG (1994) Protein kinase C activators inhibit receptor-mediated potocytosis by preventing internalization of caveolae. J Cell Biol 124:307-313. Spinella MJ, Brigle KE, Sierra EE and Goldman ID (1995) Distinguishing between folate receptor-alpha-mediated transport and reduced folate carrier-mediated transport in L1210 leukemia cells. J Biol Client 270:7842-7849. 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CRC Press,Boca Raton. 139 CHAPTER 5 SUMMARY AND PROSPECTS Membrane transporters serve many important cellular functions including uptake of extracellular nutrients, regulation of electrolytes, and transduction of intercellular signals. Since the last two decades, it is recognized that many membrane transporters not only transfer nutrients, but also control absorption and elimination of drugs (Amidon and Sadee, 1999). For pharmaceutical scientists, the highly efficient membrane transporters further provide potential targets for drug permeation enhancement. In most cases, a prodrug approach is applied by piggy-backing the drug molecules onto naturally occurring substrates of the membrane transporters while retaining the structural requirements of the transporter systems. Through the specific interaction between ligands and transporters, biopharmaceuticals of limited membrane permeability could, therefore, be shuttled across the epithelial barrier (Swaan et al., 1996). Among all facilitated transport processes, receptor-mediated endocytosis has gained extensive interest due to its ability to transport polar macromolecules such as DNA and polypeptide into the cell. In this thesis, the cellular translocation mechanism of riboflavin was investigated to explore its potential application as means for drug delivery. To unravel the involvement of receptor-mediated endocytosis components, a two-tiered approach was employed. Using pharmacological perturbants known to alter the intracellular endocytosis events (Chapter 2), we demonstrated that transepithelial transport of 140 ribofTavîn fn Caco-2ceir monolayers was mediated by microtubule-based movements and vesicular-sorting components. In addition, these studies revealed the existence of saturable, high-affinity riboflavin translocating systems on both sides of the small intestinal membranes with the basolateral transporters exhibiting two-fold higher affinity than its apical counterparts. These polarized riboflavin transporter systems not only display dissimilar kinetic properties but also respond significantly different towards the pharmacological intervention. To obtain the morphological evidence of an endocytosis mechanism, we chose BeWo cells as our model because they have better-defined cell morphology under microscope. In order to extricate the cellular internalization mechanism from the complex bi-directional transport processes, we focused exclusively on the apical uptake of riboflavin. Consistent with our results in Caco-2 cells, a specialized riboflavin translocating system was identified on the microvillous membrane of BeWo cells (Chapter 3). The putative placental riboflavin transporter exhibits high affinity towards riboflavin, with a K, value comparable with most membrane receptors. Riboflavin structural analogs studies further indicated that the isoalloxazine ring has a more essential role in ligand-transporter interactions than the D-ribose side chain. Contrary to most mammalian membrane transporters as sodium- or proton- symporters, riboflavin uptake in BeWo cells is not coupled to the electrochemical gradients of monovalent cations, but is partly associated with a DIDS-sensitive chloride conductance. The partial chloride dependency of riboflavin uptake, though awaits further investigations, is particularly interesting. It has been known that chloride is required as a co-ion for the vacuolar H^-ATPase during acidification of intracellular vesicles (Pastan 141 andWniîngfiam, 1985). Studies have shown that reptacement of extracellular chtoride with r, SChT or DIDS treatment results in inhibition of transferrin endocytosis (Bowen and Morgan, 1988). Moreover, mutation of a renal voltage-gated chloride channel (CIC-5) was identified as the major etiology for the impaired receptor-mediated endocytosis in the proximal tubular cells in Dent’s disease (George, 1998; Sasaki et al., 2001). Future studies with voltage-dependent chloride channel inhibitors and vacuolar fT-ATPase inhibitors such as bafilomycin A| will be essential to unravel the role of chloride in the overall riboflavin uptake. Nevertheless, the chloride-mediated riboflavin uptake is physiologically significant. It not only represents a unique biochemical feature of riboflavin internalization, but also provides as a potential criterion for future screening and isolation of the riboflavin transporters. To further delineate the intracellular components and events involved in the internalization of riboflavin, we synthesized a rhodamine-riboflavin conjugate to monitor its movement via fluorescent microscopy (Chapter 4). Based on our knowledge derived from the structural analog studies in BeWo cells, rhodamine was specifically attached to the ribose moiety of riboflavin without compromising the isoalloxazine interaction between riboflavin and its transporter. Mutual inhibition between rhodamine-riboflavin conjugate and riboflavin demonstrated that the conjugate retains its high affinity towards the riboflavin transporters and was internalized via a riboflavin-specific process. Using cellular organelle probes and markers for endocytic compartments, our studies clearly showed the involvement of a classical clathrin-mediated endocytosis pathway in the cellular trafficking of the rhodamine-riboflavin conjugate. 142 Combined, it was shown hr this thesis that riboflavin is, in part, internalized by a receptor-mediated endocytosis mechanism. This was evidenced by: 1) the existence of high affinity riboflavin transporters on both sides of the cell membrane, 2) the microtubule-dependent vesicular sorting of riboflavin, 3) the subcellular localization of rhodamine-riboflavin conjugate in the clathrin-mediated endosomal compartments. In spite of all previous knowledge and these promising findings, much remains to be understood in the overall cellular translocation of riboflavin. The present morphological studies only defined the receptor-mediated endocytosis components involved in the apical internalization of riboflavin. Future studies that examine other endosomal sorting routes of riboflavin such as transcytosis will be essential to integrate the findings in our uptake and transport studies. Of equal importance will be the elucidation of basolateral riboflavin translocating machineries that act in release of transported riboflavin and/or in regulation of circulatory riboflavin in the plasma. Importantly, it should be emphasized that although current studies identify the involved endocytosis processes, it does not rule out the existence of membrane riboflavin carrier protein(s). Due to the pleiotropic nature of reagents and biochemical criteria utilized to characterize riboflavin uptake/transport, it has not been possible to selectively manipulate the cellular events of interest without affect other cellular functions. Ultimate proof for a definitive mechanism will require identifîcation of the molecular identity of the putative riboflavin transporterfs). Unfortunately, compared to other membrane transporters isolated thus far, molecular cloning of riboflavin transporter(s) is rather challenging. Despite its indispensable role in cellular functions, currently no disease(s) was directly linked to malabsorption of riboflavin in humans. Therefore, isolation of 143 membrane transporters couîd onfy refy on expressfonaT ctonmg approaches. To facilitate the isolation of riboflavin transporters we have screened a variety of cell lines ranging from enterocytes to ovarian cells for enhanced riboflavin uptake. Among all the examined cells, the placental trophoblasts exhibit the highest riboflavin absorption capacity. Key studies in the future will be to identify the respective riboflavin transporter gene(s) from the cDNA libraries constructed from the placental trophoblasts. Evaluation of the pharmaceutical relevance of riboflavin transport systems awaits a deeper understanding of both the transporters and its exact translocation mechanisms. Specifically, several major questions have yet to be answered to determine the drug delivery applicability of receptor-mediated endocytosis processes of riboflavin. First and foremost, what is the size restriction of this transporter system? Our current results indicate that through a rational conjugation strategy, riboflavin transporters can accommodate molecules at least twice as big as its natural ligands (Mw. riboflavin: 376.4, rhodamine-riboflavin: 803.8). It will be of interest to assess if the system could tolerate drug adducts with macromolecular size. On the other hand, attachment of molecules onto the ribityl group although preserves the ligand specificity and affinity towards the transporters, it limits the numbers of conjugatable sites to one per riboflavin adduct. Furthermore, late endosomal sorting of riboflavin conjugates suggests a potential routing of drug adducts into the lysosomes. Therefore, if it is possible to demonstrate that riboflavin translocating system could transport macromolecular chimeras, future studies to conjugate riboflavin to drug-encapsulating vehicles such as nanoparticles or liposomes would be necessary to increase both the cellular stability of drugs and the overall capacity of delivery system. 144 In concFusion, the findings fn this thesis fdehtifîed the existence o f membrane riboflavin transporters in the human enterocytes and placental trophoblasts. 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