ARTICLE pubs.acs.org/Biomac
Fluorescence-Tagged Monolignols: Synthesis, and Application to Studying In Vitro Lignification Yuki Tobimatsu,*,† Christy L. Davidson,† John H. Grabber,‡ and John Ralph*,†,§ †
Department of Biochemistry, University of Wisconsin-Madison, Enzyme Institute, 1710 University Avenue, Madison, Wisconsin 53726, United States ‡ United States Dairy Forage Research Center, USDA-ARS, 1925 Linden Drive West, Madison, Wisconsin 53706, United States § Great Lakes Bioenergy Research Center, University of Wisconsin-Madison, Wisconsin 53706, United States
bS Supporting Information ABSTRACT: Fluorescence-tagged coniferyl alcohols, coniferyl alcohol γ-coupled by ethylenediamine spacers to dimethylaminocoumarin or nitrobenzofuran fluorophores, were tested as photoprobes to study the oxidase-mediated polymerization of monolignols. The fluorescent coniferyl alcohol derivatives readily underwent peroxidase-catalyzed in vitro copolymerization with coniferyl alcohol to yield fluorescent dehydrogenation polymers, the backbone polymers of which were structurally indistinguishable from polymers formed solely from coniferyl alcohol. To illustrate the use of the photoprobes, we successfully monitored in real time the complexation of coniferyl alcohol with horseradish apoperoxidase by F€orster resonance energy transfer (FRET) using the protein-tryptophan near the active site and a dimethylaminocoumarin moiety as donor and acceptor fluorophores. Furthermore, mixtures of fluorescence-tagged and normal coniferyl alcohols readily diffused into isolated maize cell walls and reacted with wall-bound peroxidases to form in muro artificial lignins that could be visualized by fluorescence microscopy. Thus we anticipate that fluorescence-tagged monolignols will be useful for in vitro and in vivo studies of cell wall lignification.
’ INTRODUCTION Lignins are aromatic cell wall polymers produced by the oxidative polymerization of monolignols, principally coniferyl alcohol (CA, Figure 1) and sinapyl alcohol, with typically minor amounts of p-coumaryl alcohol. The lignin polymers are most abundant in vessels, tracheids, and fibrous tissues in vascular plants where they bind, strengthen, and waterproof cell walls to provide mechanical support, enhance water transport, and help ward off plant pests. The biosynthesis and the chemical and mechanical properties of lignin have attracted significant research attention mainly because lignin is a limiting factor in numerous agro-industrial processes such as chemical pulping, forage digestibility, and the processing of lignocellulosic plant biomass into liquid biofuels.1,2 Lignification is a dynamic and complex process involving biosynthesis of monolignols inside the cell, translocation of monolignols to the apoplast, and polymerization of monolignols to form lignin. Recent studies have largely unveiled the genes, enzymes, and metabolites involved in the lignin biosynthetic pathway.3,4 Immunochemical localization of these enzymes suggested that the monolignol synthesis occurs in the cytosol.5,6 The chemical process of lignin polymerization has been wellestablished, both by structural investigations on natural and synthetic lignin polymers and by mechanistic studies of in vitro r 2011 American Chemical Society
lignin polymerization; monolignols undergo oxidative combinatorial radical coupling initiated by peroxidases or laccases located in cell walls.7,8 Genetic engineering approaches have successfully modified lignin content, composition, or both in various plant species.9,10 Despite these advances, little is known about the mechanism of monolignol translocation from the cytosol to the cell wall.11 Recent studies suggest that plant membrane proteins, including ATP-binding cassette-like (ABC) transporters, are involved in monolignol transport across the plasmalemma,1214 but other work indicates that unaided diffusion of monolignols across the plasmalemma is plausible.15 In this context, a lack of useful methods to visualize the subcellular localization and transport of monolignols is a major obstacle to fully understanding monolignol biosynthesis in plants. In addition, whereas many peroxidases and laccases capable of oxidizing monolignols have been isolated from developing xylem tissues,16,17 the specific isozymes involved in lignin polymerization are not yet clear.1,11 Both types of enzymes belong to large gene families in which the individual isozymes have significantly overlapping activities.18,19 Although Received: January 31, 2011 Revised: March 6, 2011 Published: March 16, 2011 1752
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Biomacromolecules
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Figure 1. Synthetic scheme for dimethyaminocoumarin-tagged coniferyl alcohol (CADMAC) and nitrobenzofuran-tagged coniferyl alcohol (CANBD). Conditions: (a) dihydropyran, PPTS, CH2Cl2; (b) NaBH4, EtOAc; (c) ethyl bromoacetate, KH, THF; (d) LiOH, EtOH-H2O (9:1); (e) EDCI, DMAP, DMF; (f) HCl aq; (g) EDCI, DMAP, DMF; (h) HCl aq.
several peroxidases are expressed in lignifying cells and some can generate lignin ectopically in planta upon overexpression,20,1,21 their involvement in lignin polymerization still needs to be unambiguously demonstrated. Fluorescence-based techniques have long been recognized as powerful tools in studies of biological processes. When molecules of interest are tagged with fluorescent probes, their movement and interactions in vitro or in vivo can be tracked in a highly sensitive manner using various spectroscopic techniques, such as F€orster resonance energy transfer (FRET),22,23 fluorescence anisotropy,24,25 and fluorescence correlation spectroscopy.26,22 In addition, subcellular localization of fluorescence-tagged molecules in in vitro and in vivo biological systems can be visualized by advanced microscopic imaging techniques.2729 These techniques are attractive for probing monolignolprotein interactions and possibly for visualizing monolignol transport and polymerization at the cellular level. For such applications, biologically active monolignol analogues tagged with fluorophores are required. Herein, we report synthetic protocols for preparing fluorescent dimethylaminocoumarin- (DMAC) and nitrobenzofuran- (NBD) tagged analogues of CA and demonstrate their compatibility with normal monolignols for forming synthetic lignins (dehydrogenation polymers, DHPs) via horseradish peroxidase (HRP). Finally, the
potential applications of fluorescent monolignol probes are illustrated by monitoring the complexation of CA with horseradish apoperoxidase (apoHRP) by FRET and by artificially lignifying maize cell walls for fluorescence microscopy.
’ EXPERIMENTAL SECTION General. 7-Dimethylaminocoumarin ethylenediamine 6 (trifluoroacetate salt),30,31 N-(7-nitrobenz-2-oxa-1,3,diazol-4-yl)ethylenediamine 8 (trifluoroacetate salt),32 and CA33 were synthesized according to literature methods. HRP (Type II, 200-300 U) was from Sigma-Aldrich (Milwaukee, WI), apoHRP was from Calzyme (San Luis Obispo, CA), and other chemicals were purchased from Sigma-Aldrich or Fisher Scientific (Atlanta, GA) and were used as received. Measurements. NMR spectra were acquired on a Bruker Biospin (Billerica, MA) AVANCE 500 (500 MHz) spectrometer fitted with a cryogenically cooled 5 mm TCI gradient probe with inverse geometry (proton coils closest to the sample), and spectral processing used Bruker’s Topspin 2.1 software. The central solvent peaks were used as internal reference [δH/δC: acetone, 2.04/29.8; dimethyl sulfoxide (DMSO), 2.49/39.5]. The standard Bruker implementations of 1D and 2D (gradient-selected COSY, HSQC, and HMBC) NMR experiments were used for routine structural assignments of newly synthesized compounds. Adiabatic 2D-HSQC experiments (hsqcetgpsisp2.2) for synthetic lignin 1753
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Biomacromolecules (dehydrogenation polymer, DHP) samples were carried out using the following parameters: acquired from 10 to 0 ppm in F2 (1H) with 1998 data points (acquisition time 200 ms), 200 to 0 ppm in F1 (13C) with 400 increments (F1 acquisition time 8 ms) of 64 scans with a 1.5 s interscan delay; the d24 delay was set to 0.89 ms (1/8J, J: 140 Hz). Processing used typical matched Gaussian apodization in F2 and squared cosine-bell and one level of linear prediction (32 coefficients) in F1. Ultravioletvisible (UVvis) absorption spectra were recorded on a Shimadzu BioSpec-nano spectrophotometer (Shimadzu, Kyoto, Japan) equipped with a quartz cell adapter. Fluorescence spectroscopy was conducted with a PTI QuantaMaster model C-60/2000 spectrofluorometer (Photon Technology, Lawrenceville, NJ) at 25 ( 0.1 °C and data acquisitions including spectra collections used PTI Felix32 or FelixGX software (Photon Technology). Fluorescence quantum yields (Φf) were determined according to the method described in the literature34 using anthracene (λem = 350 nm, Φf = 0.27 in EtOH, η = 1.36) or fluoresceine (λem = 450 nm, Φf = 0.92 in 0.1 N NaOH aq, η = 1.33) as standards. Gel permeation chromatography (GPC) was performed on a Shimadzu LC-20A LC system (Shimadzu, Kyoto, Japan) equipped with a photodiode array (PDA) detector (SPD-M20A; Shimadzu) using the following conditions: column: TSK gel R-M þ R-2500 (Tosoh, Tokyo, Japan); eluent: 0.1 M LiBr in dimethylformamide (DMF); flow rate: 0.5 mL min1; column oven temperature: 40 °C; sample detection: PDA response at 280 nm. The molecular weight calibration was via polystyrene standards. The data acquisition and computation used LCsolution version 1.25 software (Shimadzu). Fluorescence microscopy was performed with an Olympus BX60 epifluorescence microscope (Olympus Optical, Tokyo, Japan) equipped with an Olympus DP70 digital camera. Olympus U-MWU (excitation, 330385 nm; emission, 410430 nm) and Chroma 41020 narrow band GFP filter cubes (excitation, 480 nm; emission, 505535 nm) (Chroma Technology, Brattleboro, VT) were used for the blue and green channels, respectively.
3-[3-Methoxy-4-[(tetrahydropyran-2-yl)oxy]phenyl-2-propenal (2). Pyridinium p-toluenesulfonate (PPTS) (350 mg, 0.0014 mol)
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OMe), 3.863.95 (1H, m, THP-H5b), 4.20 (2H, t, J = 5.4 Hz, Hγ), 5.37 (1H, t, J = 7.1 Hz, THP-H1), 6.28 (1H, dt, J = 5.4, 15.9 Hz, Hβ), 6.52 (1H, d, J = 15.9 Hz, HR), 6.89 (1H, dd, J = 2.0, 8.3 Hz, H6), 7.04 (1H, d, J = 8.3, H5), 7.07 (1H, d, J = 2.0, H2). 13C NMR (acetone-d6): δ = 19.44 (THPC3), 25.99 (THP-C4), 31.03 (THP-C2), 56.15 (OMe), 62.21 (THPC5), 63.24 (Cγ), 97.98 (THP-C1), 110.89 (C2), 118.63 (C5), 119.91 (C6), 129.38 (Cβ), 129.87 (CR), 132.76 (C1), 146.85 (C4), 151.44 (C3). HR-MS (ESI) calcd for C15H20NaO4 [(M þ Na)þ]: 287.1254; found: 287.1241.
Ethyl 3-{3-Methoxy-4-[(tetrahydropyran-2-yl)oxy]phenyl2-propen-1-yl}oxy Acetate (4). Compound 3 (2.6 g, 0.01 mol) in anhydrous tetrahydrofuran (10 mL) was added to a suspension of potassium hydride (50% in mineral oil, 1.2 g, 0.015 mol) in anhydrous tetrahydrofuran (10 mL), followed by the addition of ethyl bromoacetate (1.7 mL, 0.015 mol) at 0 °C. After 30 min of stirring, the reaction mixture was warmed to 40 °C and kept under stirring at that temperature for 15 h. The reaction was quenched by the addition of aqueous ammonium chloride solution (50 mL) at 0 °C and then extracted with ethyl acetate (100 mL). The organic layer was washed with brine, dried over sodium sulfate, and evaporated under reduced pressure to give yellowish oil, which was purified by silica-gel chromatography to give compound 4 as a colorless oil, 2.3 g, 65% yield. 1H NMR (acetone-d6): δ = 1.22 (3H, t, J = 7.2, CH3CH2), 1.511.67, 1.781.85, 1.912.02 (6H, m, THP-H2, -H3, and -H4), 3.493.56 (1H, m, THP H5a), 3.84 (3H, s, OMe), 3.90 (1H, t, J = 10.5, THP H5b), 4.10 (2H, s, H10 ), 4.14 (2H, q, J = 7.2, CH3CH2-), 4.19 (2H, d, J = 6.2, Hγ), 5.38 (1H, br s, THP H1), 6.24 (1H, dt, J = 6.3, 15.9, Hβ), 6.57 (1H, d, J = 15.9, HR), 6.92 (1H, d, J = 8.2, H6), 7.05 (1H, d, J = 8.2, H5), 7.12 (1H, s, H2). 13C NMR (acetone-d6): δ = 14.45 (CH3CH2), 19.39 (THP-C3), 25.94 (THP-C4), 30.98 (THP-C2), 56.12 (OMe), 60.84 (CH3CH2-), 62.19 (THP-C5), 67.53 (C10 ), 72.25 (Cγ), 97.84 (THP-C1), 110.87 (C2), 118.40 (C5), 120.25 (C6), 124.78 (Cβ), 132.06 (C1), 133.17 (CR), 147.16 (C4), 151.37 (C3) 170.84 (C20 ). HR-MS (ESI) calcd for C19H26NaO6 [(M þ Na)þ]: 373.1622; found: 373.1631.
3-{3-Methoxy-4-[(tetrahydropyran-2-yl)oxy]phenyl-2propen-1-yl}oxy Acetic Acid (5). To a solution of compound 4
was added to a solution of coniferaldehyde (4-hydroxy-3-methoxycinnamaldehyde) 1 (5.0 g, 0.028 mol) and 3,4-dihydro-2H-pyran (5.1 mL, 0.056 mol) in anhydrous dichloromethane (40 mL) (Figure 1). After being stirred at room temperature for 3 h, distilled water was added, and the product was extracted with ethyl acetate (200 mL). The organic layer was washed with brine (3 50 mL), dried over sodium sulfate, and evaporated under reduced pressure to give a solid residue, which was recrystallized from ethanol to afford compound 2 as a yellowish solid, 6.0 g, 82% yield. 1H NMR (acetone-d6): δ = 1.541.70, 1.821.86, 1.942.01 (6H, m, THP-H2, -H3, and -H4), 3.543.58 (1H, m, THPH5), 3.823.87 (1H, m, THP-H50 ), 3.90 (3H, s, OMe), 5.52 (1H, br s, THP-H1), 6.70 (1H, dd, J = 15.9, 7.8 Hz, Hβ), 7.18 (1H, d, J = 8.3 Hz, H5), 7.23 (1H, d, J = 8.3 Hz, H6), 7.39 (1H, s, H2), 7.59 (1H, d, J = 15.9 Hz, HR), 9.65 (1H, d, J = 7.8 Hz, Hγ). 13C NMR (acetone-d6): δ = 19.31 (THP-C3), 25.85 (THP-C4), 30.87 (THP-C2), 56.31 (OMe), 62.35 (THP-C5), 97.54 (THP-C1), 112.11 (C2), 117.54 (C5), 123.81 (C6), 127.88 (Cβ), 129.32 (C1), 150.31 (C3), 151.39 (C4), 153.59 (CR), 193.90 (Cγ). HR-MS (ESI) calcd for C15H18NaO4 [(M þ Na)þ]: 285.1098; found: 285.1096.
(1.9 g, 0.0054 mol) in EtOHwater (9:1 v/v, 20 ml), lithium hydroxide monohydrate (1.1 g, 0.027 mol) was added at 0 °C. After 1 h of stirring, the reaction mixture was carefully neutralized with acetic acid and extracted with i-PrOHCH2Cl2 (2:1, v/v, 100 mL) over brine. The organic layer was dried over sodium sulfate and evaporated under reduced pressure to give compound 5 as a pale yellow oil, which was purified by silica-gel chromatography to a colorless oil, 1.6 g, 89% yield. 1H NMR (acetone-d6/D2O, 9:1, v/v): δ = 1.561.73, 1.841.98, 1.992.10 (6H, m, THP-H2, -H3, and -H4), 3.563.63 (1H, m, THP-H5a), 3.87 (3H, s, OMe), 3.97 (1H, t, J = 10.3, THP-H5b), 4.14 (2H, s, H10 ), 4.14 (2H, q, J = 7.1, H30 ), 4.26 (2H, d, J = 5.2, Hγ), 5.41 (1H, br s, THP-H1), 6.15 (1H, dt, J = 5.2, 15.8, Hβ), 6.56 (1H, d, J = 15.8, HR), 6.90 (1H, d, J = 8.2, H6), 6.95 (1H, s, H2), 7.07 (1H, d, J = 8.2, H5). 13C NMR (acetone-d6/D2O, 9:1, v/v): δ = 18.76 (THP-C3), 25.15 (THP C4), 30.22 (THP C2), 55.99 (OMe), 62.16 (THP-C5), 66.57 (C10 ), 72.19 (Cγ), 97.33 (THP-C1), 109.86 (C2), 117.34 (C5), 119.88 (C6), 124.45 (Cβ), 130.56 (C1), 134.29 (CR), 146.32 (C4), 150.13 (C3) 173.88 (C20 ). HR-MS (ESI) calcd for C17H22O6 [(M H)]: 321.1333; found: 321.1323.
3-[3-Methoxy-4-[(tetrahydropyran-2-yl)oxy]phenyl-2propenol (3). To a solution of compound 2 (5.6 g, 0.021 mol) in ethyl
Dimethyaminocoumarin-Tagged Coniferyl Alcohol (CADMAC). To a solution of compound 5 (540 mg, 1.7 mmol),
acetate (100 mL), sodium borohydride (1.6 g, 0.042 mol) was added and stirred at room temperature for 16 h. The reaction was quenched with saturated aqueous ammonium chloride solution and extracted with ethyl acetate. The organic layer was dried over sodium sulfate and evaporated under reduced pressure to give a residue, which was purified by silica-gel chromatography to afford compound 3 as a colorless oil, 4.8 g, 85% yield. 1 H NMR (acetone-d6): δ = 1.521.67, 1.791.84, 1.942.00 (6H, m, THP-H2, -H3, and -H4), 3.503.55 (1H, m, THP-H5a), 3.84 (3H, s,
7-dimethylaminocoumarin ethylenediamine trifluoroacetate 6 (810 mg, 2.0 mmol), and N-ethyl-N0 -(3-dimethylaminopropyl)carbodiimide hydrochloride (EDCI) (390 mg, 2.0 mmol) in anhydrous N,N-dimethylformamide (DMF) (10 mL) was added 4-dimethylaminopyridine (DMAP) (100 mg, 0.9 mmol) at room temperature. After being stirred for 12 h, 0.1 N aq HCl was added at 0 °C, and the reaction mixture was stirred for an additional 10 min. The reaction mixture was extracted with chloroform (200 mL), washed with saturated sodium bicarbonate 1754
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Biomacromolecules aqueous solution (100 mL), dried over sodium sulfate, and evaporated under reduced pressure. Purification by silica-gel chromatography yielded CADMAC as a yellowish solid, 720 mg, 84% yield. 1 H NMR (acetone-d6): δ = 3.05 (6H, s, NMe), 3.323.35 (4H, m, H120 and H130 ), 3.64 (2H, s, H90 ), 3.82 (2H, s, H160 ), 3.84 (3H, s, OMe), 4.09 (2H, d, J = 6.2, Hγ), 6.02 (1H, s, H30 ), 6.16 (1H, dt, J = 6.2, 15.9, Hβ), 6.47 (1H, d, J = 2.6, H80 ), 6.54 (1H, d, J = 15.9, HR), 6.68 (1H, dd, J = 2.6, 9.0, H60 ), 6.76 (1H, d, J = 8.1, H5), 6.88 (1H, dd, J = 1.9, 8.1, H6), 7.09 (1H, d, J = 1.9, H2), 7.56 (1H, d, J = 9.0, H50 ). 13 C NMR (acetone-d6): δ = 39.34 (C130 ), 40.13 (NMe), 40.29 (C120 ), 40.35 (C9), 56.15 (OMe), 69.89 (C160 ), 72.66 (Cγ), 98.50 (C80 ), 109.44 (C60 ), 109.66 (C2), 110.11 (C4a0 ), 111.06 (C30 ), 115.75 (C5), 121.01 (C6), 123.22 (Cβ), 126.82 (C5 0 ), 129.60 (C1), 133.82 (CR), 147.56 (C4), 148.43 (C3), 151.19 (C4 0 ), 153.95 (C7 0 ), 156.86 (C1a 0 ), 161.49 (C2 0 ), 169.18 (C100 ), 170.60 (C15 0 ). HRMS (ESI) calcd for C27H 31 N3NaO 7 [(M þ Na)þ]: 532.2055; found: 532.2049. Nitrobenzofuran-Tagged Coniferyl Alcohol (CANBD). To a solution of compound 5 (540 mg, 1.7 mmol), N-(7-nitrobenz-2-oxa-1,3, diazol-4-yl)ethylenediamine trifluoroacetate 8 (680 mg, 2.0 mmol) and EDCI (390 mg, 2.0 mmol) in anhydrous DMF (10 mL) was added DMAP (100 mg, 0.9 mmol) at room temperature. After being stirred for 12 h, 0.1 N HCl aq was added at 0 °C, and the reaction mixture was stirred for an additional 10 min. The reaction mixture was extracted with ethyl acetate (200 mL), washed with saturated aqueous sodium bicarbonate solution (100 mL), dried over sodium sulfate, and evaporated under reduced pressure. Purification by silica-gel chromatography yielded CANBD as an orange powder, 620 mg, 82% yield. 1H NMR (acetone-d6): δ = 3.713.79 (2H, m, H120 ) 3.793.84 (2H, m, H110 ), 3.89 (3H, s, OMe), 3.99 (2H, s, H150 ), 4.19 (2H, d, J = 6.4, Hγ), 6.19 (1H, dt, J = 6.4, 15.9, Hβ), 6.53 (1H, d, J = 8.8, H50 ), 6.56 (1H, d, J = 15.9, HR), 6.79 (1H, d, J = 8.1, H5), 6.89 (1H, dd, J = 1.8, 8.1, H6), 7.08 (1H, d, J = 1.8, H2), 8.57 (1H, d, J = 8.8, H60 ). 13C NMR (acetone-d6): δ = 38.10 (C110 ), 45.10 (C120 ), 56.12 (OMe), 69.88 (C150 ), 72.78 (Cγ), 99.53 (C50 ), 110.03 (C2), 115.76 (C5), 120.93 (C6), 122.96 (Cβ), 123.55 (C70 ), 129.39 (C10 ), 134.19 (CR), 137.83 (C60 ), 145.05 (C80 ), 145.47 (C30 ), 145.80 (C40 ), 147.62 (C4), 148.40 (C3), 171.73 (C140 ). HR-MS (ESI) calcd for C20H22N5O7 [(M þ H)þ]: 444.1514; found: 444.1493. HRP-Catalyzed Dehydrogenative Polymerization. Fluorescence-tagged CA (CANBD or CADMAC, 0.075 mmol) and CA (0.425 mmol) in 240 mL of acetone/sodium phosphate buffer (0.1 M, pH 6.5) (1:9, v/v) and a separate solution of hydrogen peroxide (0.6 mmol) in 240 mL of water were added by peristaltic pump over a 20 h period at 25 °C to 60 mL of buffer containing HRP (2.5 mg). The reaction mixture was further stirred for 4 h; then, the precipitate was collected by centrifugation (10 000g, 15 min), washed with ultrapure water (100 mL 3), and lyophilized to afford DHPs. FRET-Based Binding Study. The complexation of CADMAC and apoHRP was monitored by FRET using a PTI QuantaMaster model C-60/2000 spectrofluorometer (Photon Technology). The fluorescence emission spectra were recorded between 305 and 600 nm using 4 nm excitation and emission slits. We set the excitation wavelength to 295 nm to excite selectively tryptophan residues in the protein.35,22 The concentration of apoHRP in sodium phosphate buffer (0.1 M, pH 7.4) was determined by absorption spectrometry using a molar extinction coefficient of 20 000 M1 cm1 at 280 nm.36 The solution (2500 μL) of 10 μM apoHRP in sodium phosphate buffer (0.1 M, pH 7.4) was placed in a 1 cm quartz cuvette and set in the spectrometer at 25 ( 0.1 °C. The reaction was initiated by the addition of 150 μL of buffer solution containing 50 μM CADMAC, 0100 μM CA, and 10 μM apoHRP to the cuvette, and emission spectra were periodically recorded until no change in fluorescence intensity was observed. Preparation of Artificially Lignified Maize Cell Walls. Primary nonlignified cell walls were isolated from maize cell suspensions
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Table 1. Optical Properties of Fluorescence-Tagged (And Untagged) Coniferyl Alcoholsa compounds
λab (nm)
ε (M1 cm1)
λem (nm)
Δλ (nm)
Φf
b
b
89 72
0.44c 0.17d
CA
266
15 100
b
CADMAC CANBD
372 460
18 900 19 800
461 532
EtOH as solvent, λab: absorption maxima, ε: extinction coefficient, λem: emission maxima, Δλ: Stoke shift, Φf: quantum yield. b No fluorescence. c Anthracene as a standard (excitation at 350 nm). d Fluorescein as a standard (excitation at 450 nm). a
(Zea mays L. cv. Black Mexican), as previously described.37 Freshly prepared, fully hydrated cell walls (40 g, ∼1.0 g dry weight) were stirred in 100 mL of water with 4 mM CaCl2 (pH 5.2) and artificially lignified via in situ peroxidases adding separate solutions of monolignols and hydrogen peroxide via a peristaltic pump over a 20 h period at 25 °C. Monolignols (100 mg CA and 5 mg CADMAC or CANBD) were prepared in 50 mL of 10% (v/v) dioxane/water, and hydrogen peroxide (30%, 70 μL, ∼1.1 equiv) was prepared in 50 mL of water. Following additions, cell walls were stirred for an additional 4 h and then collected by centrifugation (3700g, 15 min). The cell walls were then washed twice by resuspending the pellet in 180 mL of water, followed by centrifugation (3700g, 15 min). Cell walls were then resuspended in water (1:1 v/v), and an aliquot was removed for fluorescence microscopy. The remaining cell walls were then further washed with 9:1 (v/v) acetone/water, oven-dried at 55 °C, and subjected to a gel-state NMR38 and Klason lignin39 analyses.
’ RESULTS AND DISCUSSION Chemistry. When designing fluorescent monolignols, we anticipated that γ-attachment of fluorophores would have minimal effects on the enzyme-mediated oxidation and coupling reactions of CA. As evidence, our group established that acylated (acetylated, p-hydroxybenzylated, and p-coumaroylated) lignins in some plant species are derived from γ-acylated monolignols.40,41 Moreover, our in vitro lignification studies demonstrated that γ-acylation42 and γ-glucosylation43,44 of monolignols do not particularly perturb key radical coupling reactions; some postcoupling rearomatization reactions are, however, altered during lignification, producing novel tetrahydrofuran products in the lignin polymer.40,42 For selection of fluorophores, special prudence was paid to select molecular structures that would remain intact during lignification because peroxidase isozymes can form radicals and modify a wide variety of phenolic and aniline derivatives.19,45 Following these considerations, two commonly used fluorophores, DMAC and NBD, were chosen for this study. Finally, the introduction of an ethylenediamine spacer was thought to be desirable to improve accessibility and flexibility of the CA moiety after conjugation with the rigid and bulky fluorophores and to increase the water solubility of the final conjugates. The synthesis started by protecting the phenolic hydroxyl group of commercially available coniferaldehyde 1 as a tetrahydropyranyl (THP) ether 2 in 82% yield. Hydride reduction of the cinnamaldehyde group gave THP-protected CA 3 in 85% yield. The free γ-hydroxyl group was then deprotonated with potassium hydride and alkylated with ethyl bromoacetate to give ester 4 in 65% yield, which was hydrolyzed with aq LiOH to give carboxymethylated CA 5 in 80% yield. DMAC and NBD fluorophores bearing ethylenediamine linkers, compounds 630,31 and 8,32 were synthesized according to literature methods. For the final assembly of the target fluorescence-tagged CAs, 1755
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Table 2. Horseradish Peroxidase-Catalyzed Polymerization of Fluorescence-Tagged (and Untagged) Coniferyl Alcohols optical propertiesc DHP
monomera
yield (weight%)
Mnb
Mwb
PDb
λab (nm)
λem (nm)
FI
CADMAC-DHP
CA & CADMAC
89
2300
6860
2.98
362
450
1
CANBD-DHP
CA & CANBD
88
2220
7900
3.56
460
531
0.514
CA-DHP
CA only
86
2330
8130
3.49
333
380
0.008
a
CADMAC and CANBD were copolymerized with CA at 15 mol % feed. b Determined by GPC, polystyrene as standards, Mn: number-average molecular weight, Mw: weight-average molecular weight, PD: polydispersity (Mw/Mn). c Dioxanewater (95:5, v/v) as solvent, λab: absorption maxima (>300 nm), λem: emission maxima, FI: relative fluorescence intensity based on emission peak integral under same conditions (50 μg/mL in dioxaneH2O 95:5, excited at the corresponding absorption maxima).
EDCI-promoted amide couplings of compound 5 with fluorophores 6 and 8 were carried out in the presence of DMAP at room temperature. After confirming complete conversion of compound 5 by thin layer chromatography, THP groups were removed by acidification with aq HCl at 0 °C, yielding the desired CADMAC and CANBD fluorescent compounds in 89 and 83% yields, respectively. Optical properties of CADMAC and CANBD were verified by UVvis absorption and fluorescence spectroscopy (Table 1, Figure S1 in the Supporting Information). As expected, CADMAC exhibited a strong bright-blue fluorescence under UV-A light, and CANBD exhibited a green fluorescence under blue light, whereas untagged CA did not show any visible fluorescence. HRP-Catalyzed Dehydrogenative Polymerization. It is important to determine whether fluorescence-tagged CAs are compatible with H-abstraction by peroxidase and the subsequent radical coupling reactions of lignification. For this purpose, we produced synthetic lignins (DHPs) using a conventional HRP/H2O2 system, in which CADMAC and CANBD were used as low-level monomer substitutes (15 mol %) for CA. Product yields, average molecular weights, and dispersity data for DHPs formed with CADMAC and CANBD were quite similar to DHPs formed with CA (Table 2). The degree of polymerization for the DHPs averaged 13 (based on the molecular weight of CA), which is comparable to literature values for conventional DHPs.46 DHP structural compositions were determined by 2D HSQC-NMR. Major lignin interunit linkages in DHPs formed with CADMAC and CANBD were typical βO4-, β5-, and ββ-substructures,38 and their distributions were similar to those in the DHP formed only with CA (Figure S2, Supporting Information). The inclusion of fluorescence-tagged CAs was clearly demonstrated by the presence of fluorophore signals in the aromatic region (Figure 2). These data indicate that fluorescent CAs were incorporated into lignin polymers with an intact fluorophore moiety. As previously demonstrated for γ-acylated and γglucosylated monolignols,40,4244 the polymerization of CADMAC and CANBD logically produces several tetrahydrofuran-type ββlinked subunits because of their lack of a free γOH for internal trapping of the bis-quinone methide intermediates formed via ββcoupling. NMR evidence of these structures in the present DHPs is not yet conclusive because of low levels (if any) of such coupling products. Optical properties of the DHPs were verified by UVvis and fluorescence spectroscopy. The optical spectra of DHPs displayed characteristic absorption and emission signals from the fluorophores, further supporting successful incorporation of fluorescent CAs into the lignin polymers (Figure 3, Table 2). The CA-DHP was weakly fluorogenic, as reported.47,48 Lignin autofluorescence was, however, negligibly small relative to DHPs with fluorescence-tagged CAs, and this was easily verified visually and by spectroscopic methods (Figure 3B, Table 2).
Monolignol-Apoperoxidase Interactions Probed by FRET. In principle, fluorescence-tagged CAs can be used in a variety of fluorescence assays to study the molecular interactions of monolignols. To examine this possibility, we explored the feasibility of a FRET-based protocol for monitoring the interactions between monolignols and HRP, a model peroxidase widely used for in vitro lignin polymerization studies. HRP has a single tryptophan residue (Trp117) that is predicted to be located within 1218 Å of the active site for peroxidation,49,50 and this can serve as an intrinsic fluorescence probe for FRET studies. Because FRET efficiency depends on the inverse sixth power of the distance between donor and acceptor fluorophores,51 monitoring fluorescence signals from the protein-tryptophan donor and the fluorescent CA acceptor can, in principle, reveal the process of HRP-monolignol complexation. In the present study, apoHRP (a heme-free HRP) was chosen instead of native HRP because the absence of hematoporphyrin facilitates the observation of the tryptophan fluorescence.52 Upon excitation at 295 nm, the fluorescence spectrum of apoHRP exhibited a characteristic emission at 329 nm for the excited state of tryptophan. The DMAC fluorophore was confirmed as an appropriate energy acceptor because its absorption spectrum overlapped with the emission spectrum of tryptophan in apoHRP (Figure 4A). After exposure of CADMAC in the apoHRP buffer solution (apoHRP/ CADMAC ratio ≈ 3.3), the complex formation was evident; the tryptophan excitation gradually decreased along with rising CADMAC emission (Figure 4B), indicating that the energy transferred from excited tryptophan sensitized the proximate DMAC probe. Moreover, the DMAC emission was blue-shifted as the complexation proceeded. Because the emission maximum of CADMAC is shifted from 482 nm in phosphate buffer to 460 nm in EtOH (Figure 4A), the blue shift observed during the reaction suggests that the probe experiences a less polar environment in the active site. It has been proposed that the binding site of peroxidase is located in a hydrophobic protein interior.53,54 Therefore, the blue-shifted DMAC emission supports the contention that CADMAC is captured in the hydrophobic binding pocket of apoHRP (Figure 5). Accordingly, the time course of CADMAC-apoHRP complexation in the absence and presence of CA was successfully traced by monitoring the fluorescence intensity ratio at 329 and 460 nm (Figure 6). Nonlabeled CA effectively suppressed FRET generation, clearly indicating that CA competes with CADMAC for the apoHRP binding site. We also monitored the reaction of apoHRP with an aminocoumarin derivative without a monolignol moiety (tert-butoxycarbonylamino derivative from compound 6). In this case, neither FRET signals nor blue-shifted DMAC emission was significant (Figure S3 in the Supporting Information), indicating negligible nonspecific 1756
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Figure 2. Aromatic regions of 2D 13C1H correlation (HSQC) NMR spectra of synthetic lignins (DHPs) produced from (A) 85% coniferyl alcohol with 15% dimethylaminocoumarin-tagged coniferyl alcohol (CADMAC-DHP), (B) 85% coniferyl alcohol with 15% nitrobenzofuran-tagged coniferyl alcohol (CANBD-DHP), and (C) 100% coniferyl alcohol (CADHP).
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Figure 3. (A) Normalized UVvis absorption spectra of synthetic lignins (DHPs). (B) Fluorescence emission spectra of DHPs under identical conditions (50 μg/mL in dioxane/water 95/5 v/v excitation at the corresponding absorption maxima listed in Table 2). Insert is a picture of DHP solutions under irradiation by UV light (365 nm). CADMACDHP: DHP prepared from 85% coniferyl alcohol with 15% dimethylaminocoumarin-tagged coniferyl alcohol; CANBD-DHP: DHP prepared from 85% coniferyl alcohol with 15% nitrobenzofuran-tagged coniferyl alcohol; CA-DHP: DHP prepared from 100% coniferyl alcohol.
absorption of the probe onto the protein surface. These results confirm that the affinity of the monolignol moiety for the proteinbinding site is the driving force for CADMAC-apoHRP complexation observed in this work. The differing specificities of peroxidases are related to the accessibility of protein-binding sites by substrates,55,56 and this has mainly been examined by computer modeling.53,54 This, however, must be confirmed experimentally; our data demonstrate the utility of the fluorescence-tagged monolignols for monitoring monolignol-peroxidase interactions. Artificially Lignified Maize Cell Walls (CW-DHPs). Finally, to test the use of fluorescence-tagged CAs for microscopy, we polymerized CA with a small amount (5% by weight) of fluorescence-tagged CAs into nonlignified maize primary walls via wall-bound peroxidases and exogenously supplied H2O2. Previous studies have demonstrated that the structure and distribution of artificial lignins formed by this model system closely mimic natural lignification in grasses.57 The successful formation of artificial lignins was confirmed by gel-state 2D-HSQC NMR of whole cell walls (Figure S4 in the Supporting Information). Klason lignin contents of CW-DHPs from
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Figure 4. (A) Absorption and fluorescence emission spectra of apoperoxidase from horseradish (apoHRP) and dimethyaminocoumarin-tagged coniferyl alcohol (CADMAC). (B) Fluorescence emission spectra following the time-course of the complexation of apoHRP and CADMAC.
Figure 5. Schematic presentation of the complexation of dimethyaminocoumarin-tagged coniferyl alcohol (CADMAC) with apoperoxidase from horseradish (apoHRP).
CADMAC and CANBD and without fluorescent monomers (only CA) were similar, averaging 12.0, 12.6, and 11.8%, respectively. As expected, CADMAC and CANBD labeled CW-DHPs and produced a blue and green fluorescence (Figure 7A,B), whereas autofluorescence from CW-DHPs prepared without fluorescent CAs or with DMAC and NBD derivatives without monolignol moieties was not significantly visible under identical microscopic conditions (Figure 7C1,C2 and Figure S5 in the Supporting Information). The fluorescence from CADMAC and CANBD was observed throughout the cell wall, 1758
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fluorescent monolignols, such as p-coumaryl and sinapyl alcohol derivatives. Their application in fluorescence-based molecular interaction studies was demonstrated by using FRET for sensing the monolignol-apoperoxidase binding process. In addition, biomimetic in vitro lignification studies revealed the compatibility of fluorescence-tagged CAs with lignin polymerization; they act as substrates for peroxidase and are integrally crosscoupled to lignin polymers, generating bright fluorescence easily seen in the presence of the autofluorescence from lignin and other cell wall components. Therefore, these fluorescence-tagged CAs might be useful for visualizing the transport and deposition of monolignols in biological systems. Overall, we anticipate that fluorescence-tagged monolignols can be used in a variety of in vitro and in vivo studies aimed at understanding lignification processes in plants.
’ ASSOCIATED CONTENT
bS Figure 6. Time course of apoperoxidase and dimethyaminocoumarintagged coniferyl alcohol (CADMAC) emission ratio (I460/I329) during their complexation in the absence and presence of coniferyl alcohol (CA).
Supporting Information. UVvis and fluorescence spectra of CADMAC, CANBD, and CA; aliphatic regions from 2D HSQC NMR spectra of DHPs; FRET-based binding assay of the reaction of a DMAC derivative with apoHRP; gel-state 2D HSQC NMR spectra of CW-DHPs; and the full set of microscopic images of CW-DHPs. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected];
[email protected].
Figure 7. Epifluorescence microscopic images of artificially lignified maize cell walls (CW-DHPs). (A) CW-DHP lignified with dimethylaminocoumarin-tagged coniferyl alcohol observed via the blue channel (excitation, 330385 nm; emission, 410430 nm). (B) CW-DHP lignified with nitrobenzofuran-tagged coniferyl alcohol observed via the green channel (excitation, 480 nm; emission, 505535 nm). (C) CW-DHP lignified only with untagged coniferyl alcohol observed via the blue (C1) and green (C2) channels.
which apparently has a single thin (
