Article pubs.acs.org/biochemistry
Ligand Binding Properties of the Lentil Lipid Transfer Protein: Molecular Insight into the Possible Mechanism of Lipid Uptake Zakhar O. Shenkarev, Daria N. Melnikova, Ekaterina I. Finkina, Stanislav V. Sukhanov, Ivan A. Boldyrev, Albina K. Gizatullina, Konstantin S. Mineev, Alexander S. Arseniev, and Tatiana V. Ovchinnikova* M. M. Shemyakin and Yu. A. Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Miklukho-Maklaya street, 16/10, 117997 Moscow, Russia
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ABSTRACT: The lentil lipid transfer protein, designated as Lc-LTP2, was isolated from Lens culinaris seeds. The protein belongs to the LTP1 subfamily and consists of 93 amino acid residues. Its spatial structure includes four α-helices (H1−H4) and a long C-terminal tail. Here, we report the ligand binding properties of Lc-LTP2. The fluorescent 2-p-toluidinonaphthalene-6-sulfonate binding assay revealed that the affinity of Lc-LTP2 for saturated and unsaturated fatty acids was enhanced with a decrease in acyl-chain length. Measurements of boundary potential in planar lipid bilayers and calcein dye leakage in vesicular systems revealed preferential interaction of Lc-LTP2 with the negatively charged membranes. Lc-LTP2 more efficiently transferred anionic dimyristoylphosphatidylglycerol (DMPG) than zwitterionic dimyristoylphosphatidylcholine. Nuclear magnetic resonance experiments confirmed the higher affinity of Lc-LTP2 for anionic lipids and those with smaller volumes of hydrophobic chains. The acyl chains of the bound lysopalmitoylphosphatidylglycerol (LPPG), DMPG, or dihexanoylphosphatidylcholine molecules occupied the internal hydrophobic cavity, while their headgroups protruded into the aqueous environment between helices H1 and H3. The spatial structure and backbone dynamics of the Lc-LTP2−LPPG complex were determined. The internal cavity was expanded from ∼600 to ∼1000 Å3 upon the ligand binding. Another entrance into the internal cavity, restricted by the H2−H3 interhelical loop and C-terminal tail, appeared to be responsible for the attachment of Lc-LTP2 to the membrane or micelle surface and probably played an important role in the lipid uptake determining the ligand specificity. Our results confirmed the previous assumption regarding the membrane-mediated antimicrobial action of Lc-LTP2 and afforded molecular insight into its biological role in the plant.
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glandin B2, sterols, molecules of organic solvents, and some drugs.4 The only covalent complex of LTP1 with the allene oxide FA 9(S),10-epoxy-10,12(Z)-octadecadienoic acid was isolated from wheat and barley seeds.7 Like many others PR proteins, LTPs exhibit antimicrobial activity against phytopathogenic bacteria and fungi. At the same time, these proteins do not display any toxicity toward plant or mammalian cells.8,9 The mechanism of LTP antimicrobial action is still obscure. In fact, LTPs cause permeabilization of model lipid bilayers and cytoplasmic membranes of pathogenic fungi.8,10 In plant genomes, LTPs are usually represented by a set of genes encoding different isoforms. Knockout of the LTP genes is accompanied by disturbance of plant vegetative growth and reproductive development.10 Plant LTPs are synthesized as preproteins with N-terminal signal peptides and are mainly extracellularly localized.5,11 Various LTP isoforms are present in different organs and tissues of plants at certain stages of
lant lipid transfer proteins (LTPs) constitute a class of small cationic proteins that belongs to a large family of pathogenesis-related (PR) proteins. The genes of these proteins are constitutively expressed in different organs and tissues of plants, but induction of LTP synthesis takes place under abiotic and biotic stress conditions.1 For this reason, LTPs are believed to be involved in plant stress defense.2,3 Structures of LTPs are stabilized by four disulfide bonds, and perhaps that is why these proteins are so resistant to high temperatures and proteolytic enzymes.4 Plant LTPs are classified into two subfamilies according to their molecular masses: LTP1s (∼9 kDa) and LTP2s (∼7 kDa).5 All representatives of the LTP1 subfamily have similar spatial structures that include four α-helices (H1−H4) and a long Cterminal tail containing several β-turns or one turn of 310-helix (H5). Side chains of the hydrophobic residues form a relatively large tunnel-like cavity containing the ligand-binding site.5,6 Most LTPs bind and transfer diverse lipid molecules in vitro. These proteins reversibly form noncovalent complexes with various ligands, including unsaturated and saturated fatty acids (FAs) with a carbon-chain length of C10−C18, acyl derivatives of coenzyme A (CoA), phospho- and galactolipids, prosta© 2017 American Chemical Society
Received: October 21, 2016 Revised: March 4, 2017 Published: March 7, 2017 1785
DOI: 10.1021/acs.biochem.6b01079 Biochemistry 2017, 56, 1785−1796
Article
Biochemistry ontogenesis.11 LTPs are considered to perform different functions in plants such as adaptation of plants to stress, lipid metabolism, embryogenesis, growth and reproduction of plants, defense signaling, symbiosis, and others.4 It is believed that the involvement of LTPs in many processes in plants is due to their ability to bind and transfer lipids and other hydrophobic molecules. In particular, plant LTPs are assumed to play an essential role in the formation of cutin and suberin wax protective layers, participating in the transfer of their precursors to the appropriate destinations in plant tissues. Formation of LTP−ligand complexes and their stability probably depend on the hydrophobic cavity size and flexibility as well as the spatial structure of the ligand.1,4 However, the detailed mechanisms of ligand binding and selectivity of LTPs remain poorly studied. The tunnel-like cavity of LTP1 runs through the whole molecule and has two entering ends: the first one (termed here as “top”) located between helices H1 and H3 and the second (“bottom”) restricted by the H2−H3 interhelical loop and C-terminal tail.12,13 A number of highresolution X-ray and nuclear magnetic resonance (NMR) structures of plant LTP1 complexes revealed that both entrances have potential sites for ligand binding.12−16 In the observed structures, the acyl chains of the bound lipids enter the cavity, and their polar headgroups are localized at one entrance or both (for doubly liganded LTP1s).15,16 In addition, how the hydrophobic ligands with limited solubility (e.g., phospholipids with two fatty acid chains) can approach and enter the cavity of the water-soluble protein remains unclear. Here, we set a goal to provide molecular insight into the mechanism of lipid uptake by LTPs and to answer the question about a preferred orientation of ligands in the hydrophobic cavity. With this aim in view, we investigated the membrane interactions and ligand binding properties of the lentil Lens culinaris lipid transfer protein (Lc-LTP2, molecular weight of 9282.7 Da, 93 amino acids) that belongs to the LTP1 subfamily.17 The data obtained by a variety of biophysical methods revealed the pronounced Lc-LTP2 selectivity toward anionic phosphatidylglycerol and lysophosphatidylglycerol lipids. The structural NMR study revealed that the lipid molecules (LPPG, DMPG, and DHPC) entered the hydrophobic cavity of Lc-LTP2 from the “top” entrance, while the other interface of the protein enclosing the “bottom” entrance appeared to be initially attached to the membrane surface. The “bottom” lipid-binding site probably plays an important role in the mechanism of lipid uptake and determines ligand specificity. The obtained data confirm previously made assumptions about the mechanism of antimicrobial action of Lc-LTP2 and its possible biological role in the plant.
intensity was measured at 437 nm with excitation at 320 nm using a model F-2710 spectrofluorimeter (Hitachi). TNS (3.5 μM) with or without a lipid (18 μM) was incubated for 1 min in a stirred cuvette containing 1 mL of the measurement buffer [175 mM mannitol, 0.5 mM K2SO4, 0.5 mM CaCl2, and 5 mM MES (pH 7.0)] before the initial fluorescence (F0) was recorded. Then Lc-LTP2 (4 μM) was added, and 2 min later, the fluorescence was recorded at equilibrium (F). The results were expressed as a percentage of the Lc-LTP2−TNS complex fluorescence calculated according to the formula [(F − F0)/FC] × 100%, where FC is the fluorescence of the Lc-LTP2−TNS complex in the absence of a lipid. Lipid Transfer Assay. The fluorescence lipid transfer assay was performed using the fluorescent probe dilution method as previously described.21 Briefly, two lipid probes, a fluorophore [1,3,5,7-tetramethyl BODIPY-labeled phosphatidylcholine (TMB-PC)] and its quencher [bis-cyclohexyl-BODIPY-labeled phosphatidylcholine (BCHB-PC)],22 were placed in small unilamellar vesicles (SUVs). Upon lipid transfer, the probes become diluted and fluorescence arises. To prepare SUVs, lipid suspensions were extruded 10 times through two stacked 100 nm polycarbonate membrane filters (Nucleopore) at ambient temperature using an Avanti Mini-extruder. The labeled vesicles (POPC, 30 μM) containing 0.8% TMB-PC and 1.6% BCHBPC in 200 μL of the buffer containing 20 mM Tris and 1 mM EDTA (pH 7.4) were incubated with 6 μL of the unlabeled vesicle samples (1 mM DMPC or DMPG produced via sonication in the same buffer). Then, 8 μL of the protein sample (360 μM in the same buffer) was added to the mixture, and the change in fluorescence intensity at 505 nm (excitation at 470 nm) was recorded as a function of time at 25 °C while the sample was being constantly stirred. The maximal fluorescence intensity (Ft) was determined by lysing vesicles in the presence of 1.5% Triton X-100. The obtained fluorescence intensity curves were normalized as follows: Fnorm (%) = (F − F0)/(Ft − F0) × 100%
where F is the recorded fluorescence intensity and F0 is the fluorescence intensity measured immediately after the addition of protein. The initial 50 s regions of the normalized curves were linearly approximated to qualitatively compare the transfer efficiency. Leakage of Dye from Lipid Vesicles. The calceinentrapped SUVs composed of POPC, POPG, or POPC and POPG (at a 1:1 molar ratio) were prepared by extrusion in a buffer containing 50 mM calcein, 10 mM HEPES, 200 mM NaCl, and 0.5 mM EDTA (pH 7.5). Untrapped calcein was removed by gel filtration on a Sephadex G-75 column. The eluted calcein-entrapped vesicles were diluted to achieve the desired lipid concentration. The leakage of calcein from the SUVs was monitored by measuring the fluorescence intensity at 520 nm (excitation at 490 nm). The maximal fluorescence intensity (Ft) was determined by lysing vesicles in the presence of 1.5% Triton X-100. The percentage of dye leakage caused by the protein was calculated as follows:
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MATERIALS AND METHODS All the used synthetic phospholipids, lysophospholipids, and lipid extracts were purchased from Avanti Polar Lipids (Alabaster, AL). FAs and 2-p-toluidinonaphthalene-6-sulfonate (TNS) were purchased from Sigma. The deuterated D2O and d4-acetic acid were purchased from CIL (Andover, MA). The recombinant Lc-LTP2 and its 15N-labeled and 13C- and 15 N-labeled analogues were overexpressed in Escherichia coli and purified as described previously.18 Lipid Binding. The ability of Lc-LTP2 to bind FAs was assessed using the TNS fluorescent probe as previously described.19,20 To prevent autoxidation, 0.01% butylated hydroxytoluene (BHT) was added to the polyunsaturated FAs, and the samples were stored at −50 °C. The fluorescence
dye leakage (%) = (F − F0)/(Ft − F0) × 100%
where F0 and F are the fluorescence intensity before and after the addition of protein, respectively. Planar Lipid Bilayers. The bilayer lipid membranes (BLM) mimicking the plasma membrane of Gram-negative bacteria (phospholipid mixture PLG−) were made of the E. coli polar lipid extract consisting of phosphatidylethanolamine, phospha1786
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(NUS) scheme and processed in qMDD software.27 The 3JHNHα, JNHβ, and 3JNHα(−1) coupling constants were measured using 3D HNHA and 3D HNHB experiments. 28 3 J Cγ C′ and 3 JC γN constants for Val, Ile, and Thr residues and 3JCδCα for Leu were calculated from the cross-peak intensities in the spin− echo difference 13C HSQC experiments.28 13C-filtered, 13Cedited NOESY-HSQC spectra with adiabatic pulses in purge elements29 were employed to directly observe the intermolecular nuclear Overhauser effect (NOE) contacts in Lc-LTP2− lipid complexes. 1H chemical shifts were measured relative to the residual protons of H2O, the chemical shift of the signal being arbitrarily chosen as 4.75 ppm at 30 °C. 15N and 13C chemical shifts were referenced indirectly. Spatial structure calculation was performed in CYANA 3.030 The library file with the topology of the LPPG molecule was created manually using MOLMOL.31 The additional pseudoatom, corresponding to all CH2 groups resonating at 1.25 ppm, was defined for the purpose of structure calculation. The lipid molecule was connected to the protein with the flexible linker of dummy residues. Intramolecular and intermolecular distance restraints were derived from the intensities of cross-peaks in 3D NOESY spectra (τm = 80 ms) via a “1/r6” calibration. 1H, 13C, and 15N backbone chemical shifts were used as an input for the TALOS+ software to predict the secondary structure.32 Torsion angle restraints and stereospecific assignments were obtained from J couplings, NOE intensities, and TALOS+ predictions. Hydrogen bonds were introduced on the basis of temperature coefficients of amide protons (Δδ1HN/ΔT) measured over a temperature range of 20−50 °C. The relaxation parameters of 15N nuclei (R1, R2, and 15N− 1 { H} NOE) were measured at 80 MHz using the standard set of 15N HSQC-based pseudo-3D experiments. Model-free analysis of relaxation data was performed in the FastModelFree software33 using an isotropic rotational model. The location and volume of cavities in the proteins were calculated using the CASTp Web server with a 1.4 Å probe radius (http://sts.bioe. uic.edu/castp/calculation.php).
tidylglycerol, and diphosphatidylglycerol (cardiolipin) in a 67:23.2:9.8 ratio (weight percent). The BLM mimicking the plasma membrane of Gram-positive bacteria (phospholipid mixture PLG+) were composed of diphosphatidylglycerol from E. coli, phosphatidylglycerol from E. coli, and soy L-αphosphatidylinositol in a 66:24:10 ratio (weight percent) in accordance with the known phospholipid composition of the plasma membrane of the Gram-positive bacterium Micrococcus lysodeikticus (currently Micrococcus luteus).23 The BLM mimicking the plasma membrane of mammalian cells were prepared from diphytanoylphosphatidylcholine (DPhyPC). Planar BLM were formed of a 5% lipid solution in n-decane by the Mueller technique24 with the 0.85 mm aperture in the Teflon partition separating two stirring compartments, each filled with 2 mL of the aqueous buffer solution containing 20 mM NaCl and 5 mM HEPES (pH 7.4). Membrane formation was observed under reflected light with a microscope. Electrical measurements were taken with Ag-AgCI electrodes connected with both compartments via agar bridges. The Lc-LTP2 protein (0.5 mM) was added 20 min after BLM formation. All experiments were performed at 21 °C. The signals were digitized with the analog input board (EL100E, ADCLab) and stored on the computer. Inner Field Compensation. The protein-induced difference in boundary potentials (ΔU) between two sides of the BLM was studied using the inner field compensation method (IFC) based on registration of the second harmonic of the capacitive current.25 In our setup, the alternating current (ac) signal with a frequency of 300 Hz and an amplitude of 50 mV was generated with a low-harmonic distortion generator (model G118). The ac signal was applied to a membrane with an electrode placed in the cis compartment of the measuring chamber. The trans compartment electrode is connected to current amplifier (model 427, Keithley Instruments Inc., Solon, OH). To remove the first harmonic, the signal was passed through a selective amplifier (model 189, Princeton Applied Research, Oak Ridge, TN). The filtering and detection of the second harmonic were performed using a lock-in amplifier (model 124A, Princeton Applied Research). The negative feedback loop was controlled by a phase of the second harmonic as a lock-in amplifier control set parameter. The output bias voltage was applied via the trans compartment electrode to a membrane; the second harmonic amplitude vanished in case the ΔU was compensated by an external bias voltage. The bias voltage signals were digitized at 10 Hz and stored on the computer for further analysis. NMR Experiments and Spatial Structure Calculation. The NMR investigation was performed using 0.3−0.5 mM samples of 15N-labeled or 13C- and 15N-labeled Lc-LTP2 in a 20 mM perdeuterated sodium acetate/acetic acid buffer solution (pH 5.6, 5% D2O) at 30 °C. LPPG micelles, DMPC or DMPG SUVs (produced via sonication), or DMPC/DHPC bicelles were added to the Lc-LTP2 sample using concentrated stock solutions. The Lc-LTP2/DMPG samples additionally contained 50 mM NaCl and 2 mM EDTA. The NMR spectra were recorded on Bruker Avance III 600 and 800 spectrometers equipped with cryoprobes. 1H, 13C, and 15N resonance assignments of Lc-LTP2 in complex with lipids were obtained using the standard set of three-dimensional (3D) tripleresonance experiments supplemented with 3D 13C HCCHTOCSY and 15N- or 13C-filtered 3D TOCSY and NOESY spectra. Triple-resonance experiments were performed using BEST-TROSY pulse sequences26 and the nonuniform sampling
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RESULTS Binding of Lipids to Lc-LTP2. In our previous work, we used the TNS fluorescent probe to study the ability of Lc-LTP2 to interact with saturated FAs with chain lengths of C12−C19, unsaturated FAs with chain lengths of C16−C18 containing one to three double bonds, lysolipids with different chain lengths (C14−C16), and jasmonic acid (JA).34 TNS was highly fluorescent when bound to the hydrophobic cavity of the protein and competed with lipid molecules for binding to LcLTP2. When the protein was added to a mixture of TNS and a lipid, the fluorescence of TNS became lower than that of the control performed by addition of Lc-LTP2 to TNS alone. This permitted us to assay and compare affinities of different lipids for LTP. In this study, we present an extended analysis with the use of fatty acids with longer chain lengths: saturated FAs with a chain length of C22 or C24 and unsaturated FAs with a chain length of C20 or C22 containing four or five double bonds. The data from both studies are summarized in Figure S1. No quenching between each of the FAs and TNS alone was detected, showing that these molecules did not interact directly (data not shown). Unsaturated FAs (C20:4 and C22:5) displaced TNS from the hydrophobic cavity of Lc-LTP2 less efficiently than palmitoleic (C16:1, cis-9), oleic (C18:1, cis-9), elaidic (C18:1, trans-9), linoleic (C18:2, all-cis-9,12), or linolenic (C18:3, all-cis-9,12,15) 1787
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Figure 1. Assay of lipid transfer between the labeled (POPC) and unlabeled liposomes: (A) unlabeled DMPG liposomes and (B) unlabeled DMPC liposomes. The fluorescence intensity increased because of the transfer of lipid molecules between the liposomes upon addition of Lc-LTP2. The fluorescence intensity reaches a plateau when the fluorescent lipid molecules are equally distributed between the liposomes.
neutral DPhyPC) was observed (Figure 2). It should be noted that the kinetics of the boundary potential for all the BLM
acid. Saturated FAs with a chain length of C22 or C24 also competed with TNS for binding to the hydrophobic pocket of Lc-LTP2 with an efficiency lower than that of shorter molecules (Figure S1). Transfer of Lipids. LTPs can transfer and/or exchange various phospholipids and glycolipids across membranes in vitro.4 The lipid transfer properties of Lc-LTP2 were investigated by fluorescence spectroscopy with the use of labeled lipids. Two types of SUVs, the labeled POPC liposomes and the unlabeled DMPC or DMPG liposomes, were mixed together. The labeled liposomes contain fluorescent lipid probe TMB-PC and its fluorescence resonance energy transfer quencher, BCHB-PC. Before LTP was added to the mixture, the fluorescence of TMB-PC was quenched. Addition of LTP causes changes in TMB-PC fluorescence. The fluorescence intensity increased steadily with time until a plateau was reached (Figure 1). This illustrated the dilution of the labeled lipids in the labeled liposomes due to the transfer activity of the protein. One possible reason for the fluorescence increase is the transfer of TMB-PC or BCHB-PC to DMPG/DMPC liposomes, which leads to the dilution of labeled lipids. It is important to notice that the percentage of fluorescent probes was low (0.8 and 1.6%); thus, most likely the transfer of matrix lipids (DMPC, DMPG, and POPC) had a primary impact on the dilution of labeled lipids and the corresponding fluorescence increase. The values of the initial slope of the fluorescence intensity curves after protein addition permitted us to assay and compare the transfer rates for different lipids. Under otherwise identical conditions and at the same lipid concentrations, Lc-LTP2 more efficiently transferred charged lipid DMPG (approximately by an order of magnitude) than DMPC, and the slopes calculated over initial linear 50 s regions of the curves were 0.13 ± 0.01 and 0.019 ± 0.002%/s, respectively. Boundary Potential Measurement. The ability of LcLTP2 to interact with membranes of different compositions was studied by boundary potential measurement with the use of BLM. The dependence of the boundary potential difference on Lc-LTP2 concentration for BLM with various lipid compositions (PLG−, 33% negatively charged phospholipids, mole percentage; PLG+, 100% negatively charged phospholipids;
Figure 2. Dependence of the boundary potential on the concentration of Lc-LTP2 for BLM of various lipid compositions: (1) DPhyPC, (2) PLG−, and (3) PLG+. The protein was added to the cis side of BLM.
showed a tendency to saturation, as the curves reached a plateau over time (data not shown). The potential values measured on the BLM containing negatively charged phospholipids (PLG− or PLG+) were significantly higher than that determined on the neutral membranes (DPhyPC). Moreover, the biggest difference in the boundary potentials was observed for the membranes composed of PLG+. The obtained data indicated that the affinity of Lc-LTP2 for the membrane surface strongly depended upon the charge of lipid molecules. Liposome Leakage Assay. The ability of Lc-LTP2 to induce the leakage of an encapsulated fluorescent marker calcein from small unilamellar vesicles was studied. SUVs composed of a zwitterionic phospholipid (POPC), an anionic phospholipid (POPG), and their mixture (1:1 POPC/POPG) were tested. Lc-LTP2 induced significant leakage of the POPG vesicles depending upon the protein concentration and reached a plateau at 40 μM (Figure 3). At the same time, no significant leakage of the POPC and POPC/POPG SUVs was observed at much higher concentrations of Lc-LTP2 [≤60 μM (data not shown)]. Despite the moderate level of the SUV leakage detected under the used experimental conditions, the obtained 1788
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protein structure. (1) The kink angle of helix H1 increased from ∼60° to ∼70°. (2) Helices H2 and H3 became longer by one amino acid residue at their C-termini. (3) The conformations of Cys4−Cys51 and Cys49−Cys88 disulfide bridges were changed. (4) Two additional backbone−backbone hydrogen bonds were formed, one between the H1−H2 and H3−H4 loops [HN Asn63···CO Leu18 (Figure 4B)] and one at the C-terminus (HN Thr86···CO Ser83). (5) Finally, the volume of the protein hydrophobic cavity increased from ∼600 to ∼1000 Å3. The unambiguous detection of intermolecular lipid−protein interactions by regular NOESY-type experiments usually is complicated by a background of intramolecular (lipid−lipid and protein−protein) NOE contacts. To facilitate this task, we employed a special 13C-filtered NOESY experiment29 with the use of 13C purge and 13C filter elements, which selects only one magnetization pathway going from a lipid (1H−12C spin pair) to a protein (1H−13C spin pair). In summary, 55 intermolecular contacts were observed in the filtered NOESY spectrum of LPPG-bound Lc-LTP2: nine contacts with the lipid methyl group at 0.85 ppm, four contacts with the α-CH2 group at 2.35 ppm, two contacts with the β-CH2 group at 1.55 ppm, and 40 contacts with other CH2 groups resonating at 1.25 ppm. No contacts were detected between the protein and the LPPG headgroup. As a result, the NOE data unambiguously defined the orientation and entrance site of the lipid molecule inside the hydrophobic cavity of Lc-LTP2 (Figure 4A). LPPG enters the cavity between helices H1 and H3 (Figure 4B, left), which explains the altered kink angle of helix H1. This “top” entrance into the cavity is crosswise restricted by the Cys4−Cys51 disulfide bridge and HN Asn63···CO Leu18 hydrogen bond (Figure 4B). The carbonyl and the first glycerol group of the lipid could contact polar uncharged side chains of helices H1 and H3 and the H3−H4 loop (Figure 4B). At the same time, the phosphoglycerol moiety of LPPG is located outside the protein and its conformation is poorly defined (Figure 4A). The acyl chain of LPPG passes through the Lc-LTP2 molecule along helix H3, and its terminal methyl group is located in the middle of the protein globule between the helices H2 and H3 and the C-terminal region on the level of the Cys49−Cys88 disulfide bond (Figure 4A,B). Interaction of Lc-LTP2 with Two Acyl-Chain Phospholipids. To investigate the binding of two-chain phospholipids by Lc-LTP2, we initially titrated the Lc-LTP2−LPPG complexes (LPR of 8:1) with DMPG or DMPC SUVs. No changes in the 1H−15N HSQC spectra were observed even with a 3-fold excess of the two acyl-chain phospholipids as compared with the lysolipid, indicating that two acyl-chain phospholipids have significantly lower affinity for Lc-LTP2 than LPPG does. Titration of the Lc-LTP2 apo form with the solution of DMPC vesicles also did not result in detectable lipid binding (Figure S2). It should be noted that the used NMR experiments did not permit detection of small (less than ∼30 μM) amounts of the Lc-LTP2−DMPC complex under the background of the apo form of the major protein. Assuming that the exchange between apo-Lc-LTP2 and the Lc-LTP2−DMPC complex is slow on the NMR time scale, we could estimate that the population of the protein in complex with DMPC is ≲10% of the total protein in the sample (0.3 mM). Obtained data indicated that under the used experimental conditions (pH 5.6) Lc-LTP2 could not efficiently take up zwitterionic lipids from the bilayer. To disrupt packing of the bilayer and facilitate the capture of the lipid by Lc-LTP2, the Lc-LTP2/DMPC sample was titrated
Figure 3. Percentage of leakage of the calcein dye from POPG LUVs upon addition of Lc-LTP2.
data proved Lc-LTP2 has the ability to permeabilize certain model membranes. Spatial Structure of the Lc-LTP2−LPPG Complex. TNS fluorescence assay revealed that lysolipid LPPG has the highest affinity for Lc-LTP2 among all studied lipids (Figure S1). Therefore, to investigate the structural basis of Lc-LTP2 ligand binding, we initially tested this anionic lysophosphatidylglycerol as the ligand of the protein. To prepare the Lc-LTP2−LPPG complexes, aliquots of a 10% LPPG solution were sequentially added to the 0.5 mM [13C,15N]Lc-LTP2 sample until the lipid:protein molar ratio (LPR) reached 8:1. Evident changes were observed in the NMR spectra of the protein upon LPPG binding. The majority of cross-peaks in 15N and 13C HSQC spectra changed their position significantly (Figures S2 and S3). The NMR data revealed that at an LPR of 1:1 all the protein in the sample was converted into the lipid-bound form. The slight decrease in the intensity of the NMR signals observed at higher LPRs may correspond to transient binding of the protein to the surface of the LPPG micelle (data not shown). The set of 20 spatial structures (Figure 4A) of the LPPGbound form of the protein was calculated from the obtained NMR data (Table S1). The secondary structure of the protein involves four α-helices: Cys4−Leu18 (H1), Pro26−Ala38 (H2), Thr42−Gly57 (H3), and Thr64−Lys73 (H4). In addition, the C-terminal region of the protein (Gly75− Phe93) encloses one turn of 310-helix [Cys88−Thr90 (H5)]. The conserved Pro13 residue induces a pronounced kink (∼70°) in helix H1, thus disrupting the α-helical conformation within the region of residues Asp10 and Leu11. Three α-helices (H1−H3) form a boat-shaped bundle that restricts the hydrophobic cavity. This cavity lined with apolar side chains is shielded from the aqueous environment by helix H4 and the C-terminal tail (Figure 4A). Like the apo form of the protein,18 the Gly23−Pro24 peptide bond of lipid-bound Lc-LTP2 is in the cis conformation. The spatial structure of the Lc-LTP2 molecule was not changed much upon LPPG binding (Figure 4C). The structures of the apo and liganded forms of the protein can be fairly well superimposed over backbone atoms of helices H1−H4 [root-mean-square deviation (RMSD) ∼ 1.5 Å], while superimposition over all backbone atoms results in a RMSD value ∼2.6 Å. In line with it, the structural comparison revealed significant differences only in the conformations of the H3−H4 interhelical loop and in the C-terminal region (Figure 4C). The binding of the LPPG molecule induced other changes in the 1789
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Figure 4. (A and B) Spatial structure of the Lc-LTP2−LPPG complex and its comparison with those of (C) unliganded Lc-LTP2 and (D) other plant LTPs in complex with ligands. (A) The sets of the 20 best structures are superimposed on the backbone atoms. The disulfide bonds are colored orange. Helices H1−H5 are color-coded. The LPPG molecule is drawn as balls and sticks. (B) Two-sided view of the Lc-LTP2−LPPG complex. The positively charged (Arg, Lys, N-terminal amino group) and negatively charged (Asp, C-terminal carboxylic group) side chains and/or moieties are colored blue and red, respectively. The “bottom” entrance into the internal hydrophobic cavity is shown with an arrow; the “top” entrance is occupied by the LPPG molecule. The polar or aromatic (Ser, Thr, Asn, and Tyr) and hydrophobic (Ala, Ile, Leu, and Val) side chains that restrict entrances into the internal cavity are colored green and yellow, respectively. The Pro residues are shown as magenta plates. The HN Asn63···CO Leu18 hydrogen bond is shown in the left panel. (C) Spatial structures of the apo form (yellow) and LPPG-bound state (blue) of Lc-LTP2 superimposed on the backbone atoms. (D) Complexes of the rice LTP1 with stearate (PDB entry 1UVC14) and of the wheat LTP1 with two molecules of LMPC (PDB entry 1BWO15).
with the “short” acyl-chain lipid DHPC. DHPC has detergentlike properties and upon being mixed with DMPC forms bicelles.35 The NMR spectra of Lc-LTP2/DMPC/DHPC samples revealed the presence of two different forms of the protein corresponding to DMPC- and DHPC-bound states (Figure S2). The relative population of the states depended on the DMPC:DHPC ratio (from 0.3 to 2.0); however, at all tested concentrations, the population of the DHPC-bound protein was 2−4 times higher. This indicates that the shortchain DHPC has a higher affinity for Lc-LTP2 than the longchain DMPC does. As a result, the assignment of the NMR signals was possible only for the major protein form, the LcLTP2−DHPC complex. The lipid transfer experiments (Figure 1) revealed that LcLTP2 could efficiently take up anionic lipids from the bilayer. Previously, using NMR spectroscopy, we observed the
formation of the Lc-LTP2−DMPG complexes upon titration of the apo form of the protein with DMPG SUVs, but the obtained complexes were unstable and segregated into the apoprotein and precipitated lipids,18 In the work presented here, we observed that the addition of 50 mM NaCl and 2 mM EDTA to the Lc-LTP2−DMPG sample [LPR of 1:8 (Figure S2)] stabilized the lipid−protein complexes for the time span that was necessary to perform the experiments for resonance assignments and detection of intermolecular NOE contacts. No intermolecular contacts were observed for α-CH2 and β-CH2 groups of DMPG, but four contacts were identified between methyl groups of the lipid (0.85 ppm) and methyl groups of residues Ala39, Ala48, Leu70, and Ile82 (Figure 5B, stars). These residues are located in the H2−H3 loop, the H4 helix, and the C-terminal tail, indicating that the two-acyl-chain lipid 1790
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Figure 5. Changes in the spatial structure and backbone dynamics of Lc-LTP2 upon ligand binding and proposed mechanism of lipid uptake. (A) Standard deviation of 13Cα and 1HN chemical shifts in the four sets of NMR chemical shifts (apoprotein and the protein liganded by LPPG, DMPG, and DHPC). The full sets of measured secondary 13Cα and 1HN chemical shifts are presented in Figure S3. The large variations (>0.25 and >0.1 ppm for 13Cα and 1HN, respectively) are colored red and blue, respectively. The residues that restrict the “top” and “bottom” entrances in the internal cavity are boxed and underlined on the protein sequence, respectively. (B) The ribbon of Lc-LTP2 (in complex with LPPG) is colored according to the obtained chemical shift variations (panel A). Residues with the observed intermolecular contacts with the methyl group of DMPG are marked with green stars. The “bottom” entrance into the internal hydrophobic cavity is shown with an arrow; the “top” entrance is occupied by the LPPG molecule. (C) The ribbon of Lc-LTP2 in the apo form (left) and liganded by LPPG (right) is colored according to obtained dynamical NMR data. The full set of measured 15N relaxation data and the results of their analysis are shown in Figure S4. The residues displaying 15N−{1H} NOE values of 18 s−2 could be subjected to exchange fluctuations on the microsecond to millisecond time scale.37 The results of the “model-free” analysis33 of the obtained 15N relaxation data confirmed the obtained qualitative conclusions (Figure S5). (D) Proposed mechanism of binding of Lc-LTP2 to the surface of the LPPG micelle or DMPG membrane (gray slab) and a probable path of lipid uptake (details in the text).
regions are located in the proximity of the acyl chain of the ligand in the Lc-LTP2−LPPG complex. Therefore, the observed variations probably reflect the adaptation of the protein cavity to the binding of ligands with different volumes of hydrophobic moieties (the number and lengths of acyl chains). Only minor changes in chemical shifts were observed on the other side of the Lc-LTP2 molecule, where the polar headgroup of the ligand exits the hydrophobic cavity. The large changes were detected for Leu11 (helix H1), Gly57 (helix H3), and Ile59 (H3−H4 loop). The small structural variations in this region of the protein indicate that bound ligands are mostly stabilized by the hydrophobic interactions of acyl chains within the internal cavity, but not by electrostatic interactions and hydrogen bonding with the polar headgroups. The obtained data also confirmed that zwitterionic phosphatidylcholine lipid (DHPC) interacted with Lc-LTP2 like anionic phosphatidyl-
adopts the same orientation inside the cavity of Lc-LTP2 as LPPG (Figure 5B). Adaptation of the Lc-LTP2 Molecule to Different Ligands. A previous NMR study18 and experiments described above resulted in four sets of NMR chemical shifts for LcLTP2, corresponding to the apo form (BMRB accession code 19365) and the protein liganded by LPPG, DMPG, and DHPC (Figure S4). To compare the spatial structure of Lc-LTP2 in different states, we used chemical shifts of 13Cα and 1HN nuclei, which are sensitive to the values of backbone dihedral angles and to the hydrogen bond strength of the amide groups, respectively.36 Analysis of variations of these chemical shifts within the four sets of NMR data (Figure 5A,B) revealed rather compact localization of the structural changes in the Lc-LTP2 molecule. The largest variations were observed at the protein N-terminus (Ile2-Ser3), the C-terminal part of helix H2 and the H2−H3 loop (Lys33−Thr41), the middle part of helix H3 (Cys49−Lys53), and the C-terminal tail (Ile78−Phe93). These 1791
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glycerols (LPPG and DMPG) did and entered the cavity from the “top” entrance. Intramolecular Mobility of Liganded and Unliganded Lc-LTP2. To investigate changes in the dynamics of Lc-LTP2 induced by ligand binding, we measured the 15N relaxation parameters (R1 and R2 rates and heteronuclear 15N−{1H} NOEs) for the apo and LPPG-bound forms of the protein (Figure S5). The qualitative analysis of these data revealed that in both cases the protein backbone did not accomplish significant picosecond to nanosecond time scale motions (average heteronuclear NOE values of 0.84 ± 0.08 and 0.84 ± 0.10, respectively). Only several residues belonging to the Nterminus of the protein, the H2−H3 and H3−H4 loops, and the C-terminal tail demonstrate increased mobility on this time scale [NOE values of 0.75, and R1R2 < 18 s−2). The obtained average values (4.3 ± 0.6 and 5.2 ± 0.3 ns, corresponding hydrodynamic radii of ∼17.5 and 18.7 Å, respectively) revealed significant deceleration of the protein rotational diffusion upon ligand binding. This is unexpected, taking into account the fact that the dimensions of the protein remain almost unchanged. Thus, the observed deceleration is related to the transient adsorption of the protein on the surface of negatively charged LPPG micelles presented in the sample (LPR of 8:1). This surface-bound state of Lc-LTP2 is relatively weakly populated, and the exchange between lipid-bound soluble and lipid-bound surface adsorbed states is fast on the NMR chemical shift time scale. Probably, this exchange process is at least partially responsible for the increase in microsecond to millisecond time scale mobility observed in the LPPG environment. The analysis of the distribution of the charged residues in the Lc-LTP2 structure (Figure 4B) revealed that almost all positively charged side chains (except Lys61) are exposed on one side of the protein. This interface formed by the fragments of helices H2 and H3, the H2−H3 loop, and the C-terminal tail demonstrates a significant increase in microsecond to millisecond mobility in the LPPG environment. Therefore, we can conclude that the Lc-LTP2 molecule binds to the surface of the detergent micelle via this interface, which encloses the “bottom” entrance of the cavity (Figure 5D).
Article
DISCUSSION
Lipid Selectivity of Lc-LTP2. Regardless of the spatial structural similarities of different plant LTPs, they are known to have different ligand binding properties and lipid transfer activities.12,38 These features are considered to be largely important for the variety of LTP functions in plants, e.g., for lipid metabolism and formation of cutin and suberin wax protective layers. Cutin is a polyester biopolymer composed mostly of hydroxy, epoxy, and α,ω-dicarboxylic derivatives of saturated C16 and unsaturated C18 FAs.39 The presence of a substantial proportion of aromatics, α,ω-dicarboxylic acids, or long-chain FAs (>C20) is typical of suberin.39 The composition of the suberin and cutin layers depends on the plant species.39 Probably, LTPs from various plants bind different lipids with diverse efficiencies and transfer them during the formation of cutin and suberin protective layers. Previously, we isolated a new lipid transfer protein, designated as Lc-LTP2, from germinated seeds of the lentil L. culinaris.17 The ligand binding, studied by a fluorescence TNS assay [ref 34 and the work presented here (see Figure S1)], revealed that lentil Lc-LTP2 can bind a wide range of lipids, including JA, saturated and unsaturated FAs with chain lengths of 12−24 carbon atoms, and zwitterionic and anionic lysolipids, with very different efficiencies. Among the saturated FAs, the affinity of Lc-LTP2 is almost linearly decreased with an increase in acyl-chain length. Thus, the highest and lowest binding capacities were observed for C12 and C24 FAs, respectively. The ability of Lc-LTP2 to bind unsaturated FAs strongly depended not only on chain length but also on the number and position of double bonds. Again the long acyl-chain FAs (C20 and C22) demonstrated lower affinity for the protein. The low affinity of Lc-LTP2 for the long-chain lipids is probably relevant to the tissue specificity of the protein as FAs with chains of >20 carbons are typical for suberin, which was found in root epidermis or endodermis and in bundle sheaths but not in seeds where Lc-LTP2 was discovered. The highest efficiency of binding was observed for unsaturated FAs with chain lengths of 16 and 18 carbon atoms (Figure S1). These particular unsaturated FAs are precursors of cutin monomers. A similar specificity for different fatty acids has been previously described for wheat,40 tobacco,19 and ginkgo LTP1s,41 and for a number of other proteins that bind and transfer lipids, for instance, elicitins42 and Bet v1 homologues.43 However, such specificity is not characteristic of all plant LTPs; for example, onion AceAMP1 does not bind lipids,44 dill Ag-LTP shows a lack of selectivity toward various FAs,34 and tomato XSP10 binds only saturated FAs.45 The fluorescence lipid transfer assay revealed that Lc-LTP2 exchanged lipids between POPC and anionic DMPG liposomes much more efficiently than between POPC and zwitterionic DMPC SUVs (Figure 1). The observed preferential affinity of Lc-LTP2 for the negatively charged lipids is in agreement with the results obtained with lysolipids [LMPG and LPPG (Figure S1)] and with the results of NMR titration experiments with DMPG and DMPC SUVs (see Results). Moreover, NMR experiments showed that the short two-chain lipid DHPC (2 × C6) interacted with the protein more efficiently than longer two-chain lipid DMPC (2 × C14) did. This correlates well with the results of the FA binding study (Figure S1), revealing that shorter, and consequently less bulky, molecules demonstrate higher affinity for the protein. It should be noted that the measurement of the relative affinity of the ligands could be 1792
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various sizes, from small short-chain FAs to bulky phospholipids. The obtained structural data do not provide a straightforward explanation for the selectivity of Lc-LTP2 for anionic lipids. The “top” entrance of the cavity accommodates only one positively charged residue (Lys61), and the amino group of its side chain does not contact the negatively charged phosphate group of the bound lipid (Figure 4B). On the other hand, three positively charged residues (Arg45, Lys81, and Lys92) are located in the proximity of the “bottom” entrance (Figure 4B). The selective interaction of Lc-LTP2 with anionic membranes (Figure 2) and the observed low efficiency of uptake and transfer of zwitterionic DMPC lipid (see Figure 1 and NMR data described above) indicate that the initial binding of the protein to the membrane surface is a necessary step required for the subsequent uptake of lipid molecules. Taking into account the fact that Lc-LTP2 binds to the surface of the LPPG micelle via an interface that includes the “bottom” entrance of the cavity (Figure 5D), we could propose the simple mechanism for the lipid uptake process that explains the charge selectivity of Lc-LTP2. In the proposed model (Figure 5D), Lc-LTP2 binds to the membrane or micelle surface by its positively charged “bottom” interface. Perhaps this interface initially functions as a lipidbinding site that aids in protein−membrane docking similar to that of lipid-binding domains of other proteins.47 Lc-LTP2− membrane binding probably is guided by selective interaction between the protein residues located at the “bottom” entrance and the negatively charged lipid headgroup. Then, the lipid penetrates the cavity and becomes captured at the “top” entrance (Figure 5D). Thus, the “bottom” entrance of Lc-LTP2 plays several important roles in the mechanism of lipid uptake: as a membrane-binding domain, it directs the protein to the membrane surface and determines the ligand selectivity, and at the same time, it functions as a transshipment element in the lipid pathway to the “top” entrance. The proposed model is in line with the results of a previous limited mutagenesis study of wheat LTP1.48 The substitution of the conserved Gln46 and Tyr80 residues (the numbers are in accordance with Lc-LTP2 numbering) belonging to the “bottom” entrance of the cavity did not influence the lipid transfer activity of the protein. At the same time, the fluorescence data revealed that the side chain of Tyr80 of wheat LTP1 is located in the proximity of the lipid (or membrane)-binding site.48 Role of Interactions with a Membrane in the Antimicrobial Activity of Lc-LTP2. Like other plant LTP1s,49,50 lentil Lc-LTP2 displays antimicrobial activity and inhibits the growth of phytopathogenic fungi and bacteria at relatively high concentrations.17,18 The mechanism of plant LTP antimicrobial action remains unclear, but the cell membrane is considered one of the most probable targets. Consistent with this hypothesis, the boundary potential measurements revealed that Lc-LTP2 interacted with the BLM of compositions mimicking those of membranes of Gram-positive (PLG+) and Gram-negative bacteria (PLG−) as well as mammal cell membranes (Figure 2). The protein demonstrated higher affinity for the membrane containing 100% negatively charged phospholipids (PLG+). Besides, the membrane permeabilization study revealed an ability of LcLTP2 to induce the leakage of calcein from negatively charged POPG liposomes (Figure 3). Interestingly, the considerable permeabilization of POPG SUVs was observed at a Lc-LTP2
biased by differences in their solubility. Thus, the higher solubility of detergent-like LPPG and DHPC molecules, as well as of short FAs, could facilitate their access to the binding site on the LTP molecule. Structure and Dynamics of Lc-LTP2−Ligand Complexes and Proposed Mechanism of Lipid Uptake. The obtained NMR data revealed that anionic and zwitterionic lipid molecules (LPPG, DMPG, and DHPC) entered the Lc-LTP2 hydrophobic cavity from the “top” entrance, formed by the residues belonging to helices H1 and H3 and the H3−H4 interhelical loop (Figures 4B and 5A,B). Such an orientation of the bound ligands is opposite to the positions of the FAs observed in the high-resolution X-ray structures of maize and rice LTP1 complexes.13,14 In these structures, the carboxylate moiety of the ligands interacted with the “bottom” entrance located between the H2−H3 loop and C-terminal tail of the proteins (Figure 4D, left; the “bottom” entrance is also marked with an arrow on the Lc-LTP2 structures shown in Figures 4 and 5). A similar (“bottom”) orientation of the ligand was also proposed as a result of the NMR study of wheat LTP1 in complex with DMPG.46 At the same time, the NMR study of barley LTP1 in complex with palmitate and palmitoyl-CoA revealed that the polar headgroups of the ligands occupy the “top” entrance12 as seen for the Lc-LTP2−LPPG complex. Previous investigations of other plant LTP1s complexes indicated that these proteins could simultaneously bind several lipids fixing their headgroups at the different entrances of the hydrophobic cavity. For example, two bound LMPC molecules occupying both “top” and “bottom” entrances were observed in the crystal structure of wheat LTP1 (Figure 4D, right).15 Similar doubly liganded complexes were described for peach LTP1 (Pru p 3).16 Moreover, the crystal structures of rice LTP1 in complex with two palmitate molecules14 and of the barley LTP1 in complex with three lysophosphocholine molecules (PDB entry 1MID) revealed that the LTP1 hydrophobic cavities could have an additional entrance localized between the H3−H4 interhelical loop and the Cterminal tail. The number and localization of the lipid-binding sites in the cavity of LTP1s probably depend on the individual properties of a protein−ligand pair. At the same time, an influence of the crystal packing and individual crystallization conditions on the occupancy of different sites should not be excluded. In this case, the structures provided by solution NMR better correspond to the LTP−lipid complexes, which are present in the plant tissues. The selectivities of Lc-LTP2 toward different FAs and phospholipids are probably determined by the protein structure and dynamics, e.g., plasticity and adaptability of the hydrophobic cavity as well as charge distribution on the surfaces that form the cavity entrances. The structural comparison of apo and liganded forms of Lc-LTP2 (Figure 5A,B) and the observed distributions of the picosecond to nanosecond and microsecond to millisecond time scale mobility (Figure 5C) permitted us to distinguish two pseudodomains of the protein with different properties. The “stable” domain is linked by the Cys14−Cys28 and Cys29−Cys74 disulfide bridges and involves the C-terminal part of helix H1, the H1−H2 loop, and the Nterminal parts of helices H2 and H4. Another “mobile” domain is formed around Cys4−Cys51 and Cys49−Cys88 disulfides and includes the C-terminus of helix H2, the H2−H3 loop, helix H3, and the C-terminal tail of the protein (Figure 5A−C). The plasticity of the latter domain is probably responsible for the adaptation of the Lc-LTP2 cavity to the ligands of 1793
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Biochemistry concentration of 40 μM, which is comparable to the IC50 values determined in the antimicrobial assays (10−40 μM).17,18 This result is consistent with the finding that plant LTPs are active against pathogenic bacteria and fungi and exhibit no toxic effects on mammalian and plant cells, outer membranes of which mainly consist of neutral zwitterionic phospholipids.51 The obtained results confirmed the previously made assumptions about the mechanism of antimicrobial action of Lc-LTP2.18 All lipid binding and transport experiments as well as experiments on the interaction of protein with model membranes revealed a pronounced selectivity of lentil Lc-LTP2 for anionic lipids. Taking into consideration the fact that the Lc-LTP2 molecule surface does not contain large hydrophobic patches (Figure 4C), we could conclude that electrostatic interactions appear to play a crucial role in the binding of protein to cell membranes of phytopathogens. Like the mode of action proposed for amphipathic membrane active antimicrobial peptides,52 the surface accumulation of lentil Lc-LTP2 destabilizes the phospholipid fatty acyl packing in the membranes of pathogenic bacteria and fungi and finally causes transient membrane opening, rupture, and ultimately lysis. However, the efficiency of membrane permeabilization by lentil Lc-LTP2 and other plant LTP1s8,44,51 is significantly lower than that reported for amphipathic antimicrobial peptides52 like LL37 or magainin. According to the obtained data, we proposed a hypothesis regarding the possible biological role of lentil Lc-LTP2. This protein is presented in both dry and germinated lentil seeds and obviously takes part in lipid metabolism and transfer of unsaturated FAs, in particular, to the place of cutin biosynthesis. Under stress conditions, including phytopathogen attack, LcLTP2 synthesis may be activated in the plant. This fact is demonstrated for other plant LTPs and representatives of defense PR proteins. At higher concentrations that could be triggered under different stress conditions, Lc-LTP2 may exhibit antimicrobial activity against phytopathogenic bacteria and fungi and complements the spectrum of defense proteins that effectively hinder the spread of infection. Our results afford further molecular insight into the biological role of lipid transfer proteins.
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Author Contributions
Z.O.S. and D.N.M. contributed equally to this work. Funding
This work was supported by the Russian Science Foundation (Project 14-50-00131). Notes
The authors declare no competing financial interest.
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ABBREVIATIONS BCHB, bis-cyclohexyl-BODIPY; BLM, bilayer lipid membranes; DHPC, 1,2-dihexanoyl-sn-glycero-3-phosphocholine; DMPC, 1,2-dimyristoyl-sn-glycero-3-phosphocholine; DMPG, 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol; DPhyPC, 1,2diphytanoyl-sn-glycero-3-phosphocholine; FA, fatty acid; JA, jasmonic acid; Lc-LTP2, second isoform of the lipid transfer protein from L. culinaris; LMPC, 1-myristoyl-2-hydroxy-snglycero-3-phosphocholine; LPPG, 1-palmitoyl-2-hydroxy-snglycero-3-phosphoglycerol; LPR, lipid:protein molar ratio; LTP, lipid transfer protein; PC, phosphatidylcholine; PDB, Protein Data Bank; PLG−, phospholipid mixture mimicking the plasma membrane of Gram-negative bacteria; PLG+, phospholipid mixture mimicking the plasma membrane of Grampositive bacteria; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3phosphocholine; POPG, 1-palmitoyl-2-oleoyl-sn-glycero-3phosphoglycerol; SUV, small unilamellar vesicle; TMB, 1,3,5,7-tetramethyl-BODIPY; TNS, 2-p-toluidinonaphthalene6-sulfonate.
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(1) Liu, F., Zhang, X., Lu, C., Zeng, X., Li, Y., Fu, D., and Wu, G. (2015) Non-specific lipid transfer proteins in plants: Presenting new advances and an integrated functional analysis. J. Exp. Bot. 66, 5663− 5681. (2) Jang, C. S., Lee, H. J., Chang, S. J., and Seo, Y. W. (2004) ) Expression and promoter analysis of the TaLTP1 gene induced by drought and salt stress in wheat (Triticum aestivum L.). Plant Sci. 167, 995−1001. (3) Jung, H. W., Kim, W., and Hwang, B. K. (2003) Three pathogeninducible genes encoding lipid transfer protein from pepper are differentially activated by pathogens, abiotic, and environmental stresses. Plant, Cell Environ. 26, 915−928. (4) Finkina, E. I., Melnikova, D. N., Bogdanov, I. V., and Ovchinnikova, T. V. (2016) Lipid transfer proteins as components of the plant innate immune system: structure, functions, and applications. Acta Naturae 8, 47−61. (5) Douliez, J.-P., Michon, T., Elmorjani, K., and Marion, D. (2000) Mini review: structure, biological and technological functions of lipid transfer proteins and indolines, the major lipid binding proteins from cereal kernels. J. Cereal Sci. 32, 1−20. (6) Sy, D., Le Gravier, Y., Goodfellow, J., and Vovelle, F. (2003) Protein stability and plasticity of the hydrophobic cavity in wheat nsLTP. J. Biomol. Struct. Dyn. 21, 15−29. (7) Bakan, B., Hamberg, M., Larue, V., Prange, T., Marion, D., and Lascombe, M. B. (2009) The crystal structure of oxylipin-conjugated barley LTP1 highlights the unique plasticity of the hydrophobic cavity of these plant lipid-binding proteins. Biochem. Biophys. Res. Commun. 390, 780−785. (8) Regente, M. C., Giudici, A. M., Villalaín, J., and de la Canal, L. (2005) The cytotoxic properties of a plant lipid transfer protein involve membrane permeabilization of target cells. Lett. Appl. Microbiol. 40, 183−189. (9) Cammue, B. P., Thevissen, K., Hendriks, M., Eggermont, K., Goderis, I. J., Proost, P., Van Damme, J., Osborn, R. W., Guerbette, F., Kader, J. C., and Broekaert, W. F. (1995) A potent antimicrobial protein from onion seeds showing sequence homology to plant lipid transfer proteins. Plant Physiol. 109, 445−455.
ASSOCIATED CONTENT
* Supporting Information S
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.6b01079. NMR spectra, secondary 13Cα and 1HN chemical shifts, statistics of spatial structure calculation, 15N relaxation rates, and results of “model-free” analysis (PDF) Accession Codes
The coordinates of the Lc-LTP2/LPPG complex have been deposited as PDB entry 5LQV. Chemical shifts of Lc-LTP2 in complex with LPPG, DMPG, and DHPC have been deposited as BMRB entries 34036, 26970, and 26971, respectively.
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REFERENCES
AUTHOR INFORMATION
Corresponding Author
*Phone: +7 495 336 44 44. E-mail:
[email protected]. ORCID
Tatiana V. Ovchinnikova: 0000-0002-5950-249X 1794
DOI: 10.1021/acs.biochem.6b01079 Biochemistry 2017, 56, 1785−1796
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DOI: 10.1021/acs.biochem.6b01079 Biochemistry 2017, 56, 1785−1796
