Interaction of Phthalates and Phenoxy Acid Herbicide Environmental

Article pubs.acs.org/crt

Interaction of Phthalates and Phenoxy Acid Herbicide Environmental Pollutants with Intestinal Intracellular Lipid Binding Proteins Vincenzo Carbone† and Tony Velkov*,‡ †

Animal Nutrition and Health, AgResearch Limited, Grasslands Research Centre, Tennent Drive, Private Bag 11008, Palmerston North 4442, New Zealand ‡ Drug Delivery, Disposition and Dynamics, Monash Institute of Pharmaceutical Sciences, Monash University, 381 Royal Parade, Parkville 3052, Victoria, Australia S Supporting Information *

ABSTRACT: Transcellular diffusion across the columnar absorptive epithelial cells (enterocytes) of the small intestine is a major route of absorption for phthalate and phenoxy acid herbicide environmental pollutants that have been associated with adverse human health effects. The biochemical mechanisms responsible for the transport of these pollutants across the enterocyte, however, remain poorly characterized. In the present study, we have shown that the innate intestinal intracellular lipid binding proteins (iLBPs), namely, intestinal (I) and liver (L)-fatty acid binding proteins (FABP) bind to phthalate and phenoxy acid herbicides. The relative affinities of the compounds were determined by fluorescence competition assays, and a 3D-QSAR model was established for L-FABP. Structural information obtained from NMR chemical shift perturbation and molecular docking experiments defined the binding sites. Differential scanning calorimetry and proteolysis experiments revealed that the binding of these compounds produces stabilizing conformational changes in the structure of I-FABP. In summary, the presented biophysical data suggests that the binding of phthalate and phenoxy acid herbicides to intestinal iLBPs may increase the cytosolic solubility of these compounds and thereby may facilitate their transport from the intestinal lumen across the enterocyte to sites of distribution and metabolism.

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INTRODUCTION Phthalate plasticizers and phenoxy acid herbicides are widespread environmental pollutants.1−3 Phthalates are found in most plastic consumer items, and it is estimated that the mean exposures are in the order 0.01−0.05 μg kg−1 d−1 for adults and higher (∼0.07 μg kg−1 d−1) for children.4 The phenoxy acid herbicides are the most used class worldwide for the control of broad leaf weeds in crops.5 In modern day life, humans ingest phenoxy acid herbicide residues residing on the skins of fruit, on grains, and vegetables.5 Meat, fish, and dairy products also contain significant amounts of these substances.5 The widespread long-term use of phthalates and phenoxy acid herbicides and the high potential for water contamination have highlighted concerns about the long-term adverse effects on human health and the aquatic ecosystem.5 Acute exposure to phenoxy acid herbicides produces moderate toxicity in mammals, and chronic exposure has been associated with soft tissue sarcomas, endocrine effects, and severe developmental effects that include abnormal fetal skeletal development and increased fetal mortality.5−9 Similarly, chronic exposure to phthalates has been correlated with a long list of detrimental human effects such as endocrine disruption, allergies, childhood obesity, reproductive effects (fetal malformations and increased mortality), and reduced male fertility.2,3,10 Phthalates and phenoxy acid herbicides are well absorbed from the gastrointestinal tract.11−15 However, to date little is known about the molecular © XXXX American Chemical Society

mechanisms that mediate the intestinal absorption of these ubiquitous environmental toxins. Phthalates and phenoxy acid herbicides have been shown to act as potent peroxisome prolifereators and peroxisome prolifereator activated receptor (PPAR) gene activators, which may have important implications for their developmental and metabolic toxicity.16−25 PPARs are ligand-activated transcription factors that are responsible for controlling the expression of genes involved in fatty acid metabolism and glucose homeostasis.26 Early reports indicated I- and L-FABP isolated from rat intestinal tissue and purified bound to phthalates following the feeding of rats with these compounds.27 Feeding phthalates to rats increases L-FABP protein levels in the liver.28,29 Moreover, the treatment with phthalates was shown to increase the expression levels of heart FABP and all three PPAR subtypes in cultured rat placental HRP-1 trophoblast cells.30 Together, these findings are coincident with the role of FABPs as intracellular shuttles for lipophilic ligands to the nucleus, where the ligand is released to a cognate PPAR, thereby affecting the transcriptional regulation of metabolic enzymes and transporters that reciprocally target the activating ligand.31−34 This opens up the possibility that FABPs may also be involved in the binding Received: May 7, 2013

A

dx.doi.org/10.1021/tx400170t | Chem. Res. Toxicol. XXXX, XXX, XXX−XXX

Chemical Research in Toxicology

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were performed as previously described using GraphPad Prism V5.0 software (GraphPad software, San Diego, CA, USA).47 NMR 1HN−15N Backbone Amide Chemical Shift Mapping. For NMR experiments, 600 μL samples of 50 μM 15N-protein were prepared in 95% H2O/10% D2O in buffer C (20 mM MES, pH 5.5; 50 mM NaCl). Compounds were titrated into the protein solution to final concentrations of 0−2.0 mM from a DMSO stock solution, with a final DMSO concentration of less than 2% (v/v). 2D 1H−15N HSQC spectra were acquired on a Varian ANOVA 600 MHz spectrometer operating at 20 °C. Spectra were processed with the software package NMRPipe and assigned using the program SPARKY.48 The combined 1H and 15N backbone amide nuclei chemical-shift changes between apo- and holoprotein assignments were calculated using the square root of the sum of the weighted squares of the 1HN and 15N backbone amide chemical shift values (eq 1):49,50

and trafficking of phthalates and phenoxy acids herbicides to PPARs. In enterocytes, I- and L-FAPB are the most abundant intracellular proteins and may constitute as much as ∼3−6% of the total cytosolic protein.35−37 In the proximal portion of the small intestine, both I- and L-FABP are expressed at high levels.35−37 In intestinal tissues, all three PPAR subtypes are present; the PPARα and PPARδ subtypes are predominantly expressed and to a lesser extent PPARγ.38−40 FABPs display a common tertiary structure, their consensus topology consists of two β-sheets, each composed of five antiparallel β-strands capped by a helix-turn-helix motif.41,42 The 10 antiparallel β-strands are organized into a β-barrel that encloses a large solvated cavity. FABPs function by binding their ligands within a water filled cavity.42,43 Available evidence suggests that the α-helical region acts as a dynamic portal, which opens to allow ligand entry.44 In this study, we have characterized the binding of a series of phthalate and phenoxy acid herbicides to both intestinal FABPs (Supporting Information, Figure S1). The presented data give substance to the potential role of the intestinal FABPs as cytosolic transporters for these major pollutants within the enterocyte to intracellular sites of PPAR mediated transcriptional regulation and detoxification pathways.

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Δδcomb =

(ωHNΔδ1H)2 + (ω NΔδ15 N)2

(1)

where Δδ1HN and Δδ15N denote the 1HN and 15N backbone amide chemical shift change between the apo- and holo-protein for a particular residue. ωi denotes the weight factor of the nucleus, which accounts for differences in sensitivity of the 1HN and 15N ωNH = 1.0; ωNH = 0.154.51 Weight factors are determined from the ratio of the average standard deviations of the chemical shifts for a given nucleus type observed for the 20 proteogenic amino acids using the BioMagResBank chemical shift database.51 The Δδcomb for each titration was normalized to the maximum Δδcomb for the given data set. Residues that displayed significant chemical shift perturbations were mapped onto the respective crystal structures of human L-FABP (PDB code: 2F73) or human IFABP (PDB code: 3AKM) to visualize the movement of backbone amides. Limited Proteolysis of I-FABP. The limited proteolysis of I-FABP (50 μM) with sequencing grade trypsin was carried out in buffer D (20 mM Tris-HCl, pH 8.0; 50 mM NaCl; and 2 mM CaCl2) for 1 h at 20 °C. All digestions were performed at a protein/protease ratio of 20:1 (w/w). Apo- or holo-fenoprop (0, 5, 10, 25, and 50 μM final fenoprop concentration) protein samples were equilibrated with the fenoprop at 20 °C for 30 min before protease was added. Digestion reactions were stopped by the addition of 5 μL of 50 mM phenylmethanesulfonyl fluoride (PMSF) followed by one volume of SDS−PAGE sample buffer (12.5% 0.5 M Tris-HCl pH 6.8; 0.005% bromophenol blue; 10% SDS; 10% glycerol; and 2% β-mercaptoethanol) and heated for 2 min at 100 °C. Samples were resolved on 4% stacking, 20% resolving polyacrylamide gels at 4 °C at a constant voltage of 80 V, using the Laemmli buffer system. Gels were stained with Coomassie Blue G-250 and destained with 50% methanol/10% acetic acid (v/v) solution. Gels were dried between cellulose sheets and scanned at 1200 dpi. Pulse Proteolysis of Ligand−I-FABP Complexes in Urea. Pulse proteolysis of drug−I-FABP complexes was performed as previously described with minor modifications to the protocol.52,53 To monitor IFABP stability in the presence of fenoprop, the protein was equilibrated with fenoprop for 1 h at 25 °C, before the addition of urea to the reaction mixture. Reaction mixtures consisted of I-FABP (30 μM) with 0.5 mM fenoprop in buffer E (20 mM Tris-HCl pH 8.0; 50 mM NaCl; and 10 mM CaCl2) and urea to a final concentration of 0−8 M. To establish equilibrium between the folded and unfolded state, reaction mixtures in urea were incubated for 24 h prior to proteolysis. Proteolyis was initiated by the addition of thermolysin to a final concentration of 0.2 mg/mL, and the mixture was incubated at 25 °C for 1 min. Proteolysis was quenched by the addition of 3 mM EDTA (pH 8) and the addition of one volume of SDS−PAGE sample buffer and heated for 2 min at 100 °C. Samples were resolved on 4% stacking, 15% resolving polyacrylamide gels at 4 °C at a constant voltage of 80 V, using the Laemmli buffer system. Gels were stained with Coomassie Blue G-250 and destained with solution containing 50% methanol and 10% acetic acid (v/v). Following drying, the protein bands were quantified densitometrically using LabImage 1D gel analysis software, V3.4 (Kapelan GmbH, www.kapelan-bioimaging.com). The fraction of folded protein (Ffold = I/I0) was determined from the intensity of the I-FABP protein band in the presence of urea (I) divided by the band intensity of I-FABP

EXPERIMENTAL PROCEDURES

Materials. Isopropyl β-D-thiogalactopyranoside (IPTG) was purchased from BioVectra (Prince Edward Island, Canada). Sequencinggrade trypsin was purchased from (Promega, NSW, Australia). Phthalate and phenoxy acid compounds were obtained from SigmaAldrich (Sydney, NSW, Australia). Escherichia coli strain BL21 Codon Plus (DE3)-RIL was purchased from Stratagene (La Jolla, CA, USA). 15 NH4Cl was purchased from Cambridge Isotopes (Melbourne, VIC, Australia). All other reagents were of the highest purity commercially available. Protein Expression and Purification. The DNA of human I- and L-FABP were synthetically generated (GeneScript, NJ, USA) and ligated into the pET45b expression vector. The expression plasmid for human L-FABP is available from the Plasmid Repository (http:// plasmid.med.harvard.edu/PLASMID/) under the plasmid identification code HsCD00073511. Following IPTG induction at a cell density of 0.6, recombinant proteins were expressed for 6 h and purified from E. coli BL21 Codon Plus (DE3)-RIL cells. 15N-labeled proteins for NMR experiments were produced by overexpression in M9 minimal media containing 15NH4Cl using the protocol of Marley et al.45 The I- and LFABP were engineered with N-terminal [His]6 affinity tags and were separated from the bulk contaminants in the soluble cell fraction by Ni2+-based immobilized metal ion affinity chromatography (IMAC) on a Ni2+ Sepharose 5 mL HisTrapHP chromatography column (GE Health Care, Sydney, N.S.W, Australia, Cat#17−5248−02). Proteins were resolved using a step gradient of 0−300 mM imidazole in buffer A (50 mM Tris-HCl, pH 8.0; 500 mM NaCl; 0.5 mM ethylenediaminetetraacetic acid (EDTA); 1 mM dithiothreitol (DTT); 5% (v/v) glycerol) at a flow rate of 5 mL/min (4 column volumes (CV) wash-out unbound sample; 0−30% imidazole over 5 CV; held 30% for 2 CV; 30−100% imidazole over 5 CV; and held 100% for 3 CV). Delipidation and further purification was achieved by hydrophobic interaction chromatography on a Phenyl HP 16/10 column (GE Health Care, Sydney, N.S.W, Australia, Cat# 17-1085-01) as previously described.46 The final purity of the proteins was ascertained by SDS− PAGE (silver staining) and in all cases was >98%. Steady-State Fluorescence Binding Assays. Fluorometric ligand binding affinity measurements were performed under steady-state conditions on a Cary Eclipse fluorescence spectrophotometer in buffer B (20 mM Tris-HCl, pH8; 50 mM NaCl) at 20 °C (Varian, Mulgrave, Victoria, Australia). Fluorometric titrations of ANS into I- and L-FABP were performed as previously described.47 All data modeling operations B



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Table 1. Binding Affinities of Phthalate and Phenoxy Acid Herbicides for Intestinal FABPsa

a

Abbreviations: a, determined by intrinsic tryptophan fluorescence assay; nb, no binding; nt, not tested. Differential Scanning Calorimetry (DSC). DSC measurements were performed on a N-DSC II calorimeter (Calorimetry Sciences Corporation, UT, U.S.A) at a heating rate of 0.5 °C/min. For the preparation of 250 μM protein sample solutions, I-FABP were extensively buffer exchanged into buffer F (20 mM Tris-HCl at pH 8.0) by ultrafiltration using Amicon Ultra 3K centrifugal concentrators and degassed prior to filling the calorimeter cells. The filtrate from the sample preparation was used as a reference buffer for the DSC measurements. The thermal transition midpoint temperatures (Tm) were calculated using CpCalc software (Calorimetry Sciences Corporation. UT. U.S.A). Model Preparation and Molecular Docking. To produce a model of inhibitor binding, compounds were docked into the ligand cavity of human I-FABP (PDB code: 3AKM) and human L-FABP (PDB code: 2F73).54 Molecular docking experiments were carried out using the program GOLD (Genetic Optimization for Ligand Docking), version 5.1 and favoring the Piecewise Linear Potential (CHEMPLP) and ChemScore rescore Fitness function using default parameters.55 The binding sites for I-FABP and L-FABP were defined as residues and waters (where applicable) that fell within 6 Å of the fatty acid constituents and the conserved residues Arg106 and Trp82 for I-FABP and Arg122 for L-FABP. The generated binding poses were inspected, and conformations were chosen for further analysis taking into account their ranking and interactions with the cavity site residues. Additional molecular visualizations and figures were prepared using the software package PYMOL (Delano Scientific, San Carlos, CA, USA).

in absence urea (I0). The denaturant concentration at which half of the protein is unfolded (Cm) and the m value (m = ΔGunf°/[denaturant]1/2, representing the denaturant dependence of ΔGunf°) were determined by fitting Ffold to eq 2:52,53

Ffold = F0(1/1 + exp(−ΔGunf °/RT ))

(2)

where F0 represents the fraction of folded protein in the absence of urea (F0 = 1.0). The global stability of the protein is ΔGunf° = −m(Cm − [urea]); R is the gas constant (1.99 cal/(mol K)); and T is the reaction temperature, 25 °C, in Kelvin (298.15 K). Because m values determined by pulse proteolysis are not entirely reliable for the calculation of ΔGunf° (due to insufficient data in the transition zone),52,53 an m value for IFABP was estimated using a statistical method that estimates m values from the molecular size of the protein.52,53 For urea denaturation, the m value is estimated by multiplying the number of residues in the protein by a value of −0.013 kcal mol−1, and the m value of I-FABP (131 residues) was estimated to be −1.7 kcal/(mol.M). ΔGunf°, the Gibbs free energy of global unfolding which represents the difference in stability between the unfolded and native state was calculated from ΔGunf° = −m·Cm. To ensure that proteolysis of the folded protein during the 1 min pulse is negligible, a control experiment was performed where the protein is digested near the Cm and the proteolytic rate is measured. Proteolysis of I-FABP in 3.5 M urea was monitored for 10 min. Changes in the intensity of the 15 kDa I-FABP band corresponding to the intact protein were plotted as a function of time and fitted to a first-order rate equation to determine the observed proteolytic rate constant (kobs).52,53 C



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Figure 1. 1HN and 15N backbone amide chemical shift perturbations on FABPs produced by phthalate and phenoxy acid binding. (A) L-FABP binding to fenoprop. (B) I-FABP binding to MEHP. Left panels: 1H−15N HSQC spectrum of the apo-FABP (red) overlaid on top of the spectrum of holo-FABP (blue). Right panels: a plot of the 1HN and 15N backbone amide chemical shift perturbations upon ligand binding versus residue number. Bottom panels: perturbed residues (>0.5 ppm perturbation) mapped onto the three-dimensional structure of each FABP. The inset shows the chemical structure of the docked ligand. 3D-QSAR Analysis. Phase from Schrö dinger was used for

FABP, which provided the best spread of activities over the different

pharmacophore elucidation and 3D-QSAR model building for L-

classes of compounds tested.56 Ki values were converted to −log10 Ki, D



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Figure 2. Binding to the phenoxy acid herbicide fenoprop protects I-FABP from proteolysis and heat denaturation. (A) DSC thermograms of I-FABP in the presence of increasing concentrations of fenoprop. The concentration of fenoprop and Tm is indicated above each thermogram. The scan rate was 0.5 °C/min. (B) Binding to fenoprop protects I-FABP from limited proteolysis. I-FABP was pre-equilibrated with increasing concentrations (0, 5, 10, 25, and 50 μM) of fenoprop and then digested with trypsin. The resultant fragments were resolved on 20% polyacrylamide gels and visualized by Coomassie Blue G250 staining. (C) Global stability of apo-I-FABP (▼) and holo-I-FABP in complex with fenoprop (○) measured by pulse proteolysis. Top panel: Ffold plotted as a function of the urea concentration. The solid line represents the nonlinear least-squares fit of Ffold to eq 2 from which Cm values were determined. Bottom panels: SDS−PAGE gel profiles of 1 min proteolytic pulse reactions of apo- and holo-I-FABP complexes in the presence of increasing concentrations of urea. The protein band corresponding to intact I-FABP and the protease thermolysin are indicated. The intensity of the IFABP band was quantified desitometrically and used to calculate the fraction of folded protein (Ffold). while the nonbinders were omitted entirely from the study. To elevate compounds with features that are essential for high-affinity binding, active ligands were defined as those with pKi > −0.8, while inactive ligands were defined as those with pKi < −0.8. In addition, conditions were relaxed so that a common pharmacophore needed to only match a subset of the actives (4 of 7 compounds). Conformers were generated for each ligand using an OPLS_2005 force field and an implicit distancedependent dielectric solvent model at a cutoff root-mean-square deviation (RMSD) of 1 Å and a maximum energy difference of 30 kcal/mol.

herbicides displayed a moderate affinity (Table 1). Notably, only the mono-substituted phthalates mono(2-ethylhexyl) phthalate (MEHP) and mono(1-methylheptyl)phthalate (MOP) bound to the FABPs, whereas the disubstituted phthalate di(2ethylhexyl) phthalate (DEHP) did not bind, suggesting the carboxylate functionality is indispensible for binding. A closer examination of the binding affinity data for the phenoxy acids revealed a consistent structure−activity relationship (SAR) trend for both I- and L-FABP, albeit L-FABP consistently displayed a higher affinity. Like their native fatty acid substrates, the phenoxy acids consist of a terminal carboxylate with a hydrophobic “tail” in the form of a benzene ring. Conformational Changes in Intestinal FABPs Induced by Binding to Phthalate and Phenoxy Acid Herbicide Compounds Monitored by 1H and 15N Backbone Amide Chemical Shift Mapping. Local conformational changes induced by ligand binding can be monitored by 1HN and 15N backbone amide chemical shift changes that are related to the change of the dihedral ϕ, ψ-angles.50 Changes in chemical shift were followed by recording the 1H−15N-HSQC spectra of each FABP in the presence of a high affinity phthalate or phenoxy acid

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RESULTS Examination of the Binding Affinity of Phthalate and Phenoxy Acid Herbicide Compounds for Intestinal FABPs by Fluorescence Displacement Assays. The binding dissociation constants (Ki) for phthalate and phenoxy acid herbicide compounds were measured fluorometrically by monitoring the competitive displacement of the fluorescent probe ANS from the ligand binding cavities of I- and L-FABP (Table 1). Binding isotherms conformed well to a one site binding model.47,57 Overall, the phthalates displayed the highest binding affinity for I- and L-FABP, whereas the phenoxy acid E



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herbicide compound (Figure 1). Mapping of the chemical shift perturbations between the apo- and holo-FABP complexes onto their three-dimensional structures revealed that the most significant perturbations were concentrated within the binding cavity or the portal region that mediates ligand entry/exit from the β-barrel cavity (Figure 1). Binding to Fenoprop Protects I-FABP from Proteolysis and Heat Denaturation. DSC thermal denaturation measurements revealed that complexation with fenoprop increased the thermal transition midpoint temperature of I-FABP from Tm = 62.7 to Tm = 67.5 (Figure 2A), which would indicate that fenoprop binding stabilizes the I-FABP structure. Similarly, the thermal transition midpoint temperature of L-FABP increased from Tm = 80.2 to Tm = 86.5, which is in line with our previously reported thermal shift results for human L-FABP binding to PPARα activator ligands.58 Figure 2B shows the proteolytic peptide pattern that evolves from the digestion of I-FABP with trypsin (Arg-C; Lys-C) in the apo- and holo-fenoprop bound forms. The apo-form was significantly more susceptible to proteolysis than the holofenoprop form. Control experiments with E.coli lac repressor, a protein that does not possess an affinity for fenoprop showed no protection against cleavage, thus ruling out the possibility that the protection observed with I-FABP is due to an inhibitory effect of fenoprop on trypsin (data not shown). Pulse proteolysis is designed to digest only target protein molecules that are unfolded. The time of the proteolyic pulse (1 min) is much shorter than the relaxation time between the folded and unfolded proteins in the equilibrium mixture; thus, only the unfolded protein molecules are digested. Because excess protease is present in the equilibrium mixture, digestion of the unfolded proteins occurs rapidly, whereas proteolysis of the folded species is slow and negligible within the time frame of the pulse. Thermolysin is the most suitable proteolytic enzyme for this assay due to its broad specificity and ability to function in the presence of high concentrations of urea, thereby ensuring complete digestion of unfolded protein at most concentrations of the denaturant. An important criteria for the success of this assay is that the folded protein is not significantly digested within the pulse period, and this can occur if the intrinsically unstructured regions are present in the native structure. Although, I-FABP displays intrinsically unstructured regions within the portal, there was no apparent digestion in the absence of denaturant within the 1 min pulse period employed for these experiments.59 Thus, in the presence of urea, folded I-FABP in the equilibrium mixture should remain intact within the 1 min pulse, providing a reliable estimate of the fraction of folded protein (Ffold). The amount of folded protein that remained after the 1 min proteolytic pulse was determined by densitometric quantification of SDS−PAGE protein bands and was used to calculate Ffold (Figure 2C). As the urea concentration is increased, the two-state equilibrium is shifted toward the unfolded, proteolytically susceptible form. In the apo-form, I-FABP remained resistant to proteolysis at urea concentrations up to 2 M, rapidly decreased in the 3−5 M concentration range, and completely disappeared at 6−7 M urea, consistent with the cooperative transition of protein unfolding. The denaturant concentration at which Ffold is 0.5 (Cm) was determined by fitting Ffold at each urea concentration to a two state equilibrium unfolding model (Figure 2C, top panel; Table 2). The midpoint of transition (Cm) is a function of the stability and the m value of the protein. Therefore, the global stability of the protein (ΔGunf°) was calculated by multiplying the Cm value determined from pulse proteolysis by the estimated m value for I-

Table 2. Stability of I-FABP Determined by Pulse Proteolysis Urea Unfolding Measurementsa

apo-I-FABP Fenoprop-IFABP a

Cm 0.5 unfold [urea] (M)

ΔG°unf (kcal/mol)

ΔΔGunf° (kcal/mol)

4.1 ± 0.2 4.9 ± 0.3

7.0 ± 0.3 8.4 ± 0.4

na 1.4

Abbreviations: na, not applicable.

FABP. Table 2 documents the stability parameters determined by pulse proteolysis for apo- and the holo-I-FABP complexes. The ΔGunf° values obtained for the apo-protein by this method were in agreement with those reported for equilibrium urea unfolding detected by spectroscopic methods.60 The calculated change in the global stability (ΔΔGunf°) in the presence of fenoprop relative to the apo-protein indicates that fenoprop binding contributes significantly to the global stability of I-FABP (Table 2). It is evident from the increase in Cm values and the apparent decrease in unfolding rate that fenoprop binding affects both the stability and kinetics of unfolding of the I-FABP structure. The docking solutions with I-FABP show that fenoprop makes a number of potentially powerful hydrophobic interactions that may bear out this stability. Molecular Docking Simulations. The NMR chemical shift perturbation experiments were complemented by molecular docking and QSAR analysis of the test compounds into the threedimensional structures of the intestinal FABPs (Figure 3). In each case, many of the residues which displayed significant

Figure 3. (A) The binding poses of phenoxy proprionic acid herbicides in L-FABP. (B) The binding poses of phenoxy proprionic acid herbicides in I-FABP. (C) Fenoprop docked into L-FABP. (D) Fenoprop docked into I-FABP. (E) Phthalates docked into L-FABP; MEHP in green and MOP in yellow. (F) Phthalates docked into IFABP, MOP in purple, and MEHP in yellow. Residues within 4 Å of the molecules are represented, with hydrogen bonds shown as dotted lines. F



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groups of MEHP and MOP fall well within 4 Å of a hydrophobic pocket of residues and side chain moieties including Leu91, Phe63, Phe48, Ile41, Phe50, Thr102, and Thr93. In I-FABP, MOP and MEHP bind in a mode/pocket similar to 11(dansylamino)undecanoic acid in 3AKM.54 As in L-FABP, the carboxylate of MEHP and MOP makes a number of hydrogen bond contacts with side chain amines of Trp82 (NE1; 2.6 Å) and Arg106 (NH1; 2.7 Å and 2.6 Å, respectively) and the side chain carboxylate of Glu51 (OE2; 2.4 and 2.7 Å, respectively). The carbonyl moiety makes hydrogen bond contact with the Gln115 (NE2; 2.7 and 2.9 Å, respectively), while the ethylhexyl and methylheptyl groups make identical interactions with aliphatic side chains as we observed in L-FABP with residues including Phe17, Leu102, Leu72, Leu78, Phe93, and Tyr117. In addition, MOP and MEHP form a π-stacking interaction with the side chain ring of Tyr70 (3.7 Å; Figure 3F). 3D-QSAR. Pharmacophore modeling of the L-FABP binders were developed to identify key features necessary for ligand potency. It is easy to observe that the most active compounds contained exclusively a hydrophobic tail and a terminal carboxylate. The generated hypotheses of phase were therefore rejected if they were unable to define the complete binding space of the selected ligands regardless of survival score. The resultant pharmacophore models generated included combinations of hydrogen bond acceptors (A), hydrophobic groups (H), the negatively charged group (N), and aromatic ring (R) and labeled as AHHR (7 hypotheses), AHNR (13 hypotheses), AHHN (98 hypotheses), and HHNR (7 hypotheses). The best hypothesis AHNR.13 was selected on the basis of overall fit with all common pharmacophores in the strongest herbicide binder−fenoprop and best validated r2 (0.81) for 11 of the 13 active herbicides (Figure 4A and B). AHNR.13 had a good survival score of (3.094), a posthoc score of (2.969), a P value of (2.659 × 10−4), and an F value of (83.1). The high affinity of fenoprop was further validated by the 3D-QSAR model (Figure 4C and D). In the context of a single high-affinity binding ligand such as

chemical shift perturbations in the NMR data were clustered around the docked ligand in the models, suggesting that the models are representative of the binding of the drugs. Across the phenoxy acid structural series, pertinent features that confer optimal binding included the length of and presence of a terminal carboxylic acid moiety and their interaction with the conserved arginine within the FABP cavity, much like the poses observed for a number of fatty acid molecules bound to FABPs.41,61 Once that pharmacophore is converted to an aliphatic ester, then binding to the FABP is abrogated (e.g., MCPP and 2,4,5-T methyl, cf. Table 1). This was also borne out in the docking simulations where there was no consistent binding poses observed with these particular compounds. Both the phenoxy propionic and phenoxy butyric acid based herbicides adopted similar orientations within the I- and L-FABP cavity (Figure 3A and B). L-FABP has hydrogen bond contacts with the side chain amines of Arg122 (NH1) and Asn111 (ND2), and the side chain hydroxyl of Thr102 (OG1), while IFABP maintains hydrogen bond contacts with the amines of Arg106 (NH1 and NH2) and Trp82 (NE1). These particular interactions are carried out almost exclusively by the carboxylate moiety of either series of herbicides. Outside of the phthalates and acifluorfen, fenoprop remains the tightest binder of L-FABP (an observation that is also attributable to I-FABP). We would therefore assume that the presence of a 5-chloro ring substituent and a 2-methyl propionic acid substituent is crucial to high binding affinity for the phenoxy series of inhibitors. In L-FABP, the methyl group of fenoprop forms a tight hydrophobic interaction in an area bordered by aliphatic residues including Ile52, Ile41, Phe50, and Ile109; however, none of the chloride moieties produce identifiable van der Waals interactions, but the benzene ring forms a tight πstacking interaction between with the side chain of Phe50 (3.6 Å; Figure 3C). The lower binding affinity of gemfibrozil is attributed to the steric bulk imposed by the 2,5-methyl substituted phenoxy terminus and pentanoic acid backbone, which promoted fewer hydrogen bond interactions with cavity residues and did not allow for the important π-stacking interaction with Phe50. The IFABP-gemfibrozil in silico screen did not produce a consistent binding model that exploited hydrogen bonding with Arg106. What was consistent (and due to the higher preponderance of polar residues in I-FABP) was the placement of the 2,5dimethylphenoxy group of gemfibrozil immediately adjacent to Arg106 in a tight hydrophobic pocket and with the benzene ring carefully π-stacking with the side chain of Tyr70. The modeled I-FABP−fenoprop complex produced hydrogen bond and van der Waals contacts in addition to those aforementioned, mediated by the ether moiety with the cavity Arg106 and the residue Glu51 (3.2 and 3.8 Å, respectively; Figure 3D). Like L-FABP, the methyl group of fenoprop is bordered predominantly by aliphatic and aromatic residues including Phe68, Tyr70, Trp82, and Phe93, and while not observed in our rigid model, we suspect that potential van der Waals contacts would be possible with the 2- and 5-chloro moieties of fenoprop and the hydroxyl groups of Tyr70 and Thr104, respectively, via the rotameric adjustments of their side chains. Phthalate Binding. Both phthalate molecules assume relatively identical orientations within L-FABP (Figure 3E). The predominantly strong interaction with the molecule is metered by a number of hydrogen bond contacts by the carboxylate and carbonyl moiety with the amine of Asn111 (ND2; 2.7 Å), the hydroxyl of Ser100 (OG; 3.0 Å), and the amine Arg122 (NH1; 3.1 Å). The ethylhexyl and methylheptyl

Figure 4. (A) Scatter plot of observed versus predicted biological activity of all compounds in the training and test set (and excluding 2 inactive molecules 2,4-D and 2,4-DP; r2 = 0.81). (B) The most active compound fenoprop overlaid with the best phase hypothesis (ANHR.13). (C) Visual representation of an atom-based 3D-QSAR model in the context of a single high-affinity ligand, where favorable hydrophobic and (D) negative and electron withdrawing groups (in blue) dominate unfavorable (red) characteristics. G



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compounds are largely ionized at the pH (7.4) used for these experiments, it would mean they would be poorly absorbed into the apical membrane of the cells. The latter scenario would also account for the unique ability of diclofop-methyl to serve as a PPAR activator, as the methyl-ester of the carboxylate is neutral, which would allow the compound to readily permeate into cellular membranes. The labile methyl ester would then become hydrolyzed to the carboxylate inside the cell, generating the active compound. Moreover, the COS-1 cell line used in one of the reports does not express FABPs, which are known to facilitate desorption of lipophilic molecules from the plasma membrane.65,68 Phthalates generally show good intestinal absorption in humans.69−71 Following oral exposure, DEHP is hydrolyzed by intestinal lipases to MEHP along with the release of the 2ethylhexonal alcoholic substituent.71−75 At low exposure levels, DEHP is mostly hydrolyzed and absorbed as MEHP,71,73,76 whereas at high concentrations the hydrolytic enzymes become saturated, and unhydrolyzed DEHP is also absorbed.71,77 It is tempting to speculate that the lower acute toxicity and lower intestinal absorption of DEHP compared to MEHP might be due to the dependency on monode-esterification of the diester and the lower affinity of intestinal transporters for DEHP such as the FABPs.75,78 Nuclear hormone receptors (NHRs) act as chemical sensors or xenosensors, which react to the presence of a ligand and increase the levels of expression of the proteins that transport and detoxify that ligand via efflux or metabolism.79 These proteins include iLBP transporters, cytochrome P450 metabolizing enzymes, and membrane efflux transporters, which physically remove toxins from the cell.79 In the case of an absorptive cell such as the enterocyte, this also reduces absorption and exposure to the rest of the body. Phthalates and phenoxy acids are potent peroxisome proliferators and PPAR activators.16−25 In addition to PPARs, phthalates have been also shown to activate the constitutive androstane receptor and pregnane X receptor.80 Since NHRs are located in the nucleus of the enterocyte, transport of these relatively insoluble lipophilic environmental pollutants from the cytoplasm to the nucleus seems to be required for activation of the PPAR transcriptional response. Initially, the intestinal FABPs were assumed to be localized to the cytoplasm for the sole function of controlling the intracellular concentration of fatty acids.43 However, in our recent reports binding to FABPs has emerged as a potential mechanism whereby a diverse range of lipophilic xenobiotics, such as orally administered drugs, can be solubilized and targeted to specific intracellular organelles and receptors in enterocytes.47,64,65 Furthermore, over a series of exemplary reports, researchers from our group and the laboratories of Schroeder and Spener have provided convincing evidence that L-FABP directly interacts with PPARα and is involved in the nucleo-cytoplasmic shuttling of activator ligands.34,58,81−87 In light of their general solubilizing function for small lipophilic molecules, broad ligand selectivity, and high tissue expression levels, the innate intestinal FABPs are obvious candidates to perform the intracellular nucleo-cytoplasmic transport role for phthalate and phenoxy acid pollutants in enterocytes. The NMR chemical shift perturbation data together with the partial proteolysis and DSC experiments suggest that the binding of these pollutants to FABPs produces a stabilizing conformational change in the portal region of the molecule. The functional importance of this conformational change has recently been demonstrated in the case of adipocyte (A)-FABP. Activator ligand binding to A-FABP induces changes

fenoprop, contouring for favorable hydrophobic and negative and electron withdrawing groups (in blue) dominated unfavorable (red) characteristics and in particular highlighted the importance of the hydrophobic methyl group and methoxy and carboxylate oxygens.

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DISCUSSION In addition to the numerous dietary lipophilic molecules, intestinal cells are challenged with a variety of lipophilic environmental xenobiotics such as phthalates and phenoxy acid herbicides that can be toxic if present at high concentrations. Accordingly, enterocytes have developed elegant biochemical signaling and metabolic pathways for the elimination of these compounds. The central hypothesis examined in this work was that intestinal FABPs bind phthalate and phenoxy acid environmental pollutants and thereby potentially facilitate their solubilization and transport across the aqueous environment of the enterocyte cytosol. A structural series of phthalate and phenoxy acid herbicides were validated as binding site ligands by a combination of NMR spectroscopy chemical shift perturbation experiments and fluorometric binding assays. Mapping of the nuclei that undergo chemical shift perturbations upon ligand titration is a sensitive indicator of the positions that are either adjacent to the binding location or at which structural changes occur upon complex formation.49,50 It is apparent that the binding of these ligands was site-specific and localized to the binding cavity and portal regions of each FABP. The binding affinities (Ki) of the test compounds measured by fluorescence competition assays revealed that the phenoxy acids displayed a considerably lower affinity for the FABPs compared to those of their native fatty acid ligands (Table 1). Importantly, the cytosolic concentration of the FABPs in the enterocyte is 0.2−0.4 mM, which is far in excess of the total fatty acid concentration leaving excess apo-FABP available for binding to lower-affinity ligands, including herbicide molecules.35−37 In contrast, the phthalates displayed a high binding affinity for both I- and LFABP (Table 1). The ability of the intestinal FABPs to bind phthalates and phenoxy acid herbicides is not surprising if one considers that these compounds bear physicochemical similarities with mammalian fatty acids in the form of a predominantly aliphatic moiety in the tail group and charged carboxylic acid headgroup. This requirement of an aliphatic tail group is also borne out of the 3D-QSAR modeling experiments suggesting that even the halides act as hydrophobic groups and not as Lewis acids in this case and suggests why in the phenoxy series of herbicides that the removal of halides of any combination (with respect to fenoprop) reduced binding affinity and which in turn matched our predicted hydrophobic interactions between the herbicide and the FABP cavity. Furthermore, phenoxy acid herbicides are structurally homologous with the classic peroxisome proliferators and hypolipidemic drugs, the fibrates, particularly clofibrate, which coincidently has demonstrated herbicidal activity.62 Similar to these compounds, the phenoxy acid herbicides act as potent peroxisomal proliferators, affecting cholesterol and triglyceride levels.20,22,63 It was reported that 2,4DP possessed a greater cholesterol lowering effect than an equimolar dose of clofibrate.22 Coincidently, we have previously reported that both I- and L-FABP avidly bind to clofibrate.47,64,65 Intriguingly, with the exception of diclofop-methyl, the phenoxy acid herbicides do not show PPAR activation in vitro within cell culture-based PPAR reporter assays.66,67 These findings suggest that the phenoxy acid herbicides may undergo conversion to an active metabolite in vivo, or more simply, because these H



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(5) Sterling, T. D., and Arundel, A. V. (1986) Health effects of phenoxy herbicides. A review. Scand. J. Work, Environ. Health 12, 161−173. (6) Manninen, A., Kangas, J., Klen, T., and Savolainen, H. (1986) Exposure of Finnish farm workers to phenoxy acid herbicides. Arch. Environ. Contam. Toxicol. 15, 107−111. (7) Green, L. M. (1987) Suicide and exposure to phenoxy acid herbicides. Scand. J. Work, Environ. Health 13, 460. (8) Hill, R. H., Jr., To, T., Holler, J. S., Fast, D. M., Smith, S. J., Needham, L. L., and Binder, S. (1989) Residues of chlorinated phenols and phenoxy acid herbicides in the urine of Arkansas children. Arch. Environ. Contam. Toxicol. 18, 469−474. (9) Johnson, C. C., Feingold, M., and Tilley, B. (1990) A meta-analysis of exposure to phenoxy acid herbicides and chlorophenols in relation to risk of soft tissue sarcoma. Int. Arch. Occup. Environ. Health 62, 513−520. (10) Hatch, E. E., Nelson, J. W., Stahlhut, R. W., and Webster, T. F. (2012) Association of endocrine disruptors and obesity: perspectives from epidemiological studies. Int. J. Androl. 33, 324−332. (11) Arnold, E. K., and Beasley, V. R. (1989) The pharmacokinetics of chlorinated phenoxy acid herbicides: a literature review. Vet. Hum. Toxicol. 31, 121−125. (12) Kohli, J. D., Khanna, R. N., Gupta, B. N., Dhar, M. M., Tandon, J. S., and Sircar, K. P. (1974) Absorption and excretion of 2,4,5trichlorophenoxy acetic acid in man. Arch. Int. Pharmacodyn. Ther. 210, 250−255. (13) Kohli, J. D., Khanna, R. N., Gupta, B. N., Dhar, M. M., Tandon, J. S., and Sircar, K. P. (1974) Absorption and excretion of 2,4dichlorophenoxyacetic acid in man. Xenobiotica 4, 97−100. (14) Wittassek, M., and Angerer, J. (2008) Phthalates: metabolism and exposure. Int. J. Androl. 31, 131−138. (15) McKee, R. H., El-Hawari, M., Stoltz, M., Pallas, F., and Lington, A. W. (2002) Absorption, disposition and metabolism of di-isononyl phthalate (DINP) in F-344 rats. J. Appl. Toxicol. 22, 293−302. (16) Kobayashi, T., Niimi, S., Kawanishi, T., Fukuoka, M., and Hayakawa, T. (2003) Changes in peroxisome proliferator-activated receptor gamma-regulated gene expression and inhibin/activinfollistatin system gene expression in rat testis after an administration of di-n-butyl phthalate. Toxicol. Lett. 138, 215−225. (17) Ren, H., Aleksunes, L. M., Wood, C., Vallanat, B., George, M. H., Klaassen, C. D., and Corton, J. C. (2010) Characterization of peroxisome proliferator-activated receptor alpha–independent effects of PPARalpha activators in the rodent liver: di-(2-ethylhexyl) phthalate also activates the constitutive-activated receptor. Toxicol. Sci. 113, 45− 59. (18) Valles, E. G., Laughter, A. R., Dunn, C. S., Cannelle, S., Swanson, C. L., Cattley, R. C., and Corton, J. C. (2003) Role of the peroxisome proliferator-activated receptor alpha in responses to diisononyl phthalate. Toxicology 191, 211−225. (19) Hurst, C. H., and Waxman, D. J. (2003) Activation of PPARalpha and PPARgamma by environmental phthalate monoesters. Toxicol. Sci. 74, 297−308. (20) Vainio, H., Nickels, J., and Linnainmaa, K. (1982) Phenoxy acid herbicides cause peroxisome proliferation in Chinese hamsters. Scand. J. Work, Environ. Health 8, 70−73. (21) Lovekamp-Swan, T., and Davis, B. J. (2003) Mechanisms of phthalate ester toxicity in the female reproductive system. Environ. Health. Perspect. 111, 139−145. (22) Vainio, H., Linnainmaa, K., Kahonen, M., Nickels, J., Hietanen, E., Marniemi, J., and Peltonen, P. (1983) Hypolipidemia and peroxisome proliferation induced by phenoxyacetic acid herbicides in rats. Biochem. Pharmacol. 32, 2775−2779. (23) Lapinskas, P. J., Brown, S., Leesnitzer, L. M., Blanchard, S., Swanson, C., Cattley, R. C., and Corton, J. C. (2005) Role of PPARalpha in mediating the effects of phthalates and metabolites in the liver. Toxicology 207, 149−163. (24) Corton, J. C., and Lapinskas, P. J. (2005) Peroxisome proliferatoractivated receptors: mediators of phthalate ester-induced effects in the male reproductive tract? Toxicol. Sci. 83, 4−17. (25) Lampen, A., Zimnik, S., and Nau, H. (2003) Teratogenic phthalate esters and metabolites activate the nuclear receptors PPARs

within the portal region that stabilizes a conformational nuclear localization signal (NLS) situated in the αII helix consisting of Lys21, Arg30, and Lys31.88 This in turn leads to nuclear import of the A-FABP, where it can release its ligand cargo to its cognate NHR, PPARγ.31,33 Coincidently, the NLS motif is present in the sequence of I-FABP suggesting that portal stabilizing ligands such as phthalates and phenoxy acid pollutants may trigger the nucleo-cytoplasmic shuttling of I-FABP.58 An interesting facet of mammalian xenobiotic response pathways is the multiple layers of functional redundancy across each of the biochemical components.79 The several layers of functional overlap and redundancy provide the framework for broad specificity systems that can cope with challenges from diverse xenochemicals. Presumably, the ability of both innate intestinal FABPs to bind to these pollutants is reflective of this functional redundancy in the enterocyte and serves as a fail-safe mechanism to facilitate host protection. To the best of our knowledge, this is the first report of the specific binding interaction of human intestinal FABPs with phthalate and phenoxy acid herbicide pollutants. These data provide a detailed account of the binding determinants of these xenobiotics and provide new insight into the potential mechanism(s) by which these pollutants are absorbed and transported through the intestine. More importantly, the data suggest intestinal FABPs may be in effect part of a general cellular detoxification pathway for these ubiquitous environmental pollutants.

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ASSOCIATED CONTENT

* Supporting Information S

Chemical structures of phthalate and phenoxy acid herbicide compounds used in this study. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Phone: +61-3-9903-9539. Fax: +61-3-9903-9582. E-mail: [email protected]. Funding

T.V. is an Australian National Health and Medical Research Council Industry Career Development Research Fellow. Notes

The authors declare no competing financial interest.

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ABBREVIATIONS DEHP, di(2-ethylhexyl)phthalate; FABP, fatty acid binding protein;; iLBP, intracellular lipid binding protein;; 2D-1H−15NHSQC, two-dimensional amide proton and nitrogen heteronuclear single-quantum correlation; MEHP, mono(2ethylhexyl)phthalate; MOP, mono(1-methylheptyl)phthalate

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REFERENCES

(1) Heudorf, U., Mersch-Sundermann, V., and Angerer, J. (2007) Phthalates: toxicology and exposure. Int. J. Hyg. Environ. Health 210, 623−634. (2) Crinnion, W. J. (2010) Toxic effects of the easily avoidable phthalates and parabens. Altern. Med. Rev. 15, 190−196. (3) Meeker, J. D., Sathyanarayana, S., and Swan, S. H. (2009) Phthalates and other additives in plastics: human exposure and associated health outcomes. Philos. Trans. R. Soc., B. 364, 2097−2113. (4) Koch, H. M., and Calafat, A. M. (2009) Human body burdens of chemicals used in plastic manufacture. Philos. Trans. R. Soc., B. 364, 2063−2078. I



Article

and induce differentiation of F9 cells. Toxicol. Appl. Pharmacol. 188, 14− 23. (26) Lemberger, T., Desvergne, B., and Wahli, W. (1996) Peroxisome proliferator-activated receptors: a nuclear receptor signaling pathway in lipid physiology. Annu. Rev. Cell. Dev. Biol. 12, 335−363. (27) Kanda, T., Ono, T., Matsubara, Y., and Muto, T. (1990) Possible role of rat fatty acid-binding proteins in the intestine as carriers of phenol and phthalate derivatives. Biochem. Biophys. Res. Commun. 168, 1053− 1058. (28) Kawashima, Y., Nakagawa, S., Tachibana, Y., and Kozuka, H. (1983) Effects of peroxisome proliferators on fatty acid-binding protein in rat liver. Biochim. Biophys. Acta 754, 21−27. (29) Kawashima, Y., Nakagawa, S., and Kozuka, H. (1982) Effects of some hypolipidemic drugs and phthalic acid esters on fatty acid binding protein in rat liver. J. Pharmacobiodyn. 5, 771−779. (30) Xu, Y., Cook, T. J., and Knipp, G. T. (2005) Effects of di-(2ethylhexyl)-phthalate (DEHP) and its metabolites on fatty acid homeostasis regulating proteins in rat placental HRP-1 trophoblast cells. Toxicol. Sci. 84, 287−300. (31) Ayers, S. D., Nedrow, K. L., Gillilan, R. E., and Noy, N. (2007) Continuous nucleocytoplasmic shuttling underlies transcriptional activation of PPARgamma by FABP4. Biochemistry 46, 6744−6752. (32) Schug, T. T., Berry, D. C., Shaw, N. S., Travis, S. N., and Noy, N. (2007) Opposing effects of retinoic acid on cell growth result from alternate activation of two different nuclear receptors. Cell 129, 723− 733. (33) Tan, N. S., Shaw, N. S., Vinckenbosch, N., Liu, P., Yasmin, R., Desvergne, B., Wahli, W., and Noy, N. (2002) Selective cooperation between fatty acid binding proteins and peroxisome proliferatoractivated receptors in regulating transcription. Mol. Cell. Biol. 22, 5114− 5127. (34) Wolfrum, C., Borrmann, C. M., Borchers, T., and Spener, F. (2001) Fatty acids and hypolipidemic drugs regulate peroxisome proliferator-activated receptors alpha - and gamma-mediated gene expression via liver fatty acid binding protein: a signaling path to the nucleus. Proc. Natl. Acad. Sci. U.S.A. 98, 2323−2328. (35) Kanda, T., Fujii, H., Fujita, M., Sakai, Y., Ono, T., and Hatakeyama, K. (1995) Intestinal fatty acid binding protein is available for diagnosis of intestinal ischaemia: immunochemical analysis of two patients with ischaemic intestinal diseases. Gut 36, 788−791. (36) Lowe, J. B., Sacchettini, J. C., Laposata, M., McQuillan, J. J., and Gordon, J. I. (1987) Expression of rat intestinal fatty acid-binding protein in Escherichia coli. Purification and comparison of ligand binding characteristics with that of Escherichia coli-derived rat liver fatty acid-binding protein. J. Biol. Chem. 262, 5931−5937. (37) Poirier, H., Niot, I., Degrace, P., Monnot, M. C., Bernard, A., and Besnard, P. (1997) Fatty acid regulation of fatty acid-binding protein expression in the small intestine. Am. J. Physiol. 273, G289−295. (38) Lemberger, T., Braissant, O., Juge-Aubry, C., Keller, H., Saladin, R., Staels, B., Auwerx, J., Burger, A. G., Meier, C. A., and Wahli, W. (1996) PPAR tissue distribution and interactions with other hormonesignaling pathways. Ann. N.Y. Acad. Sci. 804, 231−251. (39) Braissant, O., Foufelle, F., Scotto, C., Dauca, M., and Wahli, W. (1996) Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR-alpha, -beta, and -gamma in the adult rat. Endocrinology 137, 354−366. (40) Escher, P., Braissant, O., Basu-Modak, S., Michalik, L., Wahli, W., and Desvergne, B. (2001) Rat PPARs: quantitative analysis in adult rat tissues and regulation in fasting and refeeding. Endocrinology 142, 4195− 4202. (41) Furuhashi, M., and Hotamisligil, G. S. (2008) Fatty acid-binding proteins: role in metabolic diseases and potential as drug targets. Nat. Rev. Drug Discovery 7, 489−503. (42) Hamilton, J. A. (2002) How fatty acids bind to proteins: the inside story from protein structures. Prostaglandins, Leukotrienes Essent. Fatty Acids 67, 65−72. (43) Schroeder, F., Jolly, C. A., Cho, T. H., and Frolov, A. (1998) Fatty acid binding protein isoforms: structure and function. Chem. Phys. Lipids. 92, 1−25.

(44) Hodsdon, M. E., and Cistola, D. P. (1997) Discrete backbone disorder in the nuclear magnetic resonance structure of apo intestinal fatty acid-binding protein: implications for the mechanism of ligand entry. Biochemistry 36, 1450−1460. (45) Marley, J., Lu, M., and Bracken, C. (2001) A method for efficient isotopic labeling of recombinant proteins. J. Biomol. NMR 20, 71−75. (46) Velkov, T., Lim, M. L., Capuano, B., and Prankerd, R. (2008) A protocol for the combined sub-fractionation and delipidation of lipid binding proteins using hydrophobic interaction chromatography. J. Chromatogr., B 867, 238−246. (47) Velkov, T., Chuang, S., Wielens, J., Sakellaris, H., Charman, W. N., Porter, C. J., and Scanlon, M. J. (2005) The interaction of lipophilic drugs with intestinal fatty acid-binding protein. J. Biol. Chem. 280, 17769−17776. (48) Delaglio, F., Grzesiek, S., Vuister, G. W., Zhu, G., Pfeifer, J., and Bax, A. (1995) NMRPipe: a multidimensional spectral processing system based on UNIX pipes. J. Biomol. NMR 6, 277−293. (49) Schumann, F. H., Riepl, H., Maurer, T., Gronwald, W., Neidig, K. P., and Kalbitzer, H. R. (2007) Combined chemical shift changes and amino acid specific chemical shift mapping of protein-protein interactions. J. Biomol. NMR 39, 275−289. (50) Shuker, S. B., Hajduk, P. J., Meadows, R. P., and Fesik, S. W. (1996) Discovering high-affinity ligands for proteins: SAR by NMR. Science 274, 1531−1534. (51) Doreleijers, J. F., Mading, S., Maziuk, D., Sojourner, K., Yin, L., Zhu, J., Markley, J. L., and Ulrich, E. L. (2003) BioMagResBank database with sets of experimental NMR constraints corresponding to the structures of over 1400 biomolecules deposited in the Protein Data Bank. J. Biomol. NMR 26, 139−146. (52) Park, C., and Marqusee, S. (2006) Quantitative determination of protein stability and ligand binding by pulse proteolysis. Curr. Protoc. Protein Sci., Chapter 20, Unit 20.11. (53) Park, C., and Marqusee, S. (2005) Pulse proteolysis: a simple method for quantitative determination of protein stability and ligand binding. Nat. Methods 2, 207−212. (54) Laguerre, A., Wielens, J., Parker, M. W., Porter, C. J., and Scanlon, M. J. (2011) Preparation, crystallization and preliminary X-ray diffraction analysis of two intestinal fatty-acid binding proteins in the presence of 11-(dansylamino)undecanoic acid. Acta. Crystallogr., Sect. F 67, 291−295. (55) Verdonk, M. L., Cole, J. C., Hartshorn, M. J., Murray, C. W., and Taylor, R. D. (2003) Improved protein-ligand docking using GOLD. Proteins 52, 609−623. (56) Dixon, S. L., Smondyrev, A. M., Knoll, E. H., Rao, S. N., Shaw, D. E., and Friesner, R. A. (2006) PHASE: a new engine for pharmacophore perception, 3D QSAR model development, and 3D database screening: 1. Methodology and preliminary results. J. Comput.-Aided Mol. Des. 20, 647−671. (57) Velkov, T., Lim, M. L., Horne, J., Simpson, J. S., Porter, C. J., and Scanlon, M. J. (2009) Characterization of lipophilic drug binding to rat intestinal fatty acid binding protein. Mol. Cell. Biochem. 326, 87−95. (58) Velkov, T. (2013) Interactions between human liver fatty acid binding protein and peroxisome proliferator activated receptor selective drugs. PPAR Res., 938401. (59) Hodsdon, M. E., and Cistola, D. P. (1997) Ligand binding alters the backbone mobility of intestinal fatty acid-binding protein as monitored by 15N NMR relaxation and 1H exchange. Biochemistry 36, 2278−2290. (60) Ropson, I. J., Gordon, J. I., and Frieden, C. (1990) Folding of a predominantly beta-structure protein: rat intestinal fatty acid binding protein. Biochemistry 29, 9591−9599. (61) Sacchettini, J. C., Gordon, J. I., and Banaszak, L. J. (1989) Crystal structure of rat intestinal fatty-acid-binding protein. Refinement and analysis of the Escherichia coli-derived protein with bound palmitate. J. Mol. Biol. 208, 327−339. (62) Lahey, K. A., Yuan, R., Burns, J. K., Ueng, P. P., Timmer, L. W., and Kuang-Ren, C. (2004) Induction of phytohormones and differential gene expression in citrus flowers infected by the fungus Colletotrichum acutatum. Mol. Plant-Microbe Interact. 17, 1394−1401. J



Article

(63) Kawashima, Y., Katoh, H., Nakajima, S., Kozuka, H., and Uchiyama, M. (1984) Effects of 2,4-dichlorophenoxyacetic acid and 2,4,5-trichlorophenoxyacetic acid on peroxisomal enzymes in rat liver. Biochem. Pharmacol. 33, 241−245. (64) Chuang, S., Velkov, T., Horne, J., Wielens, J., Chalmers, D. K., Porter, C. J., and Scanlon, M. J. (2009) Probing the fibrate binding specificity of rat liver fatty acid binding protein. J. Med. Chem. 52, 5344− 5355. (65) Velkov, T., Horne, J., Laguerre, A., Jones, E., Scanlon, M. J., and Porter, C. J. (2007) Examination of the role of intestinal fatty acidbinding protein in drug absorption using a parallel artificial membrane permeability assay. Chem. Biol. 14, 453−465. (66) Maloney, E. K., and Waxman, D. J. (1999) trans-Activation of PPARalpha and PPARgamma by structurally diverse environmental chemicals. Toxicol. Appl. Pharmacol. 161, 209−218. (67) Takeuchi, S., Matsuda, T., Kobayashi, S., Takahashi, T., and Kojima, H. (2006) In vitro screening of 200 pesticides for agonistic activity via mouse peroxisome proliferator-activated receptor (PPAR)alpha and PPARgamma and quantitative analysis of in vivo induction pathway. Toxicol. Appl. Pharmacol. 217, 235−244. (68) Hsu, K. T., and Storch, J. (1996) Fatty acid transfer from liver and intestinal fatty acid-binding proteins to membranes occurs by different mechanisms. J. Biol. Chem. 271, 13317−13323. (69) Koch, H. M., Bolt, H. M., and Angerer, J. (2004) Di(2ethylhexyl)phthalate (DEHP) metabolites in human urine and serum after a single oral dose of deuterium-labelled DEHP. Arch. Toxicol. 78, 123−130. (70) Kessler, W., Numtip, W., Volkel, W., Seckin, E., Csanady, G. A., Putz, C., Klein, D., Fromme, H., and Filser, J. G. (2012) Kinetics of di(2ethylhexyl) phthalate (DEHP) and mono(2-ethylhexyl) phthalate in blood and of DEHP metabolites in urine of male volunteers after single ingestion of ring-deuterated DEHP. Toxicol. Appl. Pharmacol. 264, 284− 291. (71) Kluwe, W. M. (1982) Overview of phthalate ester pharmacokinetics in mammalian species. Environ. Health Perspect. 45, 3−9. (72) Schmid, P., and Schlatter, C. (1985) Excretion and metabolism of di(2-ethylhexyl)phthalate in man. Xenobiotica 15, 251−256. (73) Albro, P. W. (1986) Absorption, metabolism, and excretion of di(2-ethylhexyl) phthalate by rats and mice. Environ. Health Perspect. 65, 293−298. (74) Albro, P. W., and Thomas, R. O. (1973) Enzymatic hydrolysis of di-(2-ethylhexyl) phthalate by lipases. Biochim. Biophys. Acta 306, 380− 390. (75) White, R. D., Carter, D. E., Earnest, D., and Mueller, J. (1980) Absorption and metabolism of three phthalate diesters by the rat small intestine. Food Cosmet. Toxicol. 18, 383−386. (76) Albro, P. W., Corbett, J. T., Schroeder, J. L., Jordan, S., and Matthews, H. B. (1982) Pharmacokinetics, interactions with macromolecules and species differences in metabolism of DEHP. Environ. Health Perspect. 45, 19−25. (77) Teirlynck, O. A., and Belpaire, F. (1985) Disposition of orally administered di-(2-ethylhexyl) phthalate and mono-(2-ethylhexyl) phthalate in the rat. Arch. Toxicol. 57, 226−230. (78) Ljungvall, K., Tienpont, B., David, F., Magnusson, U., and Torneke, K. (2004) Kinetics of orally administered di(2-ethylhexyl) phthalate and its metabolite, mono(2-ethylhexyl) phthalate, in male pigs. Arch. Toxicol. 78, 384−389. (79) Kliewer, S. A., Goodwin, B., and Willson, T. M. (2002) The nuclear pregnane X receptor: a key regulator of xenobiotic metabolism. Endocr. Rev. 23, 687−702. (80) Wyde, M. E., Kirwan, S. E., Zhang, F., Laughter, A., Hoffman, H. B., Bartolucci-Page, E., Gaido, K. W., Yan, B., and You, L. (2005) Di-nbutyl phthalate activates constitutive androstane receptor and pregnane X receptor and enhances the expression of steroid-metabolizing enzymes in the liver of rat fetuses. Toxicol. Sci. 86, 281−290. (81) Hostetler, H. A., Balanarasimha, M., Huang, H., Kelzer, M. S., Kaliappan, A., Kier, A. B., and Schroeder, F. (2010) Glucose regulates fatty acid binding protein interaction with lipids and peroxisome proliferator-activated receptor alpha. J. Lipid Res. 51, 3103−3116.

(82) Hostetler, H. A., Kier, A. B., and Schroeder, F. (2006) Very-longchain and branched-chain fatty acyl-CoAs are high affinity ligands for the peroxisome proliferator-activated receptor alpha (PPARalpha). Biochemistry 45, 7669−7681. (83) Hostetler, H. A., McIntosh, A. L., Atshaves, B. P., Storey, S. M., Payne, H. R., Kier, A. B., and Schroeder, F. (2009) L-FABP directly interacts with PPARalpha in cultured primary hepatocytes. J. Lipid Res. 50, 1663−1675. (84) Huang, H., Starodub, O., McIntosh, A., Atshaves, B. P., Woldegiorgis, G., Kier, A. B., and Schroeder, F. (2004) Liver fatty acid-binding protein colocalizes with peroxisome proliferator activated receptor alpha and enhances ligand distribution to nuclei of living cells. Biochemistry 43, 2484−2500. (85) Huang, H., Starodub, O., McIntosh, A., Kier, A. B., and Schroeder, F. (2002) Liver fatty acid-binding protein targets fatty acids to the nucleus. Real time confocal and multiphoton fluorescence imaging in living cells. J. Biol. Chem. 277, 29139−29151. (86) Schroeder, F., Petrescu, A. D., Huang, H., Atshaves, B. P., McIntosh, A. L., Martin, G. G., Hostetler, H. A., Vespa, A., Landrock, D., Landrock, K. K., Payne, H. R., and Kier, A. B. (2008) Role of fatty acid binding proteins and long chain fatty acids in modulating nuclear receptors and gene transcription. Lipids 43, 1−17. (87) Wolfrum, C. (2007) Cytoplasmic fatty acid binding protein sensing fatty acids for peroxisome proliferator activated receptor activation. Cell. Mol. Life Sci. 64, 2465−2476. (88) Gillilan, R. E., Ayers, S. D., and Noy, N. (2007) Structural basis for activation of fatty acid-binding protein 4. J. Mol. Biol. 372, 1246−1260.

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